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NDT Advance Access published online on August 1, 2008
Nephrology Dialysis Transplantation, doi:10.1093/ndt/gfn436
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© The Author [2008]. Published by Oxford University Press on behalf of ERA-EDTA. All rights reserved. For Permissions, please e-mail: journals.permissions@oxfordjournals.org


Evidence for involvement of nonesterified fatty acid-induced protonophoric uncoupling during mitochondrial dysfunction caused by hypoxia and reoxygenation

Thorsten Feldkamp1, Joel M. Weinberg2, Markus Hörbelt1, Christina Von Kropff1, Oliver Witzke1, Jens Nürnberger1 and Andreas Kribben1

1 Department of Nephrology, University Hospital Essen, University Duisburg-Essen, 45122 Essen, Germany 2 Division of Nephrology, Department of Internal Medicine, Veterans Affairs Ann Arbor Healthcare System and University of Michigan, Ann Arbor, MI 48109, USA

Correspondence and offprint requests to: Thorsten Feldkamp, Department of Nephrology, University Hospital Essen, University of Duisburg-Essen, Hufelandstr. 55, 45122 Essen, Germany. Tel: +49-201-723-2552; Fax: +49-201-723-5633; E-mail: thorsten.feldkamp{at}uni-due.de



   Abstract
Top
 Abstract
Introduction
 Subjects and methods
 Results
 Discussion
 References
 
Background. Proximal tubules subjected to hypoxia in vitro under conditions relevant to ischaemia in vivo develop an energetic deficit that is not corrected even after full reoxygenation. We have provided evidence that accumulation of nonesterified fatty acids (NEFA) is the primary reason for this energetic deficit. In this study, we have further investigated the mechanism for the NEFA-induced energetic deficit.

Methods. Mitochondrial membrane potential ({Delta}{psi}) was measured in digitonin-permeabilized, freshly isolated proximal tubules by safranin O uptake. Addition of the potassium/proton exchanger nigericin enables the determination of the mitochondrial proton motive force ({Delta}p) and the proton gradient ({Delta}pH). ATP was measured luminometrically and NEFA colorimetrically.

Results. Tubule ATP content was depleted after hypoxia and recovered incompletely, even after full reoxygenation. Mitochondrial safranin O uptake was decreased in proximal tubules after hypoxia and reoxygenation (H/R). This decrease was attenuated by delipidated bovine serum albumin (dBSA) or citrate. Addition of nigericin increased safranin O uptake of mitochondria in normoxic proximal tubules, but not in proximal tubules after H/R. Addition of dBSA restored the effect of nigericin to increase mitochondrial safranin O uptake. Addition of the NEFA oleate had the same impact on mitochondrial safranin O uptake as subjecting proximal tubules to H/R.

Conclusion. The mechanism of the NEFA-induced energetic deficit in freshly isolated rat proximal tubules induced by H/R is characterized by impaired ATP production after full reoxygenation, impaired recovery of {Delta}{psi} and {Delta}p, abrogation of {Delta}pH and sensitivity to citrate, consistent with involvement of the tricarboxylate carrier. The data support the concept that protonophoric uncoupling by NEFA movement on anion carriers plays a critical role in proximal tubule mitochochondrial dysfunction after H/R.

Keywords: acute kidney injury; hypoxia/reoxygenation; mitochondrial damage; nonesterified fatty acids; proximal tubule



   Introduction
Top
 Abstract
 Introduction
Subjects and methods
 Results
 Discussion
 References
 
