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Am J Physiol Renal Physiol 290: F465-F477, 2006. First published September 13, 2005; doi:10.1152/ajprenal.00305.2005
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Accumulation of nonesterified fatty acids causes the sustained energetic deficit in kidney proximal tubules after hypoxia-reoxygenation

Thorsten Feldkamp,1,2 Andreas Kribben,2 Nancy F. Roeser,1 Ruth A. Senter,1 and Joel M. Weinberg1

1Division of Nephrology, Department of Internal Medicine, Veterans Affairs Ann Arbor Healthcare System and University of Michigan, Ann Arbor, Michigan; and 2Division of Nephrology and Hypertension, Department of Internal Medicine, University Hospital Essen, Essen, Germany

Submitted 29 July 2005 ; accepted in final form 31 August 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Kidney proximal tubules exhibit decreased ATP and reduced, but not absent, mitochondrial membrane potential ({Delta}{psi}m) during reoxygenation after severe hypoxia. This energetic deficit, which plays a pivotal role in overall cellular recovery, cannot be explained by loss of mitochondrial membrane integrity, decreased electron transport, or compromised F1F0-ATPase and adenine nucleotide translocase activities. Addition of oleate to permeabilized tubules produced concentration-dependent decreases of {Delta}{psi}m measured by safranin O uptake (threshold for oleate = 0.25 µM, 1.6 nmol/mg protein; maximal effect = 4 µM, 26 nmol/mg) that were reversed by delipidated BSA (dBSA). Cell nonesterified fatty acid (NEFA) levels increased from <1 to 17.4 nmol/mg protein during 60- min hypoxia and remained elevated at 7.6 nmol/mg after 60 min reoxygenation, at which time ATP had recovered to only 10% of control values. Safranin O uptake in reoxygenated tubules, which was decreased 85% after 60-min hypoxia, was normalized by dBSA, which improved ATP synthesis as well. dBSA also almost completely normalized {Delta}{psi}m when the duration of hypoxia was increased to 120 min. In intact tubules, the protective substrate combination of {alpha}-ketoglutarate + malate ({alpha}-KG/MAL) increased ATP three- to fourfold, limited NEFA accumulation during hypoxia by 50%, and lowered NEFA during reoxygenation. Notably, dBSA also improved ATP recovery when added to intact tubules during reoxygenation and was additive to the effect of {alpha}-KG/MAL. We conclude that NEFA overload is the primary cause of energetic failure of reoxygenated proximal tubules and lowering NEFA substantially contributes to the benefit from supplementation with {alpha}-KG/MAL.

acute renal failure; kidney; mitochondrial membrane potential

THE RENEWED APPRECIATION OF the central role played by mitochondria in cell death and the multiplicity of ways in which it is regulated has led to reassessment and clarification of a number of longstanding observations that, although well established, had remained phenomenological and poorly understood (21, 22, 41, 66, 98). During ischemia, the combination of inhibition of mitochondrial and peroxisomal {beta}-oxidation and continued phospholipid hydrolysis mediated by normal and increased phospholipase activity (47, 54, 56, 57, 60, 61) and unopposed by ATP-requiring reesterification (50, 91) leads to intracellular accumulation of nonesterified fatty acids (NEFA; see Refs. 26, 42, 82, 86, 95, 97) and their coenzyme A (CoA; see Refs. 32 and 75) and carnitine esters (32, 55, 56, 67, 75). These metabolites, in turn, have been implicated in progression of cell injury, with multiple effects on mitochondrial function including "detergent" actions on membrane structure (32), uncoupling (3, 30, 33, 72, 90, 92, 93), inhibition of adenine nucleotide transport (51, 71, 75, 90, 94), promotion of the mitochondrial permeability transition (5, 27, 44, 87), and activation of apoptotic pathways (68). However, the necessary roles of these various processes relative to each other and to the multiplicity of other simultaneous events in the complex milieu of the ischemic cell during the progression of mitochondrial and cell injury has remained incompletely defined in most cases.

In kidney proximal tubule cells, which are major sites of ischemic cell damage during acute renal failure (40), NEFA accumulate progressively during ischemia in vivo (42, 97) and hypoxia in vitro (26, 60, 82, 86, 95) via both calcium-dependent and -independent mechanisms (26, 60, 61, 82, 86, 95). Zager and coworkers (96) have demonstrated that treatment of isolated tubules with exogenous phospholipase A2 or with arachidonate during hypoxia/reoxygenation (H/R) further impairs recovery of ATP during reoxygenation despite at the same time decreasing plasma membrane damage. Subsequent studies documented persistent mitochondrial dysfunction in isolated tubules subjected to H/R (81, 83, 84) that plays a pivotal role in overall cellular recovery (85). The energetic deficit is characterized by intact function of the electron transport chain (17), absence of cytochrome c release (84), preserved activity of the mitochondrial F1FO-ATPase and adenine nucleotide translocase (18), and partial but incomplete recovery of mitochondrial membrane potential ({Delta}{Psi}m; see Refs. 18, 19, 83, 84). Mitochondrial function can be improved by supplementing tubules with specific tricarboxylic acid cycle metabolites such as {alpha}-ketoglutarate and malate individually and in combination during hypoxia or reoxygenation (17–19, 83, 84).

To better characterize the mitochondrial defect, we have performed studies on proximal tubules exposed to a concentration of digitonin that selectively permeabilized the plasma membrane and allowed access for exogenously provided substrates, nucleotides, and probes to structurally intact mitochondria without perturbation by the glycoside (18, 19). This permitted the use of safranin O for more quantitative and dynamic measurements of {Delta}{Psi}m. These studies showed that, despite the maintenance of electron transport and adenine nucleotide translocase and F1FO-ATPase activity, neither optimal substrate delivery to mitochondria nor availability of excess ATP fully restores {Delta}{Psi}m in tubules permeabilized at the end of H/R, suggesting the contribution of an inner mitochondrial membrane leak to the persistent deenergization1 (18). In the present work, we demonstrate that the mitochondrial deenergization is completely reversible and caused by NEFA. Moreover, the benefit provided by protective tricarboxylic acid cycle metabolites can be in large part explained by their effects to lower NEFA levels. These observations define NEFA accumulation as the primary cause of mitochondrial dysfunction and the energetic deficit that develop in proximal tubules during H/R and ischemia-reperfusion.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
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Materials. Female New Zealand White rabbits (1.5–2.0 kg) were obtained from Harlan (Indianapolis, IN). Type I collagenase was from Worthington Biochemical (Lakewood, NJ). Percoll was purchased from Amersham Biosciences (Piscataway, NJ). HPLC-grade acetonitrile was from Fisher Scientific (Pittsburgh, PA). 5,5',6,6'-Tetrachloro-1,1',3,3'-tetraethylbenzimidazocarbocyanine iodide (JC-1) was supplied by Molecular Probes (Eugene, OR). High-purity digitonin was purchased from Calbiochem (catalog no. 300411; San Diego, CA). NEFA-C assay kits were from WAKO Diagnostics (Richmond, VA). Delipidated BSA (dBSA) was from Sigma-Aldrich (catalog no. A6003; St. Louis, MO). All other reagents and chemicals were of the highest grade available from Sigma-Aldrich. Aqueous stock solutions of experimental reagents were all pH adjusted so as not to alter the final pH values of the experimental medium. Regents that required solubilization in ethanol or DMSO were delivered from ≥1,000x stock solutions.

Isolation of tubules. Proximal tubules were prepared from kidney cortex of female New Zealand White rabbits by collagenase digestion and centrifugation on self-forming Percoll gradients as described (17, 80, 83–85).

Experimental procedure. Incubation conditions were similar to our previous studies (17–19, 81, 83–85). Tubules were suspended at 3.0–5.0 mg tubule protein/ml in a 95% air-5% CO2-gassed medium containing (in mM) 110 NaCl, 2.6 KCl, 25 NaHCO3, 2.4 KH2PO4, 1.25 CaCl2, 1.2 MgCl2, 1.2 MgSO4, 5 glucose, 4 sodium lactate, 0.3 alanine, 5 sodium butyrate, 2 glycine, and 1.0 mg/ml bovine gelatin (75 bloom; solution A). After 15–30 min preincubation at 37°C, tubules were resuspended in fresh solution A with experimental agents and regassed with either 95% air-5% CO2 (normoxic controls) or 95% N2-5% CO2 (hypoxia). During hypoxia, solution A was kept at pH 6.9 to simulate tissue acidosis during ischemia in vivo (81) and omitted glucose, lactate, alanine, and butyrate. These incubation conditions result in near-anoxic conditions. However, it is not possible to confirm the presence of complete anoxia in the flasks, so we use the term hypoxia to describe the oxygen deprivation. After 60 or 120 min, samples were removed for analysis. The remaining tubules were pelleted and then resuspended in fresh 95% air-5% CO2-gassed, pH 7.4 solution A with experimental agents as needed. In all studies except those for measurement of NEFA, sodium butyrate in solution A was replaced with 2 mM heptanoic acid. To assure availability of purine precursors for ATP resynthesis, 250 µM AMP was added at the start of reoxygenation (81, 84). After either 60 or 120 min of reoxygenation, samples were removed again for analysis. {alpha}-Ketoglutarate + malate ({alpha}-KG/MAL, 4 mM each) was added from concentrated stock solutions of neutralized sodium salts of the two substrates during either hypoxia, reoxygenation, or both periods to allow assessment of the behavior of substrate-protected tubules (17, 19, 83, 84).