Proximal tubule injury plays a critical role in the pathogenesis of acute kidney injury (AKI), which is most commonly caused by renal ischaemia [1]. Freshly isolated proximal tubules subjected to hypoxia in vitro under conditions relevant to ischaemia in vivo develop a mitochondrial energetic deficit that is not corrected even after full reoxygenation [2,3]. Because fully differentiated kidney proximal tubules have minimal glycolytic capacity, they are dependent on mitochondrial metabolism for ATP synthesis [4]. The limitation of the restoration of cellular ATP during reoxygenation plays a pivotal role in overall cellular recovery [5]. Nonesterified fatty acids (NEFA) progressively accumulate during renal ischaemia in vivo [6] and during hypoxia of isolated proximal tubules in vitro [7,8]. We have provided evidence that accumulation of NEFA is the primary reason for mitochondrial dysfunction and for the energetic deficit that develops in proximal tubules during hypoxia and reoxygenation (H/R) [9]. To further study the mechanism of NEFA-induced mitochondrial dsyfunction, we analysed the impact of H/R and NEFA on the mitochondrial driving force for ATP production and the involvement of integral proteins of the inner mitochondrial membrane in this mechanism.

NEFA limit mitochondrial ATP production by decreasing mitochondrial membrane potential ({Delta}{psi}) [9]. However, mitochondrial {Delta}{psi} is only one part of the driving force for ATP production over the inner mitochondrial membrane; the other part is the mitochondrial proton gradient ({Delta}pH). Taken together, {Delta}{psi} and {Delta}pH are termed proton motive force ({Delta}p), which is the complete driving force responsible for mitochondrial ATP production [10]. Usually, {Delta}pH is only a small part of {Delta}p. Therefore, measurement of {Delta}{psi} should be sufficient to assess the mitochondrial capability to produce ATP. However, it is conceivable that the relative contributions of {Delta}pH and {Delta}{psi} to {Delta}p change after H/R or in the presence of NEFA. The inner mitochondrial membrane has ATP-dependent potassium channels, which are opened during ATP depletion after hypoxia [11]. An increase in potassium permeability would decrease {Delta}{psi} and concomitantly increase {Delta}pH, leaving {Delta}p unchanged [10]. If net {Delta}p remained unchanged, it could not account for the energetic deficit that develops during H/R. To confirm that it is sufficient to measure {Delta}{psi} after H/R and to study the effect of hypoxia and NEFA on mitochondrial ATP production in more depth, we measured mitochondrial {Delta}p after H/R and in the presence of exogenous NEFA. The measurement of {Delta}p then allows us to determine {Delta}pH by calculating the delta of {Delta}p and {Delta}{psi}. We have recently optimized approaches for quantitative and dynamic measurements of mitochondrial {Delta}{psi} in permeabilized isolated tubules using safranin O [12]. In this study, we measure mitochondrial {Delta}p using safranin O, and show that {Delta}{psi} is the predominant component of {Delta}p even after H/R or in the presence of NEFA and that H/R and NEFA affect {Delta}p and {Delta}pH in the same way. This confirms that the mechanism of mitochondrial perturbation induced by H/R and NEFA is the same, which adds to the evidence that NEFA accumulation is indeed the primary cause for the energetic deficit after H/R.

It has been shown that NEFA can decrease {Delta}{psi} by acting 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 integral proteins (e.g. anion carriers) in the inner mitochondrial membrane [13,14]. We have provided evidence that the glutamate aspartate carrier and the adenine nucleotide carrier are involved in the NEFA-induced decrease in {Delta}{psi} [15]. In this study, we investigated whether other carriers, e.g. the tricarboxylate carrier, are involved in this process. The tricarboxylate carrier is located in the inner mitochondrial membraneand is involved in the mitochondrial exchange of a tricarboxylate (e.g. citrate) for another tricarboxylate, dicarboxylate or phosphoenolpyruvate (PEP) [16]. To address whether the tricarboxylate carrier could be involved in the NEFA-induced decrease in {Delta}{psi}, we assessed modification of the process by citrate using both normoxic tubules treated with exogenous NEFA and tubules de-energized as a result of H/R. Our data further support a major role for shuttling of NEFA by mitochondrial anion carriers in the pathogenesis of the important NEFA-induced mitochondrial dysfunction in proximal tubules after H/R.

Hitherto, the development of the energetic deficit during H/R has been demonstrated in freshly isolated rabbit proximal tubules [2,3,9]. We wanted to assess whether the development of the energetic deficit after H/R is a fundamental cellular response to H/R in general that develops in other species as well. We show here that freshly isolated rat proximal tubules develop the same energetic deficit as rabbit tubules after H/R and that NEFA also play a critical role in the development of this energetic deficit in rat proximal tubules.



   Subjects and methods
Top
 Abstract
 Introduction
 Subjects and methods
Results
 Discussion
 References
 
Materials
All compounds were of the highest available purity and obtained from Sigma (Taufkirchen, Germany) if not otherwise indicated.

Animals
Male Sprague-Dawley rats (Charles River, Sulzfeld, Germany) weighing 240–320 g were used for these studies. Animal use protocols for the studies in this manuscript adhered to our animal law and were approved by the Institutional Animal Care and Use Committee of the University of Duisburg-Essen.

Preparation of isolated proximal tubules
Renal proximal tubules were freshly isolated as previously described, with slight modification [17,18]. Briefly, rats were anaesthetized with xylazin (6 mg/kg bodyweight i.p.) and ketamine (120 mg/kg bodyweight i.p.), and the kidneys flushed with 40 ml of oxygenated solution A containing (in mM) NaCl, 112; NaHCO3, 20; KCl, 5; CaCl2, 1.6; Na2HPO4, 2; MgSO4, 1.2; glucose, 5; HEPES, 10; mannitol, 10; glutamine, 1; sodium butyrate, 1; sodium lactate, 1; pH 7.05, 4°C with the addition of 2000 I.E. heparin. Subsequently, perfusion was continued with 30 ml of oxygenated solution A containing 5 mg of collagenase [type A, specific activity (SA): 0.228 U/mg; Lot 70128622, Roche Diagnostics GmbH, Mannheim, Germany] and 12.5 mg of hyaluronidase (Roche Diagnostics GmbH Mannheim, Germany). After perfusion, the kidneys were decapsulated, removed and transferred to ice-cold solution A. The renal cortices were dissected and minced on a cold petri dish. After two washes with solution A, the tissue was incubated for 30 min in 30 ml of oxygenated solution A containing 10 mg collagenase (SA as described above) and 7.5 mg of hyaluronidase. Separation of the tubules was checked regularly under the microscope. When the tubules were separated, they were washed and placed in 15 ml of ice-cold solution A containing 0.5 g of bovine serum albumin (Serva, Heidelberg, Germany) for 20 min. After filtering the tissue and two washes to remove albumin, the tubules were separated using 45% Percoll (Pharmacia, Uppsala, Sweden). After centrifugation for 10 min at 11 000 x g, the proximal tubules were recovered from the lowest band. This band was composed primarily (>95%) of proximal tubules and contained no glomeruli. This suspension was washed three times to remove the Percoll.

Experimental procedure
Tubules were resuspended at 1.0 mg tubule protein/ml in a 95% air/5% CO2-gassed medium containing (in mM) NaCl, 106; NaHCO3, 30; KCl, 5; CaCl2, 1; Na2HPO4, 2; MgSO4, 1; glucose, 5; HEPES, 10; sodium butyrate, 10; alanine, 1; sodium lactate, 4; glycine, 2 and pH 7.05, 4°C, and gassed on ice for 5 min with 95% O2/5% CO2 (solution B). Thereafter, the flask was capped with a rubber stopper and the tubule suspension was allowed to warm up for 10 min at room temperature. After equilibration at room temperature, tubules were placed in a shaking water bath at 37°C for 10 min, whereafter pH had increased to 7.35. After the equilibration period at 37°C, samples were removed for analysis (t = 0 min) and tubules were resuspended in fresh solution B and regassed with either 95% air/5% CO2 (normoxic controls) or 95% N2/5% CO2 (hypoxia). During hypoxia, solution B was kept at pH 6.9 to simulate tissue acidosis during ischaemia in vivo [19], and omitted glucose, butyrate, alanine and lactate. These incubation conditions result in near anoxic conditions. As it is not possible to readily confirm the presence of complete anoxia in the flasks, we use the term hypoxia to describe the condition of oxygen deprivation. After 60 min, samples were again removed for analysis. The remaining tubules were pelleted and then resuspended in fresh 95% air/5% CO2-gassed, pH 7.4 solution B. Sodium butyrate in solution B was replaced with 2 mM heptanoic acid during reoxygenation, and, to assure availability of purine precursors for ATP resynthesis, 250 µM AMP was included [2,19]. After 10 min and 60 min of reoxygenation, samples were removed again for analysis.