Measurement of ATP levels. Samples of tubule cell suspension were immediately deproteinized in TCA, neutralized with trioctylamine-CFC 113, and stored at –20°C as previously described (81). Purine nucleotides and their metabolites in 20-µl aliquots of the neutralized extracts were separated and quantified using a reverse-phase ion-pairing, gradient HPLC method as previously described (17).

JC-1 fluorescence. JC-1 was added to the suspensions for 15 min at the end of reoxygenation followed by sampling for measurement of fluorescence at 488 nm excitation and 535 and 595 nm emission as previously described (19).

Measurement of {Delta}{Psi}m with safranin O. Measurements were done as previously described (18, 19). At the end of the experimental period, tubules were pelleted, washed three times in an ice-cold solution containing (in mM) 110 NaCl, 25 Na-HEPES, pH 7.2, 1.25 CaCl2, 1.0 MgCl2, 1.0 KH2PO4, 3.5 KCl, 5.0 glycine, 5% polyethylene glycol (average mol wt 8,000), and 2.0 mg/ml bovine gelatin, and maintained at 4°C until use. For the safranin O uptake measurements, the tubules were resuspended at a final concentration of 0.10–0.15 mg/ml in an intracellular buffer-type solution containing 120 mM KCl, 1 mM KH2PO4, 2 mM EGTA, 5 µM safranin O, 100 µg digitonin/mg protein, 4 mM potassium succinate, and 10 mM K-HEPES, pH 7.2 at 37°C (solution B) supplemented with other experimental reagents that are described with the data. Succinate was used as the substrate during safranin O uptake in these studies because it supports {Delta}{Psi}m slightly better in both permeabilized normoxic control tubules and after H/R than {alpha}-KG/MAL (19). Fluorescence was followed at 485 nm excitation, 586 nm emission using Photon Technology International (Lawrenceville, NJ) Deltascan and Alphascan fluorometers, equipped with temperature-controlled, magnetically stirred cuvette holders. Uptake of safranin O in the matrix of energized mitochondria results in quenching of its fluorescence, so the measured signal decreases. To make it easier to follow the tracings relative to high and low {Delta}{Psi}m, they are inverted in Figs. 1–12. Initial safranin O uptake is slower than its subsequent movements because of the time required to rewarm and permeabilize the tubules (19). For studies done on normoxic, control tubules, all experiments used tubules from the same suspension, so variability between cuvettes was limited to pipetting differences and was under 1–2%. For studies comparing tubules subjected to different experimental conditions in separate flasks before sampling for safranin O, protein concentrations were targeted to be the same as for the normoxic control and were always within 10% of each other. For studies where average changes of net safranin O uptake between groups were compared, values were calculated as differences of fluorescence between time 0 and the point of maximal uptake during the period of observation and were factored for the amount of tubule protein in the sample. Safranin O uptake by isolated mitochondria is well documented to be a linear function of {Delta}{Psi}m at >80–90 mV (1, 11, 34). Using valinomycin to induce K+ diffusion potentials, we confirmed this to be true for mitochondria in permeabilized tubules (Fig. 1). However, K+ diffusion calibrations were not available for every experiment, so data are reported in terms of safranin O uptake only.



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  		Fig. 1. Relationship between safranin O uptake by permeabilized tubules and mitochondrial membrane potential ({Delta}{Psi}m). Tubules were permeabilized in a K+-free medium containing 200 mM sucrose, 2 mM EGTA, 10 mM Na-HEPES, 1 mM sodium phosphate, 4 mM sodium succinate, 100 µg digitonin/mg protein, 0.5 mg/ml delipidated BSA (dBSA), and 5 µM safranin O, pH 7.2 at 37°C. When safranin O uptake was complete, 100 ng/ml valinomycin was added, followed by incremental additions of KCl to produce the indicated diffusion potentials calculated using the Nernst equation and assuming a matrix K+ concentration of 150 mM. Carbonyl cyanide p-(trifluoromethoxyphenyl)hydrazone (FCCP, 5 µM) was added to end the sequence. The inset compares the calculated diffusion potentials (mV) with the corresponding safranin O uptakes measured as the differences between fluorescence at each point and time 0.

 


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  		Fig. 12. Effects of dBSA and {alpha}-KG/MAL during H/R on ATP and mitochondrial energization of unpermeabilized tubules. Tubules were subjected to either normoxic control incubation or to 60-min hypoxia then 60-min reoxygenation followed by measurement of ATP levels (A) and 595/535-nm JC-1 ratio (B). Test agents were present in the incubation medium either only during hypoxia or only during reoxygenation as indicated. Values are percentages of those for normoxic control tubules incubated for the same durations given as means ± SE for 5–7 experiments. *P < 0.01 vs. corresponding NA group, except ATP and JC-1 ratio for dBSA addition during hypoxia, which were P < 0.05. #P < 0.05 vs. corresponding group with addition of test agent during hypoxia. Average normoxic control values were as follows: ATP = 15.8 ± 1.2 nmol/mg protein; 595/535-nm JC-1 ratio = 14.0 ± 0.8.

 
ATP production by permeabilized tubules. Tubules were suspended in solution B as for safranin O uptake measurements with further addition to solution B of the following reagents to couple ATP production to conversion of NADP to highly fluorescent NADPH: 10 mM glucose, 10 U/ml hexokinase, 0.2 mM NADP, 5 U/ml glucose-6-phosphate dehydrogenase, 30 µM diadenosine-5'-pentaphosphate to inhibit adenylate kinase (49), and 1 mM ouabain. The rate of ATP production was followed as formation of NADPH at 360 nm excitation/450 nm emission. Standard curves to calibrate fluorescence changes relative to ATP formation were done in the absence of tubules by addition of known amounts of ATP and NADPH, which gave identical results. Also, experiments using tubules were ended with addition of 5 µM ATP as an internal standard. Safranin O fluorescence was followed simultaneously at 465 nm excitation/586 nm emission.

Measurement of NEFA. Lipids were extracted at the end of hypoxia or H/R as previously described (42, 82) by Bligh and Dyer (4) except for the use of 2 M KCl instead of water to phase the layers. Tubule suspension (3 ml) was vortexed into an ice-cold solution of 7.5 ml methanol and 3.75 ml chloroform. After 15 min with additional mixing several times, 3.75 ml ice-cold chloroform were added with vigorous mixing followed by 3.75 ml ice-cold 2 M KCl with mixing. The suspension was centrifuged to separate the two phases. The top layer was discarded, and the bottom chloroform layer containing the extracted lipids was carefully removed, dried down under N2, and stored at –80°C until assayed. For assays, lipids were washed in chloroform and concentrated to an 80-µl volume taking care not to lose any material. NEFA were assayed enzymatically using 20-µl volumes of sample by conversion to acyl-CoA esters using acyl-CoA synthetase followed by oxidation of the acyl-CoA by acyl-CoA oxidase with production of hydrogen peroxide that is then measured colorimetrically (NEFA-C kit; WAKO Chemicals). The assay detects heptanoate, precluding its use as a substrate, but does not detect butyrate so butyrate was used as the lipid substrate during reoxygenation for those studies.

Statistics. Paired and unpaired t-tests were used as appropriate. Where experiments consisted of multiple groups, they were analyzed statistically by ANOVA for repeated-measure or independent group designs as needed. Individual group comparisons for the multigroup studies were then made using the Holm-Sidak test for multiple comparisons (SigmaStat 3; SPSS, Chicago, IL). P < 0.05 was considered to be statistically significant. Data shown are either means ± SE of no less than three to five experiments or are tracings representative of the behavior in that many experiments.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
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Tubule cell ATP levels and NEFA after H/R. Tubules were studied during 60-min hypoxia followed by 120-min reoxygenation with and without supplemental {alpha}-KG/MAL. During 60 min hypoxia without extra substrates, ATP levels decreased to 3.4% of the normoxic control level (Fig. 2A). With no extra substrates during hypoxia or reoxygenation (H/R NES groups in Fig. 2A), ATP recovered to 10.0% of normoxic control levels at 60-min reoxygenation and to 24.5% at 120 min of reoxygenation. Supplementation with {alpha}-KG/MAL during only reoxygenation (H NES, R {alpha}-KG/MAL) increased ATP recovery to 50% at 60-min reoxygenation in these studies, with complete recovery at 120-min reoxygenation. Inclusion of {alpha}-KG/MAL in the medium during hypoxia maintained slightly but significantly higher ATP levels (4.29% of control with {alpha}-KG/MAL vs. 3.42% without, P < 0.05) and promoted ATP recovery during the first 60 min of reoxygenation (H {alpha}-KG-MAL, R NES). The most rapid recovery of ATP was seen in tubules supplemented with {alpha}-KG/MAL during both hypoxia and reoxygenation (H {alpha}-KG/MAL, R {alpha}-KG/MAL), as described previously (17–19, 83–85).



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  		Fig. 2. Tubule cell ATP and nonesterified fatty acid levels during hypoxia and reoxygenation. Tubules were subjected to 60-min hypoxia (H) with either no extra substrates (NES) or 4 mM {alpha}-ketoglutarate + malate ({alpha}-KG/MAL) followed by 120 min of reoxygenation (R) with either no extra substrates or {alpha}-KG/MAL. They were sampled for ATP (A) and nonesterified fatty acids (B) at the end of hypoxia (60' H data) and than again after 60 and 120 min of reoxygenation (60' R and 120' R data). Values are means ± SE for n = 3–4 experiments. P < 0.05 vs. corresponding NES flask (*) and vs. value of same flask for the previous sampling period (#).