ATP content
ATP content of proximal tubules was measured by a luciferase assay [20] using the ATP-Bioluminescence Assay Kit (Roche Diagnostics GmbH, Mannheim, Germany) with the method suggested by the manufacturer. At the given time points 0.5 ml medium was taken, added to 50 µl of 10 M perchloric acid, vigorously mixed and immediately frozen at –80°C. After thawing, samples were diluted 1:10 000 in buffer containing 100 mM Tris and 4 mM EDTA, pH 7.75 and mixed immediately with equal amounts of luciferase reagent. Emitted light of the luciferase was measured in a luminometer (Berthold Detection Systems, Pforzheim, Germany).

LDH release
LDH release was measured in a 0.5 ml aliquot of the tubule suspension at the given time points. Immediate centrifugation at 3000 x g for 1 min and lysis of the pellet permitted the measurement of LDH activity separately in the supernate and in the pellet as previously described [17,21]. LDH activity was measured photometrically and calculated by dividing LDH activity in the supernatant by total LDH activity (supernatant + pellet). The results are expressed as a percentage of total and supernatant LDH [22].

Measurement of NEFA
Lipids were extracted as previously described [6,9] by Bligh and Dyer [23] 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 was 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. 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 GmbH, Neuss, Germany).

Mitochondrial membrane potential ({Delta}{psi}), proton gradient ({Delta}pH) and proton motive force ({Delta}p)
At the end of the experimental procedure, tubules were pelleted, washed once in an ice-cold solution containing (in mM) 110 NaCl, 25 NaHEPES, pH 7.2, 1.25 CaCl2, 1.0 MgCl2, 1.0 KH2PO4, 3.5 KCl, 5.0 glycine and 5% polyethylene glycol (average MW 8000), and held in it at 4°C until use. For the safranin O uptake measurements, the tubules were resuspended at a final concentration of 0.05–0.1 mg/ml in an intracellular buffer-type solution containing 120 mM KCl, 1 mM KH2PO4, 2 mM EGTA, 5 µM safranin O, 200 µg digitonin/mg protein (Calbiochem, Catalogue No. 300411), 10 mM K-HEPES, pH 7.2 at 37°C (solution C) supplemented as needed for specific experiments with succinate (4 mM) and other experimental reagents that are described with the data [12,24]. Succinate was used as the substrate during safranin O uptake because it strongly supports energization of mitochondria in both permeabilized normoxic control tubules and after H/R [12]. For the addition of exogenous NEFA, oleate, in a concentration of 1 µM was used. Fluorescence was followed at 490 nm excitation, 581 nm emission using a F-2500 Fluorescence Spectrophotometer (Hitachi High-Technologies Corporation, Tokyo, Japan), equipped with temperature controlled and magnetically stirred cuvette holders, as illustrated by the tracings shown in the Results section. Uptake of safranin O into 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}, they are inverted in the figures shown. 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 prior to 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. If changes in fluorescence between different groups were compared, fluorescence changes were factored by protein [9,15].

For the measurement of {Delta}p, 25 nM nigericin was added after the measurement of {Delta}{psi}. Nigericin is an ionophore that exchanges potassium ions for protons across the mitochondrial inner membrane. In the presence of nigericin, if [K+]in = [K+]out, then [H+]in = [H+]out and {Delta}pH = 0. The concentration in the buffer was 120 mM, which approximates to [K+]in. Since {Delta}pH is abolished by nigericin in the medium, the electron transport chain compensates for the drop in {Delta}pH by pumping more protons, so that {Delta}{psi} increases and {Delta}pH is quickly converted entirely into {Delta}{psi}. If {Delta}pH is zero, {Delta}{psi} is equal to {Delta}p, thus allowing us to determine {Delta}p by the measurement of {Delta}{psi} [25]. {Delta}pH can then be determined by the delta of {Delta}p and {Delta}{psi}, since the summation of {Delta}{psi} and {Delta}pH is {Delta}p [10].