 
NEFA increased from 0.2 ± 0.1 nmol/mg protein in normoxic control tubules to 17.4 ± 0.4 at the end of 60-min hypoxia (Fig. 2B). {alpha}-KG/MAL during hypoxia reduced NEFA accumulation by 53%. NEFA decreased during the first 60 min of reoxygenation irrespective of the presence of {alpha}-KG/MAL. However, with {alpha}-KG/MAL, the magnitude of decrease was greater so that NEFA levels in the "H NES, R {alpha}-KG/MAL" group were 47% of those in the tubules with no extra substrate during hypoxia or reoxygenation (H/R NES). A further decrease occurred during the second 60 min of reoxygenation in the H NES, R {alpha}-KG/MAL group. In the "H/R NES" group, NEFA did not decrease during the second 60 min of reoxygenation. Tubules that had received {alpha}-KG/MAL during hypoxia ("H {alpha}-KG/MAL, R NES" and "H {alpha}-KG/MAL, R {alpha}-KG-MAL" groups), which suppressed NEFA accumulation at that time, had the most complete recovery of NEFA levels during subsequent reoxygenation.

Effects of NEFA addition to permeabilized tubules on {Delta}{Psi}m. The major NEFA that accumulate during hypoxia are palmitate, stearate, linoleate, oleate, and arachidonate (82). Figure 3 shows the effects of oleate on {Delta}{Psi}m measured with safranin O and on ATP production in permeabilized tubules. Oleate (4–5 µM; 4 µM = 26 nmol/mg protein) maximally decreased {Delta}{Psi}m (Fig. 3, A and B) and blocked ATP production (Fig. 3C). As little as 0.5 µM had a substantial effect, and some decrease of {Delta}{Psi}m was consistently seen at concentrations as low as 0.25 µM (1.6 nmol/mg protein, data not shown). The decreases of {Delta}{Psi}m and inhibition of ATP production were prevented and reversed by 0.5 mg/ml (7.57 µM) dBSA. Arachidonate, linoleate, and palmitate, like oleate, all decreased {Delta}{Psi}m, with arachidonate and linoleate being slightly more active than oleate and palmitate slightly less (data not shown). In the absence of exogenous oleate, tubules also displayed some improvement of {Delta}{Psi}m and ATP production with dBSA, consistent with a contribution of endogenous NEFA to lowering {Delta}{Psi}m.



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  		Fig. 3. Oleate-induced decreases of {Delta}{Psi}m and ATP production. A: representative fluorescence tracings of safranin O uptake by mitochondria in permeabilized tubules starting with either no additions (NA) to the regular solution B safranin O uptake medium, 0.5 mg/ml dBSA, 0.5 µM oleate, 5 µM oleate, or 5 µM oleate + dBSA. dBSA was also added at 750 s during the experiments that did not start with it. The relative positions of the tracings at 750 s are the same as the order in the legend. B: group data for net safranin O uptake during studies like those in A. Labeling is the same as for A. Data are safranin O uptakes measured as the change in fluorescence between time 0 and the point of maximal uptake during the period before addition of dBSA for each condition as a percentage of maximum safranin O uptake by the corresponding sample starting with dBSA and no oleate. Values are means ± SE for 7 experiments. *P < 0.01 vs. dBSA group. C: ATP production rates during safranin O uptake with and without oleate and dBSA. Conditions and labeling were the same as in A except for the inclusion of glucose, hexokinase, NADP, glucose-6-phosphate deyhydrogenase, diadenosine-5'-pentaphosphate, and ouabain and monitoring of NADPH fluorescence at 360-nm excitation/450-nm emission to follow ATP production. Tracing is truncated to start at 400 s of incubation to more clearly show the period of ATP production beginning with addition of 100 µM ADP at 600 s. Experiments ended with addition 5 µM ATP at 900 s, showing the response to a known amount of ATP.

 
Tubule cell ATP levels and mitochondrial energization measured with JC-1 after H/R. As in the studies in Fig. 2, 60 min of hypoxia resulted in decreased ATP recovery that was improved by {alpha}-KG/MAL during reoxygenation (Fig. 4A). Extending the hypoxic period to 120 min resulted in almost no ATP recovery during the subsequent 60 min of reoxygenation, which was only minimally improved by {alpha}-KG/MAL during reoxygenation (data not shown). Inclusion of {alpha}-KG/MAL during both 120 min hypoxia and the 60 min of reoxygenation gave some recovery of ATP, but it remained weak (Fig. 4A). The 595/535-nm JC-1 ratios (Fig. 4B) and the protein-factored 595 nm signal (Fig. 4C) followed patterns similar to the ATP levels but, as described previously (19), were less impaired relative to the normoxic control values than the ATP levels.



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  		Fig. 4. Tubule cell ATP and mitochondrial energization measured using 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazocarbocyanine iodide (JC-1) after H/R. Tubules were subjected to either 60 or 120 min of hypoxia followed by 60-min reoxygenation or to normoxic control incubation for the same durations. The 60-min hypoxic tubules had either no extra substrate during both hypoxia and reoxygenation (H/R NES) or no extra substrate during hypoxia and {alpha}-KG/MAL during reoxygenation (H NES, R {alpha}-KG/MAL); 120 min hypoxic tubules had either no extra substrate during both hypoxia and reoxygenation (H/R NES) or {alpha}-KG/MAL during both hypoxia and reoxygenation (H {alpha}-KG/MAL, R {alpha}-KG/MAL). ATP levels (A), the ratio of JC-1 emissions at 595/535 nm (B), and the protein-factored 595-nm JC-1 emission (C) were measured at the end of reoxygenation. Values are percents of those for the normoxic control tubules given as means ± SE for 5 experiments using 60-min hypoxia and 4 experiments using 120-min hypoxia. *P < 0.01 vs. H/R NES group with the same duration of hypoxia except 120' H/60' R {alpha}-KG/MAL JC-1 595-nm value, which was 0.02. #P < 0.01 vs. corresponding 60-min hypoxia groups except 120' H/60' R NES ATP and JC-1 595-nm values, which were 0.04. Average normoxic control values were as follows: ATP = 16.2 ± 1.5 nmol/mg protein; 595/535-nm JC-1 ratio = 23.8 ± 2.4; protein-factored 595-nm JC-1 signal [in 106 counts/s(cps)/mg protein] = 3.1 ± 0.4. All values for the H/R studies shown were significantly less than the corresponding normoxic controls.

 
Restoration of {Delta}{Psi}m and ATP production after H/R in permeabilized tubules with addition of dBSA. The deceased {Delta}{Psi}m indicated by the changes of JC-1 uptake at the end of H/R (Fig. 4) persists when tubules are permeabilized at the end of experimental maneuvers and {Delta}{Psi}m is assessed using safranin O (18, 19). Safranin O uptake by permeabilized tubules provides additional opportunities to evaluate the determinants of {Delta}{Psi}m and to introduce modifying maneuvers (e.g., Fig. 3 and Refs. 18 and 19). {Delta}{Psi}m measured at the end of 60 min hypoxia and 60 min reoxygenation using safranin O uptake was markedly decreased relative to normoxic controls (Figs. 5A and 6A), and tubules that were supplemented with {alpha}-KG/MAL during reoxygenation had better safranin O uptake than unsupplemented tubules (Fig. 5A). Uptake was not improved by addition of cytochrome c to the safranin O uptake medium (Fig. 5C) but, consistent with previous observations (18, 19), was increased, though not normalized, by inclusion of 2 mM ATP (Fig. 5B). Under every condition studied, including the normoxic controls, 0.5 mg/ml dBSA improved {Delta}{Psi}m (Figs. 5, A and B, and 6A) and ATP production (Fig. 6, B and C). After H/R, safranin O uptake was restored by inclusion of dBSA to an average of >90% of the level in normoxic controls (Fig. 7). The benefit of dBSA was seen even when it was added late during safranin O uptake (Fig. 8). Albumin can be an antioxidant (29); however, the effect of dBSA was not reproduced by the superoxide scavenger Tiron (37), the superoxide dismutase mimetic Tempol (43), or GSH (Fig. 9).



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  		Fig. 5. Effects of ATP, dBSA, and cytochrome c on {Delta}{Psi}m after 60-min hypoxia and 60-min reoxygenation measured using safranin O uptake. Representative fluorescence tracings of mitochondrial safranin O uptake in tubules subjected to normoxic control (NC) incubation or to 60-min hypoxia and 60-min reoxygenation (H/R) with no extra substrate during hypoxia and either no extra substrate or {alpha}-KG/MAL during reoxygenation as described for the 60-min hypoxia studies in Fig. 4, which included these experiments. A and B: effects of dBSA and ATP. Tracings are from the same experiment and are directly comparable. dBSA, 0.5 mg/ml delipidated BSA present during safranin O uptake. In all studies in B, 2 mM ATP was also present during safranin O uptake. C: effect of cytochrome c on normoxic control and hypoxia/reoxygenation tubules. Cytochrome c (10 µM) was present during safranin O uptake in the indicated experiments.

 


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  		Fig. 6. ATP production by permeabilized tubules after H/R. Tubules were subjected to 60-min hypoxia and 60-min reoxygenation or to normoxic control incubation. A and B: simultaneous measurements of {Delta}{Psi}m using safranin O uptake (A) and ATP production as NADPH fluorescence in the presence of glucose, hexokinase, NADP, glucose-6-phosphate deyhydrogenase, diadenosine-5'-pentaphosphate, and ouabain (B). The medium also contained 1 mM Mg2+ because Mg2+ was required for maximal F1FO-ATPase/ATP synthase activity after H/R (18). ADP (100 µM) was added at 600 s. Experiments ended with 5 µM ATP at 900 s, showing the NADPH response to a known amount of ATP. H/R NES, 60-min hypoxia + 60-min reoxygenation with no extra substrates. Mitochondrial safranin O uptake by the H/R NES tubules studied without dBSA is better than in the Fig. 5 experiments because injury was milder in this series of studies, and the extra reagents used for the ATP production measurements slightly improve safranin O uptake after H/R. C: group averages for ATP production rates in studies such as those done in A and B with additional comparison of behavior without and with 1 mM Mg2+. Values are means ± SE for n = 5–6 calculated as percentages of the ATP production rate of the corresponding normoxic control in the presence of dBSA and Mg2+. P < 0.01 vs. corresponding normoxic control groups (*) and vs. corresponding H/R dBSA groups (#). All +Mg values are significantly greater than the corresponding –Mg values. Mg2+ did not affect safranin O uptake (data not shown).

 


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  		Fig. 7. Group averages for studies of dBSA effects on net safranin O uptake after H/R. Tubules were subjected to either normoxic control incubation or to 60-min hypoxia and 60-min reoxygenation followed by measurement of mitochondrial safranin O uptake as in Fig. 5, A and B, which was from the same series of experiments. Data for each experimental condition are calculated as percentages of the safranin O uptakes by paired normoxic control samples with dBSA. NA, no experimental additions during safranin O uptake; ATP, 2 mM ATP present during safranin O uptake. Values are means ± SE for n = 5. P < 0.001 vs. normoxic control (*) and vs. corresponding condition without dBSA (+). P < 0.05 vs. corresponding H/R NES group (#).

 


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  		Fig. 8. Reversal of deceased {Delta}{Psi}m after H/R by late addition of dBSA during safranin O uptake. Representative fluorescence tracings of mitochondrial safranin O uptake in tubules subjected to 60-min hypoxia and 60-min reoxygenation (H/R) or normoxic control incubation as described for the studies in Fig. 5. dBSA (0.5 mg/ml) was added at 850 s of safranin O uptake.

 


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  		Fig. 9. Comparison of the effects of antioxidants and dBSA on H/R-induced dissipation of {Delta}{Psi}m. Tubules were subjected to 60-min hypoxia and 60-min reoxygenation without extra substrates followed by measurements of safranin O uptake as in Figs. 5 and 8. Additions of the indicated test agents were made after 250 s of safranin O uptake.

 
Figure 10 shows the results of studies addressing quantitative aspects of the dBSA effects during safranin O uptake. In normoxic control tubules, as little as 0.05 mg/ml (0.76 µM) was sufficient to maximize {Delta}{Psi}m (Fig. 10A). After H/R, full effects were not seen until 0.5 mg/ml (7.57 µM) dBSA (Fig. 10B). These data are consistent with the presence of higher levels of NEFA after H/R that require more of the binding capacity of the dBSA. To further assess this issue, normoxic control and H/R tubules were treated with 0.5 mg/ml dBSA during safranin O uptake, and then oleate was added incrementally until dissipation of {Delta}{Psi}m began (Fig. 10C). In the normoxic controls, an average of 21.9 ± 1.2 µM oleate was needed to produce a decrease of {Delta}{Psi}m in the presence of the dBSA. After H/R, 8.33 ± 0.83 of oleate was required (P < 0.001, n = 4).



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  		Fig. 10. Titrations of dBSA effects on {Delta}{Psi}m. Tubules were subjected to 60-min hypoxia and 60-min reoxygenation without extra substrates or to normoxic control incubation. A and B: dBSA at the concentrations indicated (mg/ml) was added at the points marked. C: dBSA (0.5 mg/ml, 7.57 µM) was present from the start of mitochondrial safranin O uptake. Oleate was added at the indicated points (concentrations given in µM) followed by additional dBSA (0.5 mg/ml). H/R was 60-min hypoxia and 60-min reoxygenation without extra substrates.

 
Given the robust effect of dBSA to virtually completely restore {Delta}{Psi}m in the 60-min hypoxia experiments (Figs. 5–10), we sought to determine whether benefit would be maintained with the more severe 120-min insult that results in almost no ATP recovery during the subsequent 60 min of reoxygenation (Fig. 4A). Mitochondria in permeabilized tubules had no detectable safranin O uptake after hypoxia for 120 min and reoxygenation without {alpha}-KG/MAL, irrespective of the presence of ATP during the safranin uptake period (Fig. 11, A and B). Absent safranin O uptake (Fig. 11, A and B), despite some persistence of {Delta}{Psi}m detectable by JC-1 (Fig. 4, B and C) in these 120-min hypoxia studies, is explained by the insensitivity of safranin O to {Delta}{Psi}m <80–90 mV (Fig. 1 and Ref. 34). Tubules that received {alpha}-KG/MAL during hypoxia and reoxygenation showed improvement of mitochondrial safranin O uptake when ATP was present, but uptake remained relatively low (Fig. 11B). Based on the concentration-dependence information in Fig. 10B for the effects of dBSA on the 60 min hypoxic tubules, we used 2.0 mg/ml dBSA for these 120 min studies. The dBSA almost completely normalized mitochondrial safranin O uptake under all conditions (Fig. 11, A–C).



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  		Fig. 11. Restoration of {Delta}{Psi}m after 120-min hypoxia and 60-min reoxygenation by dBSA. A and B: representative tracings of safranin O uptake by mitochondria in tubules subjected to normoxic control incubation or 120-min hypoxia followed by 60-min reoxygenation with either no extra substrates or {alpha}-KG/MAL (4 mM each) present during hypoxia and reoxygenation as described for the 120-min hypoxia studies in Fig. 4, which included these experiments. In all studies in B, 2 mM ATP was present during safranin O uptake. C: group averages for studies of dBSA effects on net safranin O uptake after 120-min hypoxia and 60-min reoxygenation. H/R conditions were as described for A and B. Data for each experimental condition are calculated as percentages of the net safranin O uptakes by paired normoxic control samples with dBSA. Values are means ± SE for n = 4. *P < 0.002 vs. normoxic control. #P < 0.003 vs. corresponding H/R NES group. +P < 0.001 vs. corresponding condition without dBSA.

 
Benefit of dBSA for recovery of mitochondrial energization and ATP by unpermeabilized tubules. Under the permeabilized conditions used for the studies in Figs. 5– 11, added dBSA potentially has direct access to the mitochondria. Figure 12 summarizes studies in which dBSA was tested and compared with {alpha}-KG/MAL for its efficacy in the absence of permeabilization. When present during 60 min of hypoxia, 10 mg/ml dBSA provided only a small amount of benefit compared with {alpha}-KG/MAL on the subsequent recovery of ATP and of mitochondrial energization assessed by JC-1 uptake during reoxygenation (Fig. 12). dBSA had much more benefit when provided during reoxygenation but was still somewhat less effective than {alpha}-KG/MAL. The combination of dBSA and {alpha}-KG/MAL during hypoxia was no more effective than {alpha}-KG/MAL alone, but the two maneuvers were additive when provided during reoxygenation (Fig. 12).


   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
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 REFERENCES
 
A number of prior studies have investigated the accumulation of NEFA during kidney ischemia and hypoxia of freshly isolated proximal tubules (Table 1). The results of the previous work and the current study are remarkably similar considering the different preparations and assays used, with levels of 7–30 nmol/mg protein reached at the end of 30- to 60-min insults. When oleate was added to permeabilized normal tubules (Fig. 3 and similar studies not shown), decreases of {Delta}{Psi}m were consistently detectable with 1.6 nmol/mg protein and were maximal at 26 nmol/mg protein. Thus total levels of NEFA in hypoxic tubules were in the range that produces substantial dissipation of {Delta}{Psi}m. In cells, most NEFA are considered to be bound and may not necessarily be available to interact with mitochondria (69). High levels of fatty acid-binding protein are present in kidney (35). However, the ability of dBSA to strongly restore {Delta}{Psi}m in the tubules after H/R indicates that, despite the intracellular binding by fatty acid-binding proteins, the increased levels of NEFA measured in the whole cells after H/R and ischemia-reperfusion are available to interact with mitochondria and modify their function.


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  		Table 1. NEFA levels in kidney tissue and isolated tubules during hypoxia-ischemia and reoxygenation-reperfusion

 
Deleterious effects of NEFA on mitochondrial function have been recognized for over five decades (12, 13, 23, 25, 38, 53, 62–64, 90, 91) and can involve a number of primary and secondary actions of the NEFA and their metabolites (51, 71, 75, 90, 94). Most recent data favor the concept that deenergization under the conditions of the present studies derives from the ability of long-chain free fatty acids to act as rapidly reversible protonophoric uncouplers by entering the mitochondrial matrix in their protonated, nonpolar form followed by dissociation of the proton and transport of the deprotonated fatty acid ion out of the matrix by anion carriers, including the adenine nucleotide translocase, the glutamate/aspartate cotransporter, the dicarboxylate and tricarboxylate carriers, and the uncoupling proteins (31, 45, 72, 93). This results in a {Delta}{Psi}m-dissipating, sustained cycle of proton entry to the matrix with resultant decreased ATP production despite continued respiration (i.e., uncoupling).