Statistics
Paired and unpaired t-tests were used as appropriate. Where experiments consisted of multiple groups, they were analysed statistically by analysis of variance 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, USA). P < 0.05 was considered to be statistically significant. Data shown are either mean ± SE of no less than 3–5 experiments or are tracings representative of the behaviour in that many experiments.



   Results
Top
 Abstract
 Introduction
 Subjects and methods
 Results
Discussion
 References
 
ATP content, LDH release and NEFA content of proximal tubules during H/R
Tubules were studied after 60 min of hypoxia followed by 10 and 60 min of reoxygenation. Tubule ATP content decreased from 9.54 ± 0.88 to 0.63 ± 0.07 nmol/mg protein after 60 min of hypoxia without extra substrates (Figure 1). Tubule ATP content recovered to only 1.30 ± 0.11 nmol/mg protein after 10 min of reoxygenation and to 2.20 ± 0.29 nmol/mg protein after 60 min of reoxygenation (Figure 1). During the first 60 min of normoxia, tubule ATP content remained constant and increased to 12.90 ± 1.00 during the second 60 min period, when tubules were supplemented with AMP (Figure 1). Tubule LDH release after hypoxia and 60 min of reoxygenation was not increased compared to normoxia (10.7% versus 11.1, n = 6, n.s.). Proximal tubules sampled after hypoxia and 60 min of reoxygenation had much higher NEFA content compared to proximal tubules studied under normoxic conditions (7.0 ± 0.5 versus 2.7 ± 0.3 nmol/mg protein, n = 6, P < 0.01).


Figure 1
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  		Fig. 1 Tubule ATP content after 60 min of hypoxia and different duration of reoxygenation. Tubules were subjected to 60 min of hypoxia and to 10 (for a total duration of 70 min) or 60 min (for a total duration of 120 min) of reoxygenation (H/R). Control tubules were subjected to the corresponding duration of normoxia. Values are means ± SEM, n = 12, *P < 0.01 versus normoxia.

 
Mitochondrial membrane potential ({Delta}{psi}) of proximal tubule mitochondria after H/R with and without addition of dBSA
Mitochondrial safranin O uptake by normoxic tubules with addition of delipidated bovine serum albumin (dBSA) was set as 100%. Albumin was used to bind NEFA and thereby prevented the interaction of NEFA with the mitochondria. Normoxic tubules without addition of dBSA had a mitochondrial safranin O uptake of 62.3 ± 2.5%. Safranin O uptake in tubules subjected to 60 min of hypoxia and 10 min of reoxygenation was decreased to 3.7 ± 2.1% (P < 0.01 versus normoxic control, n = 6, Figure 2A and B). Addition of dBSA to tubules subjected to 60 min of hypoxia and 10 min of reoxygenation resulted in an increase of safranin O uptake to 62.6 + 7.2% (P < 0.01 versus H/R without dBSA, n = 6, Figure 2A and B).


Figure 2
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  		Fig. 2 Mitochondrial membrane potential ({Delta}{psi}) after hypoxia and reoxygenation (H/R). Left panel: representative fluorescence tracings of safranin O uptake by mitochondria from digitonin-permeabilized proximal tubules respiring on succinate. Mitochondrial safranin O uptake was measured after proximal tubules had been subjected to 60 min of hypoxia and 10 min of reoxygenation (H/R) or to the corresponding duration of normoxia. When indicated, delipidated bovine serum albumin (dBSA) were present during measurement of safranin O uptake. Relative fluorescence was calculated by dividing fluorescence by the fluorescence after initial equilibration. Right panel: group data for net safranin O uptake during studies under the same conditions as in the left panel. Data are safranin O uptakes calculated as the difference in fluorescence at the beginning and the point of maximal change in fluorescence as a percentage of maximum safranin O uptake by normoxic mitochondria with dBSA present during the measurement. Values are means ± SEM of six experiments. *P < 0.01 versus normoxia without dBSA (Nor), #< 0.01 versus H/R without dBSA.