Initial work with NEFA-induced mitochondrial dysfunction identified it as the major factor in spontaneous deterioration of isolated mitochondria ("aging"; see Refs. 12, 13, 23, 25, 38, 53, 62–64, 90, 91). As a result, dBSA came to be widely included during both preparation of mitochondria and in the media used for their study, usually without assessment of function in the absence of dBSA, thus obscuring the contribution of NEFA to pathological states. As in the present work, early studies of mitochondria isolated from ischemic liver in the absence of dBSA documented substantial mitochondrial dysfunction that could be entirely reversed by inclusion of dBSA (6, 7). Mitochondria isolated in the absence of dBSA from ischemic heart displayed increased rates of proton leak that were corrected by dBSA (9). Kidney mitochondria isolated during the first 3 h of reperfusion showed little functional abnormality but were prepared (89) or studied (77) only in the presence of dBSA. The interpretation of studies of isolated mitochondria in general is subject to some uncertainty as to whether observed effects were present before cellular disruption or have been modified by redistribution and loss of metabolites during isolation. The present work using permeabilized cells and also documenting efficacy of dBSA in unpermeabilized cells provides a unique demonstration of the critical role of NEFA in mitochondrial dysfunction.

In normoxic control tubules, decreases of {Delta}{Psi}m were readily detectable with addition of 0.25 µM oleate (1.6 nmol/mg protein). Albumin has seven binding sites for fatty acids (52) of which three share the highest affinity (8, 14, 74). In the presence of 7.57 µM dBSA, addition of 22 µM oleate was required to begin to dissipate {Delta}{Psi}m in normal mitochondria, suggesting that only these three highest-affinity sites bind fatty acids sufficiently to keep the free fatty acid concentration available to the mitochondria under the levels required for uncoupling. After H/R, the amount of exogenous oleate needed to decrease {Delta}{Psi}m in the presence of 7.57 µM dBSA was decreased to 8 µM, which would imply the presence of 14 µM excess fatty acid in those samples. Based on the concentration of total tubule protein during the safranin O measurements of 0.15 mg/ml, this would be 93 nmol fatty acid/mg protein, which is far greater than the 7 nmol/mg protein measured during reoxygenation or the 17 nmol/mg protein at the end of hypoxia. This indicates that, despite the permeabilization conditions, the efficacy of dBSA for trapping endogenously generated fatty acids is reduced or sensitivity to the NEFA has increased.

dBSA added to suspensions of intact tubules (Fig. 12) also alleviated mitochondrial deenergization and improved recovery of ATP during reoxygenation. Not surprisingly, and in contrast to the nearly complete recovery produced in the permeabilized cells, dBSA addition to intact cells was only partially protective, but its effect was substantial. The weaker effects of dBSA on NEFA-induced mitochondrial dysfunction in the intact cells are likely because of restriction of most of the protein to the extracellular space, limiting its effects to remove NEFA acting on the mitochondria. Although the dBSA can be endocytosed (20), that process will be constrained in the isolated tubules after H/R by both ATP depletion and limited access to the apical surface. Internalized albumin remains compartmentalized and not fully available to bind mitochondria-associated NEFA. Mouse proximal tubule cells in primary culture were protected by dBSA during cyanide-induced ATP depletion (70), but, in multiple prior studies of dBSA effects using intact freshly isolated tubules, dBSA was either ineffective or toxic (26, 95–97). The different behavior in the present studies is likely because of the presence of glycine, which is abundant in vivo and is cytoprotective by acting to suppress development of a plasma membrane pore that forms during ATP depletion states and mediates rapid cell killing (16, 48, 79). Despite worsening ATP depletion during reoxygenation (96), as expected from their deenergizing effects, exogenous, unsaturated fatty acids decrease ATP depletion-associated plasma membrane damage in isolated tubules (2, 46, 95, 96). This has been attributed to an effect of the fatty acids to suppress cellular phospholipases (2, 95), but it could conceivably also involve actions at the plasma membrane similar to those of glycine, perhaps at the same site. The prior studies that reported toxic effects of dBSA or failed to see benefit were done without glycine (26, 95–97) and therefore were subject to aggravating effects on plasma membrane damage from removal by the dBSA of membrane-associated, unsaturated, fatty acids. This would have obscured any later benefit on energetic function during reoxygenation from lowering intracellular NEFA.

Supplementing intact tubules with specific tricarboxylic acid cycle metabolites during hypoxia or reoxygenation strongly modifies development of the energetic deficit (17–19, 83–85). {alpha}-KG/MAL can produce ATP anaerobically via substrate level phosphorylation and is beneficial when provided during either hypoxia or reoxygenation (Figs. 2, 4, and 12 and Refs. 83 and 84). Succinate, which does not produce ATP anaerobically, is of benefit only during reoxygenation (83, 84). {alpha}-KG/MAL decreased NEFA accumulation during both the hypoxic period and during reoxygenation by 50% (Fig. 2). The effect during reoxygenation is complex because it occurs in the setting of generalized metabolic recovery with large increases of ATP levels. The effect of {alpha}-KG/MAL during hypoxia is more striking because the substrates produced an even larger change of NEFA levels in the setting of profound ATP depletion. {alpha}-KG/MAL maintains slightly higher ATP levels during hypoxia by supporting mitochondrial anaerobic ATP production (83, 84), and this effect was observed in the present studies (Fig. 2A). It has previously been shown using ATP-depleted LLC-PK1 cells that, at low ATP levels, very small increments of ATP can suppress the large increases of NEFA that occur with maximal ATP depletion (76), likely by promoting reesterification, inhibiting phospholipase activation (50, 78), or a combination of both effects. Taken in the context of its ability to promote anaerobic ATP production and maintain small increments of ATP levels during hypoxia, the striking effect of {alpha}-KG/MAL to decrease NEFA at that time may represent an example of the effect of very low levels of ATP to limit NEFA accumulation under physiologically more relevant injury conditions. Although we believe that these considerations strongly argue that the effect of {alpha}-KG/MAL on NEFA results from promotion of reesterification, we cannot exclude a contribution from inhibition of phospholipase activity by {alpha}-KG/MAL either via a direct effect of the substrates on the enzyme or indirectly by maintenance of ATP. Further studies will be needed to assess these possibilities. Irrespective of the mechanism for the {alpha}-KG/MAL-induced decreases of NEFA, given the importance of NEFA for the energetic deficit, the decreases of their levels produced by {alpha}-KG/MAL are likely to play a major role in the protective effects of the substrates.

The effects of {alpha}-KG/MAL and dBSA on intact tubules were additive, but showed different patterns. In contrast to {alpha}-KG-MAL, which provides similar benefit to promote recovery during reoxygenation when present during either hypoxia or reoxygenation (Fig. 12 and Ref. 84), dBSA was much more effective during reoxygenation than during hypoxia. Unlike {alpha}-KG/MAL, dBSA did not increase ATP during hypoxia. In the studies in Fig. 12, ATP levels decreased to 2.5 ± 0.1% of normoxic control values during hypoxia. dBSA did not significantly change this (3.0 ± 0.1% of normoxic control), but ATP was significantly increased (P < 0.05) by {alpha}-KG/MAL (4.4 ± 0.3%) and {alpha}-KG/MAL + dBSA (4.4 ± 0.3%). Whether the weaker effect of dBSA provided during hypoxia to promote recovery during reoxygenation is related to lesser ability to remove NEFA from the cell in the absence of normal intracellular trafficking and metabolism or to its inability to promote anaerobic ATP production and maintain slightly higher ATP during hypoxia like {alpha}-KG/MAL remains to be determined. In this regard, however, it is of interest that {alpha}-KG/MAL + dBSA during hypoxia did not promote recovery any better than {alpha}-KG/MAL during hypoxia (Fig. 12).

In vivo, most fatty acids circulate bound to albumin (65, 73). Amounts bound vary, but the ratio of total NEFA/albumin is generally <1, leaving substantial residual binding capacity (65). Reperfusion with dBSA can remove accumulated NEFA after ischemia of the intact kidney (24, 42). The studies here with the unpermeabilized tubules indicate that lowering NEFA with extracellular dBSA can have a substantial beneficial effect on mitochondrial function and, as a result, cellular recovery. However, even with the optimal delivery conditions possible for isolated tubules and the use of delipidated albumin, protection of unpermeabilized tubules was incomplete. Under in vivo reperfusion conditions where circulating albumin is not delipidated, where the cellular burden of NEFA is higher because of the density of the tissue, and where delivery of albumin may be limited by reperfusion abnormalities, the benefit is likely to be decreased. Moreover, without sufficient glycine, dBSA could aggravate injury by removing protective unsaturated fatty acids from the plasma membrane (2, 46, 95). In this regard, it is of interest that albumin infusion is highly neuroprotective against focal cerebral ischemic damage, but this has been attributed to effects of the albumin to deliver and replenish polyunsaturated fatty acids lost from cell membranes during ischemia (15).