 
Proton motive force ({Delta}p) and mitochondrial membrane potential ({Delta}{psi}) in proximal tubule mitochondria after H/R and in the presence of the NEFA, oleate
Addition of nigericin (25 nM), a hydrogen/potassium exchanger, to digitonin-permeabilized proximal tubules converts {Delta}pH to {Delta}{psi}, as illustrated in Figure 3. If {Delta}pH is zero, {Delta}{psi} and {Delta}p are equal (Figure 3) [25]. According to the formula {Delta}p = {Delta}{psi} + {Delta}pH, the delta between {Delta}p and {Delta}{psi} is {Delta}pH [10]. Addition of dBSA during the measurement of mitochondrial safranin O uptake in normoxic control tubules resulted in an increase of {Delta}{psi} and {Delta}p (P < 0.05, n = 5–11, Figure 4). Mitochondrial safranin O uptake in proximal tubules after H/R was decreased with and without nigericin (P < 0.05, n = 5–11, Figure 4), which indicates that {Delta}{psi} and {Delta}p are both affected in the same way. Addition of dBSA during safranin O uptake increased both {Delta}{psi} and {Delta}p (P < 0.05, n = 5–11, Figure 4). The NEFA oleate (1 µM) decreased {Delta}{psi} and {Delta}p in the same way, and this effect was reversed by binding of oleate with dBSA (P < 0.05, n = 5–11, Figure 4).


Figure 3
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  		Fig. 3 Illustration of the conversion of the mitochondrial proton gradient ({Delta}pH) to the mitochondrial membrane potential ({Delta}{psi}) for determination of the mitochondrial protonmotive force ({Delta}p). Representative fluorescence tracing for the illustration of the measurement of {Delta}p by mitochondrial safranin O uptake. After allowing maximal uptake of safranin O, addition of nigericin increased safranin O uptake by conversion of {Delta}pH to {Delta}{psi}. In the presence of nigercin {Delta}pH is zero, making {Delta}{psi} the only component of {Delta}p, thus {Delta}{psi} equals {Delta}p under this condition. {Delta}pH can be determined by the delta of {Delta}{psi} before and after nigericin addition (see the Subjects and method section).

 

Figure 4
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  		Fig. 4 Mitochondrial membrane potential ({Delta}{psi}) and protonmotive force ({Delta}p) after hypoxia and reoxygenatio (H/R) and in the presence of nonesterified fatty acids (NEFA). Net safranin O uptake of mitochondria from digitonin-permeabilized proximal tubules respiring on succinate are measured under the following conditions: safranin O uptake after proximal tubules had been subjected to 60 min of hypoxia and 10 min of reoxygenation (H/R) or the corresponding duration of normoxia (Nor). Delipidated bovine serum albumin (dBSA) or 1 µM of the NEFA oleate (Ol) were present during measurement of safranin O uptake where indicated. Safranin O uptakes were measured as in Figure 2. For the measurement of {Delta}p, 25 nM nigericin was added as illustrated in Figure 3. Values are means ± SEM of 5–11 experiments. *P < 0.01 versus respective normoxia group without further addition (Nor), #<0.01 versus respective H/R group without dBSA (H/R), +<0.01 versus respective normoxia group in the presence of oleate without dBSA (Nor Ol).

 
Citrate ameliorates NEFA-induced decrease of mitochondrial membrane potential ({Delta}{psi})
Oleate decreased mitochondrial safranin O uptake from 67.9 ± 3.2% of maximal uptake in the presence of dBSA to 42.5 ± 1.2% (Figure 5, P < 0.01, n = 15). Including dBSA with oleate prevented the oleate-induced decrease of safranin O uptake (91.3 ± 4.4% of maximal uptake, n = 15, P < 0.01 versus oleate without dBSA, Figure 5). Citrate ameliorated the oleate-induced decrease of mitochondrial safranin O uptake (62.5 ± 2.0% of maximal uptake, P < 0.01 versus oleate without citrate, n = 9, Figure 5).