These considerations also bear on organ harvesting and preservation procedures where use of delipidated albumin is possible and the composition of perfusion solutions can be defined. Early approaches to machine perfusion used various plasma and serum protein preparations, including albumin as perfusate (88), but difficulties in their preparation and standardization, cost, and availability of effective alternate protein-free solutions led to discontinuation of the practice. The studies here suggest that use of delipidated albumin as a fatty acid binder in preservation solutions merits reconsideration, but as part of a combined strategy that would include otherwise optimized preservation solutions, supplemented with protective substrates for their separate actions to decrease NEFA accumulation and glycine to avoid any deleterious effects of removing unsaturated fatty acids that are protecting against plasma membrane damage.

Based on measurements of lactate dehydrogenase release, several compounds with activity as phospholipase inhibitors have been reported to ameliorate hypoxic injury in isolated proximal tubules, including mepacrine (10), dibucaine (10), bromoenol lactone (61), and ONO-RS-082 (95). However, these agents and other phospholipase inhibitors have not been useful for studying the role of NEFA in our model. Benefit of these compounds for metabolic recovery in the prior studies was not described, and their effects to limit NEFA accumulation were either minimal (10), absent (60), or not reported. In our rabbit tubule preparation, neither bromoenol lactone nor ONO-RS-082 improved the energetic deficit, but they deenergized mitochondria and decreased ATP under normoxic conditions (data not shown), so benefit from phospholipase inhibition could have been obscured. Although butacaine slightly improves the energetic deficit (19), we have not observed consistent effects of mepacrine or dibucaine (data not shown). The NEFA increases in our studies occur despite the use of pH 6.9 during hypoxia, which mildly inhibits NEFA accumulation (82), probably by decreasing Ca2+-dependent phospholipase activity (60).

Our experiments are relatively short term, with durations of hypoxia and reoxygenation totaling up to 180 min. However, recent studies by Portilla and coworkers and others (28, 39, 54, 56, 58, 59) have established that defects of both mitochondrial and peroxisomal {beta}-oxidation of fatty acids persist for prolonged periods, are further aggravated by downregulation of enzyme gene transcription controlled by peroxisome proliferator-activated receptor-{alpha}, and can be alleviated by peroxisome proliferator-activated receptor-{alpha} ligands in both ischemia-reperfusion and toxic models with improvement of renal function and prevention of necrotic and apoptotic cell death of the proximal tubule. Given the multiple potential actions of fatty acids and their metabolites in cells, it is not possible to attribute this global benefit of stimulation of fatty acid oxidation simply to alleviation of mitochondrial dysfunction, but, considering the central roles of mitochondria in both energetics and cell death regulation, the NEFA effects on them are likely to be contributing.

In summary, we have shown that the energetic deficit that develops in proximal tubules during H/R and plays a pivotal role in their ability to recover can be explained by NEFA-induced mitochondrial deenergization. Moreover, the promotion of recovery by tricarboxylic acid cycle metabolites such as {alpha}-KG/MAL is also likely mediated in large part by the same mechanism, since the substrates strikingly decrease NEFA accumulation, even during hypoxia. These observations provide unique insight into a primary process that accounts for the tubule injury that develops during ischemic acute failure and the nature of the cellular response to ischemia in general.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
These studies were supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-34275 and the Department of Veterans' Affairs.


   ACKNOWLEDGMENTS
 
We thank Tiffany Ostrowski for technical assistance and Dr. M. A. Venkatachalam for helpful discussions.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. M. Weinberg, Nephrology Division, Dept. of Internal Medicine, Rm. 1560, MSRB II, Univ. of Michigan Medical Center, Ann Arbor, MI 48109-0676 (e-mail: wnberg{at}umich.edu)

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.

1 We use "deenergization" in this paper to refer to decreased proton motive force across the inner mitochondrial membrane. Although we directly measure only changes of {Delta}{Psi}m, it accounts for >80% of the proton motive force in mitochondria respiring on succinate or complex I-dependent substrates (36) under conditions similar to those used in the present studies, and we have confirmed this to be the case in our model under both normal and injury conditions (data not shown). Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Akerman KE and Wikstrom MK. Safranine as a probe of the mitochondrial membrane potential. FEBS Lett 68: 191–197, 1976.[CrossRef][Web of Science][Medline]
  2. Alhunaizi AM, Yaqoob MM, Edelstein CL, Gengaro PE, Burke TJ, Nemenoff RA, and Schrier RW. Arachidonic acid protects against hypoxic injury in rat proximal tubules. Kidney Int 49: 620–625, 1996.[Web of Science][Medline]
  3. Bernardi P, Penzo D, and Wojtczak L. Mitochondrial energy dissipation by fatty acids. Mechanisms and implications for cell death. Vitam Horm 65: 97–126, 2002.[Web of Science][Medline]
  4. Bligh EG and Dyer WJ. A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37: 911–917, 1959.[Medline]
  5. Bodrova ME, Brailovskaya IV, Efron GI, Starkov AA, and Mokhova EN. Cyclosporin A-sensitive decrease in the transmembrane potential across the inner membrane of liver mitochondria induced by low concentrations of fatty acids and Ca2+. Biochemistry (Mosc) 68: 391–398, 2003.[CrossRef][Medline]
  6. Boime I, Smith EE, and Hunter FE Jr. Stability of oxidative phosphorylation and structural changes of mitochondria in ischemic rat liver. Arch Biochem Biophys 128: 704–715, 1968.[CrossRef][Web of Science][Medline]
  7. Boime I, Smith EE, and Hunter FE Jr. The role of fatty acids in mitochondrial changes during liver ischemia. Arch Biochem Biophys 139: 425–443, 1970.[CrossRef][Web of Science][Medline]
  8. Bojesen IN and Bojesen E. Binding of arachidonate and oleate to bovine serum-albumin. J Lipid Res 35: 770–778, 1994.[Abstract]
  9. Borutaite V, Morkuniene R, Budriunaite A, Krasauskaite D, Ryselis S, Toleikis A, and Brown GC. Kinetic analysis of changes in activity of heart mitochondrial oxidative phosphorylation system induced by ischemia. J Mol Cell Cardiol 28: 2195–2201, 1996.[CrossRef][Web of Science][Medline]
  10. Bunnachak D, Almeida ARP, Wetzels JFM, Gengaro P, Nemenoff RA, Burke TJ, and Schrier RW. Ca2+ uptake, fatty acid, and LDH release during proximal tubule hypoxia: effects of mepacrine and dibucaine. Am J Physiol Renal Fluid Electrolyte Physiol 266: F196–F201, 1994.[Abstract/Free Full Text]
  11. Cancherini DV, Trabuco LG, Reboucas NA, and Kowaltowski AJ. ATP-sensitive K+ channels in renal mitochondria. Am J Physiol Renal Physiol 285: F1291–F1296, 2003.[Abstract/Free Full Text]
  12. Chefurka W. Oxidative phosphorylation in in vitro aged mitochondria. I. Factors controlling the loss of dinitrophenol-stimulated adenosine triphosphatase activity and respiratory control in mouse liver mitochondria. Biochemistry 5: 3887, 1966.
  13. Chefurka W and Dumas T. Oxidative phosphorylation in in vitro aged mitochondria. II. Dinitrophenol-stimulated adenosine triphosphatase activity and fatty acid content of mouse liver mitochondria. Biochemistry 5: 3904, 1966.
  14. Cistola DP, Small DM, and Hamilton JA. C-13 Nmr-studies of saturated fatty-acids bound to bovine serum-albumin. I. The filling of individual fatty-acid binding-sites. J Biol Chem 262: 10971–10979, 1987.[Abstract/Free Full Text]
  15. de Turco EBR, Belayev L, Liu YT, Busto R, Parkins N, Bazan NG, and Ginsberg MD. Systemic fatty acid responses to transient focal cerebral ischemia: influence of neuroprotectant therapy with human albumin. J Neurochem 83: 515–524, 2002.[CrossRef][Web of Science][Medline]
  16. Dong Z, Patel Y, Saikumar P, Weinberg JM, and Venkatachalam MA. Development of porous defects in plasma membranes of ATP-depleted Madin-Darby canine kidney cells and its inhibition by glycine. Lab Invest 78: 657–668, 1998.[Web of Science][Medline]
  17. Feldkamp T, Kribben A, Roeser NF, Senter RA, Kemner S, Venkatachalam MA, Nissim I, and Weinberg JM. Preservation of complex I function during hypoxia-reoxygenation-induced mitochondrial injury in proximal tubules. Am J Physiol Renal Physiol 286: F749–F759, 2004.[Abstract/Free Full Text]
  18. Feldkamp T, Kribben A, and Weinberg JM. F1F0-ATPase activity and ATP dependence of mitochondrial energization in proximal tubules after hypoxia/reoxygenation. J Am Soc Nephrol 16: 1742–1751, 2005.[Abstract/Free Full Text]
  19. Feldkamp T, Kribben A, and Weinberg JM. Assessment of mitochondrial membrane potential in proximal tubules after hypoxia-reoxygenation. Am J Physiol Renal Physiol 288: F1092–F1102, 2005.[Abstract/Free Full Text]
  20. Gekle M. Renal tubule albumin transport. Annu Rev Physiol 67: 573–594, 2005.[CrossRef][Web of Science][Medline]
  21. Green DR and Kroemer G. The pathophysiology of mitochondrial cell death. Science 305: 626–629, 2004.[Abstract/Free Full Text]
  22. Halestrap A. Biochemistry: a pore way to die. Nature 434: 578–579, 2005.[CrossRef][Medline]
  23. Heinski DR and Cooper C. Studies on the action of bovine serum albumin on aged rat liver mitochondria. J Biol Chem 235: 3573–3579, 1960.[Free Full Text]
  24. Hsueh W and Needleman P. Sites of lipase activation and prostaglandin synthesis in isolated, perfused rabbit hearts and hydronephrotic kidneys. Prostaglandins 16: 661–681, 1978.[CrossRef]
  25. Hulsmann WC, Elliott WB, and Slater EC. The nature and mechanism of action of uncoupling agents present in mitochrome preparations. Biochim Biophys Acta 39: 267–276, 1960.[Medline]
  26. Humes HD, Nguyen VD, Cieslinski DA, and Messana JM. The role of free fatty acids in hypoxia-induced injury to renal proximal tubule cells. Am J Physiol Renal Fluid Electrolyte Physiol 256: F688–F696, 1989.[Abstract/Free Full Text]
  27. Hunter DR, Haworth RA, and Southard JH. Relationship between configuration, function, and permeability in calcium-treated mitochondria. J Biol Chem 251: 5069–5077, 1976.[Abstract/Free Full Text]
  28. Huss JM, Levy FH, and Kelly DP. Hypoxia inhibits the peroxisome proliferator-activated receptor alpha/retinoid X receptor gene regulatory pathway in cardiac myocytes — a mechanism for O2-dependent modulation of mitochondrial fatty acid oxidation. J Biol Chem 276: 27605–27612, 2001.[Abstract/Free Full Text]
  29. Iglesias J, Abernethy VE, Wang Z, Lieberthal W, Koh JS, and Levine JS. Albumin is a major serum survival factor for renal tubular cells and macrophages through scavenging of ROS. Am J Physiol Renal Physiol 277: F711–F722, 1999.[Abstract/Free Full Text]
  30. Jezek P, Zackova M, Ruzicka M, Skobisova E, and Jaburek M. Mitochondrial uncoupling proteins: facts and fantasies. Physiol Res 53: S199–S211, 2004.[Web of Science][Medline]
  31. Kamp F and Hamilton JA. pH gradients across phospholipid-membranes caused by fast flip-flop of unionized fatty-acids. Proc Natl Acad Sci USA 89: 11367–11370, 1992.[Abstract/Free Full Text]
  32. Katz AM and Messineo FC. Lipid-membrane interactions and the pathogenesis of ischemic damage in the myocardium. Circ Res 48: 1–16, 1981.[Free Full Text]
  33. Korge P, Honda HM, and Weiss JN. Effects of fatty acids in isolated mitochondria: implications for ischemic injury and cardioprotection. Am J Physiol Heart Circ Physiol 285: H259–H269, 2003.[Abstract/Free Full Text]
  34. Kowaltowski AJ, Cosso RG, Campos CB, and Fiskum G. Effect of Bcl-2 overexpression on mitochondrial structure and function. J Biol Chem 277: 42802–42807, 2002.[Abstract/Free Full Text]
  35. Lam KT, Borkan S, Claffey KP, Schwartz JH, Chobanian AV, and Brecher P. Properties and differential regulation of 2 fatty-acid binding-proteins in the rat-kidney. J Biol Chem 263: 15762–15768, 1988.[Abstract/Free Full Text]
  36. Lambert AJ and Brand MD. Inhibitors of the quinone-binding site allow rapid superoxide production from mitochondrial NADH: ubiquinone oxidoreductase (complex I). J Biol Chem 279: 39414–39420, 2004.[Abstract/Free Full Text]
  37. Ledenev AN, Konstantinov AA, Popova E, and Ruuge EK. A simple assay of the superoxide generation rate with tiron as an Epr-visible radical scavenger. Biochem Int 13: 391–396, 1986.[Web of Science][Medline]
  38. Lehninger AL and Remmert LF. Endogenous uncoupling and swelling agent in liver mitochondria and its enzymic formation. J Biol Chem 234: 2459–2464, 1959.[Free Full Text]
  39. Li S, Wu P, Yarlagadda P, Vadjunec NM, Proia AD, Harris RA, and Portilla D. PPAR{alpha} ligand protects during cisplatin-induced acute renal failure by preventing inhibition of renal FAO and PDC activity. Am J Physiol Renal Physiol 286: F572–F580, 2004.[Abstract/Free Full Text]
  40. Lieberthal W and Nigam SK. Acute renal failure. I. Relative importance of proximal vs. distal tubular injury. Am J Physiol Renal Physiol 275: F623–F631, 1998.[Abstract/Free Full Text]
  41. Logue SE, Gustafsson AB, Samali A, and Gottlieb RA. Ischemia/reperfusion injury at the intersection with cell death. J Mol Cell Cardiol 38: 21–33, 2005.[CrossRef][Web of Science][Medline]
  42. Matthys E, Patel Y, Kreisberg J, Stewart JH, and Venkatachalam MA. Lipid alterations induced by renal ischemia: pathogenic factor in membrane damage. Kidney Int 26: 153–161, 1984.[Web of Science][Medline]
  43. Mitchell JB, Samuni A, Krishna MC, DeGraff WG, Ahn MS, Samuni U, and Russo A. Biologically-active metal-independent superoxide-dismutase mimics. Biochemistry 29: 2802–2807, 1990.[CrossRef][Medline]
  44. Mittnacht S Jr, Sherman SC, and Farber JL. Reversal of ischemic mitochondrial dysfunction. J Biol Chem 254: 9871–9878, 1979.[Free Full Text]
  45. Mokhova EN and Khailova LS. Involvement of mitochondrial inner membrane anion carriers in the uncoupling effect of fatty acids. Biochemistry (Mosc) 70: 159–163, 2005.[CrossRef][Medline]
  46. Moran JH, Mitchell LA, and Grant DF. Linoleic acid prevents chloride influx and cellular lysis in rabbit renal proximal tubules exposed to mitochondrial toxicants. Toxicol Appl Pharmacol 176: 153–161, 2001.[CrossRef][Web of Science][Medline]
  47. Nakamura H, Nemenoff RA, Gronich JH, and Bonventre JV. Subcellular characteristics of phospholipase A2 activity in the rat kidney. Enhanced cytosolic, mitochondrial, and microsomal phospholipase A2 enzymatic activity after renal ischemia and reperfusion. J Clin Invest 87: 1810–1818, 1991.[Web of Science][Medline]
  48. Nishimura Y and Lemasters JJ. Glycine blocks opening of a death channel in cultured hepatic sinusoidal endothelial cells during chemical hypoxia. Cell Death Differ 8: 850–858, 2001.[CrossRef][Web of Science][Medline]
  49. Ouhabi R, Boue-Grabot M, and Mazat JP. Mitochondrial ATP synthesis in permeabilized cells: assessment of the ATP/O values in situ. Anal Biochem 263: 169–175, 1998.[CrossRef][Web of Science][Medline]
  50. Parce JW, Cunningham CC, and Waite M. Mitochondrial phospholipase A2 activity and mitochondrial aging. Biochemistry 17: 1634–1639, 1978.[CrossRef][Medline]
  51. Parce JW, Spach PI, and Cunningham CC. Deterioration of rat liver mitochondria under conditions of metabolite deprivation. Biochem J 188: 817–822, 1980.[Web of Science][Medline]
  52. Petitpas I, Grune T, Bhattacharya AA, and Curry S. Crystal structures of human serum albumin complexed with monounsaturated and polyunsaturated fatty acids. J Mol Biol 314: 955–960, 2001.[CrossRef][Web of Science][Medline]
  53. Polis BD and Shmukler HW. Mitochrome isolation and physicochemical properties. 2. Enzymatic effects. J Biol Chem 227: 419–440, 1957.[Free Full Text]
  54. Portilla D. Role of fatty acid beta-oxidation and calcium independent phophsolipase A2 in ischemic acute renal failure. Curr Opin Nephrol Hypertens 8: 473–477, 1999.[CrossRef][Web of Science][Medline]
  55. Portilla D. Carnitine palmitoyl-transferase enzyme inhibition protects proximal tubules during hypoxia. Kidney Int 52: 429–437, 1997.[Web of Science][Medline]
  56. Portilla D. Energy metabolism and cytotoxicity. Semin Nephrol 23: 432–438, 2003.[CrossRef][Web of Science][Medline]
  57. Portilla D and Creer MH. Plasmalogen phospholipid hydrolysis during hypoxic injury of rabbit proximal tubules. Kidney Int 47: 1087–1094, 1995.[Web of Science][Medline]
  58. Portilla D, Dai G, McClure T, Bates L, Kurten R, Megyesi J, Price P, and Li S. Alterations of PPAR{alpha} and its coactivator PGC-1 in cisplatin-induced acute renal failure. Kidney Int 62: 1208–1218, 2002.[CrossRef][Web of Science][Medline]
  59. Portilla D, Dai G, Peters JM, Gonzalez FJ, Crew MD, and Proia AD. Etomoxir-induced PPAR{alpha}-modulated enzymes protect during acute renal failure. Am J Physiol Renal Physiol 278: F667–F675, 2000.[Abstract/Free Full Text]
  60. Portilla D, Mandel LJ, Bar-Sagi D, and Millington DS. Anoxia induces phospholipase A2 activation in rabbit renal proximal tubules. Am J Physiol Renal Fluid Electrolyte Physiol 262: F354–F360, 1992.[Abstract/Free Full Text]
  61. Portilla D, Shah SV, Lehman PA, and Creer MH. Role of cytosolic calcium-independent plasmalogen-selective phospholipase A2 in hypoxic injury to rabbit proximal tubules. J Clin Invest 93: 1609–1615, 1994.[Web of Science][Medline]
  62. Pressman BC and Lardy HA. Influence of potassium and other alkali cations on respiration of mitochondria. J Biol Chem 197: 547–556, 1952.[Free Full Text]
  63. Pressman BC and Lardy HA. Effect of surface active agents on the latent ATPase of mitochondria. Biochim Biophys Acta 21: 458–466, 1956.[Medline]
  64. Pullman ME and Racker E. Spectrophotometric studies of oxidative phosphorylation. Science 123: 1105–1107, 1956.[Free Full Text]
  65. Richieri GV and Kleinfeld AM. Unbound free fatty-acid levels in human serum. J Lipid Res 36: 229–240, 1995.[Abstract]
  66. Rizzuto R, Bernardi P, and Pozzan T. Mitochondria as all around players in the calcium game. J Physiol 529: 37–47, 2000.[Abstract/Free Full Text]
  67. Schonefeld M, Noble S, Bertorello AM, Mandel LJ, Creer MH, and Portilla D. Hypoxia-induced amphiphiles inhibit renal Na+,K+-ATPase. Kidney Int 49: 1289–1296, 1996.[Web of Science][Medline]
  68. Scorrano L, Penzo D, Petronilli V, Pagano F, and Bernardi P. Arachidonic acid causes cell death through the mitochondrial permeability transition. Implications for tumor necrosis factor-{alpha} apoptotic signaling. J Biol Chem 276: 12035–12040, 2001.[Abstract/Free Full Text]
  69. Shabalina IG, Jacobsson A, Cannon B, and Nedergaard J. Native UCP1 displays simple competitive kinetics between the regulators purine nucleotides and fatty acids. J Biol Chem 279: 38236–38248, 2004.[Abstract/Free Full Text]
  70. Sheridan AM, Schwartz JH, Kroshian VM, Tercyak AM, Laraia J, Masino S, and Lieberthal W. Renal mouse proximal tubular cells are more susceptible than MDCK cells to chemical anoxia. Am J Physiol Renal Fluid Electrolyte Physiol 265: F342–F350, 1993.[Abstract/Free Full Text]
  71. Shrago E, Shug A, Elson C, Spennett T, and Crosby C. Regulation of metabolite transport in rat and guinea-pig liver-mitochondria by long-chain fatty acyl coenzyme-A esters. J Biol Chem 249: 5269–5274, 1974.[Abstract/Free Full Text]
  72. Skulachev VP. Uncoupling: new approaches to an old problem of bioenergetics. Biochim Biophys Acta 1363: 100–124, 1998.[Medline]
  73. Spector AA. Fatty-acid binding to plasma albumin. J Lipid Res 16: 165–179, 1975.[Abstract]
  74. Spector AA, John K, and Fletcher JE. Binding of long-chain fatty acids to bovine serum albumin. J Lipid Res 10: 56–67, 1969.[Abstract]
  75. Vandervusse GJ, Glatz JFC, Stam HCG, and Reneman RS. Fatty-acid homeostasis in the normoxic and ischemic jeart. Physiol Rev 72: 881–940, 1992.[Free Full Text]
  76. Venkatachalam MA, Patel YJ, Kreisberg JI, and Weinberg JM. Energy thresholds that determine membrane integrity and injury in a renal epithelial cell line (LLC-PK1). Relationships to phospholipid degradation and unesterified fatty acid accumulation. J Clin Invest 81: 745–758, 1988.[Web of Science][Medline]
  77. Vogt MT and Farber E. On the molecular pathology of ischemic renal cell death. Am J Pathol 53: 1–26, 1968.[Web of Science][Medline]
  78. Waite M, Scherphof GL, Boshouwers FM, and Van Deenen LL. Differentiation of phospholipases A in mitochondria and lysosomes of rat liver. J Lipid Res 10: 411–420, 1969.[Abstract]
  79. Weinberg JM, Davis JA, Abarzua M, and Rajan T. Cytoprotective effects of glycine and glutathione against hypoxic injury to renal tubules. J Clin Invest 80: 1446–1454, 1987.[Web of Science][Medline]
  80. Weinberg JM, Nissim I, Roeser NF, Davis JA, Schultz S, and Nissim I. Relationships between intracellular amino acid levels and protection against injury to isolated proximal tubules. Am J Physiol Renal Fluid Electrolyte Physiol 260: F410–F419, 1991.[Abstract/Free Full Text]
  81. Weinberg JM, Roeser NF, Davis JA, and Venkatachalam MA. Glycine-protected, hypoxic, proximal tubules develop severely compromised energetic function. Kidney Int 52: 140–151, 1997.[Web of Science][Medline]
  82. Weinberg JM, Venkatachalam MA, Goldberg H, Roeser NF, and Davis JA. Modulation by Gly, Ca, and acidosis of injury-associated unesterified fatty acid accumulation in proximal tubule cells. Am J Physiol Renal Fluid Electrolyte Physiol 268: F110–F121, 1995.[Abstract/Free Full Text]
  83. Weinberg JM, Venkatachalam MA, Roeser NF, and Nissim I. Mitochondrial dysfunction during hypoxia/reoxygenation and its correction by anaerobic metabolism of citric acid cycle intermediates. Proc Natl Acad Sci USA 97: 2826–2831, 2000.[Abstract/Free Full Text]
  84. Weinberg JM, Venkatachalam MA, Roeser NF, Saikumar P, Dong Z, Senter RA, and Nissim I. Anaerobic and aerobic pathways for salvage of proximal tubules from hypoxia-induced mitochondrial injury. Am J Physiol Renal Physiol 279: F927–F943, 2000.[Abstract/Free Full Text]
  85. Weinberg JM, Venkatachalam MA, Roeser NF, Senter RA, and Nissim I. Energetic determinants of tyrosine phosphorylation of proximal tubule focal adhesion proteins. Am J Pathol 158: 2153–2164, 2001.[Abstract/Free Full Text]
  86. Wetzels JFM, Wang X, Gengaro PE, Nemenoff RA, Burke TJ, and Schrier RW. Glycine protection against hypoxic but not phospholipase A2-induced injury in rat proximal tubules. Am J Physiol Renal Fluid Electrolyte Physiol 264: F94–F99, 1993.[Abstract/Free Full Text]
  87. Wieckowski MR and Wojtczak L. Fatty acid-induced uncoupling of oxidative phosphorylation is partly due to opening of the mitochondrial permeability transition pore. FEBS Lett 423: 339–342, 1998.[CrossRef][Web of Science][Medline]
  88. Wight JP, Chilcott JB, Holmes MW, and Brewer N. Pulsatile machine perfusion vs. cold storage of kidneys for transplantation: a rapid and systematic review. Clin Transplant 17: 293–307, 2003.[CrossRef][Web of Science][Medline]
  89. Wilson DR, Arnold PE, Burke TJ, and Schrier RW. Mitochondrial calcium accumulation and respiration in ischemic acute renal failure in the rat. Kidney Int 25: 519–526, 1984.[Web of Science][Medline]
  90. Wojtczak L. Effect of long-chain fatty acids and acyl-CoA on mitochondrial permeability, transport, and energy-coupling processes. J Bioenerg Biomembr 8: 293–311, 1976.[Web of Science][Medline]
  91. Wojtczak L and Lehninger AL. Formation and disappearance of an endogenous uncoupling factor during swelling and contraction of mitochondria. Biochim Biophys Acta 51: 442–448, 1961.[Medline]
  92. Wojtczak L and Schonfeld P. Effect of fatty acids on energy coupling processes in mitochondria. Biochim Biophys Acta 1183: 41–57, 1993.[Medline]
  93. Wojtczak L and Wieckowski MR. The mechanisms of fatty acid-induced proton permeability of the inner mitochondrial membrane. J Bioenerg Biomembr 31: 447–455, 1999.[CrossRef][Web of Science][Medline]
  94. Wojtczak L and Zaluska H. Inhibition of translocation of adenine nucleotides through mitochondrial membranes by oleate. Biochem Biophys Res Commun 28: 76–81, 1967.[CrossRef][Web of Science][Medline]
  95. Zager RA, Burkhart KM, Conrad DS, Gmur DJ, and Iwata M. Phospholipase A2-induced cytoprotection of proximal tubules: potential determinants and specificity for ATP depletion-mediated injury. J Am Soc Nephrol 7: 64–72, 1996.[Abstract]
  96. Zager RA, Conrad DS, and Burkhart K. Phospholipase A2: a potentially important determinant of adenosine triphosphate levels during hypoxic-reoxygenation tubular injury. J Am Soc Nephrol 7: 2327–2339, 1996.[Abstract]
  97. Zager RA, Gmur DJ, Bredl CR, and Eng MJ. Temperature effects on ischemic and hypoxic renal proximal tubular injury. Lab Invest 64: 766–776, 1991.[Web of Science][Medline]
  98. Zoratti M, Szabo D, and De Marchi U. Mitochondrial permeability transitions: how many doors to the house? Biochim Biophys Acta Bioenerget 1706: 40–52, 2005.[CrossRef]



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