Figure 5
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  		Fig. 5 Mitochondrial membrane potential ({Delta}{psi}) during addition of oleate. Left panel: representative fluorescence tracings of safranin O uptake by mitochondria from digitonin-permeabilized normoxic proximal tubules respiring on succinate in the presence of oleate (1 µM) during safranin O measurement. When indicated, citrate or delipidated bovine serum albumin (dBSA) were present during measurement of mitochondrial safranin O uptake. Right panel: group data for net safranin O uptake during studies under the same conditions as in the left panel. Safranin O uptakes were measured as in Figure 2. Values are means ± SEM of nine experiments. *P < 0.01 versus no further addition (NFA).

 
Citrate ameliorates H/R-induced decrease of mitochondrial membrane potential ({Delta}{psi})
H/R decreased mitochondrial safranin O uptake to 4.6 ± 1.0% of maximal uptake (P < 0.01 versus normoxic tubules in the absence of dBSA, n = 12, Figure 6). dBSA substantially reversed the decrease of safranin O uptake induced by H/R (62.0 ± 2.2% of maximal uptake, Figure 6, n = 12, P < 0.01 versus H/R without BSA). Citrate ameliorated the decrease of mitochondrial safranin O uptake induced by H/R (16.1 ± 1.3%, n = 12, P < 0.01 versus H/R without citrate, Figure 6).


Figure 6
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  		Fig. 6 Mitochondrial membrane potential ({Delta}{psi}) after hypoxia and reoxygenation (H/R). Left panel: representative fluorescence tracings of safranin O uptake by mitochondria from digitonin-permeabilized proximal tubules respiring on succinate. Safranin O uptake was measured after proximal tubules had been subjected to 60 min of hypoxia and 10 min of reoxygenation (H/R). When indicated, citrate or delipidated bovine serum albumine (dBSA) were present during measurement of safranin O uptake. Right panel: group data for net safranin O uptake during studies under the same conditions as in the left panel. Safranin O uptakes were measured as in Figure 2. Values are means ± SEM of 12 experiments. *P < 0.01 versus no further addition (NFA).

 


   Discussion
Top
 Abstract
 Introduction
 Subjects and methods
 Results
 Discussion
References
 
Isolated proximal tubules subjected to H/R under conditions relevant to ischaemia-reperfusion in vivo develop a persistent mitochondrial energetic deficit despite an intact plasma membrane (Figure 1) [3,19]. This energetic deficit results in a greatly decreased ATP production even after full reoxygenation (Figure 1) and plays a pivotal role in overall cellular recovery [5,19]. The energetic deficit is characterized by preserved function of the electron transport chain [26], absence of cytochrome c release [3], preserved activity of the mitochondrial F1Fo-ATPase and ADP/ATP carrier [24], and partial, but incomplete recovery of mitochondrial {Delta}{psi} [2,3,12]. We have shown that this energetic deficit is primarily caused by the accumulation of NEFA [9]. This has previously been studied in rabbit proximal tubules [2,3,5,7,9,12,15,19,24,26]. In the present study, we have investigated additional important aspects of the mechanism of the NEFA-induced energetic deficit and the resulting ATP production after H/R using ratproximal tubules.

Previous work [9,15,27] assessing NEFA effects has focused on changes of {Delta}{psi}, but {Delta}{psi} is only one part of the driving force for ATP production, which is termed proton motive force ({Delta}p) [10]. The proton motive force has two components {Delta}{psi} and {Delta}pH, which add up to {Delta}p [10]. Under physiological conditions, {Delta}pH is only a small part of {Delta}p [25], so measurement of {Delta}{psi} should be sufficient to determine mitochondrial ATP production. However, it is not known whether {Delta}{psi} is still the predominant component of {Delta}p during mitochondrial injury. It is conceivable that during reoxygenation the proportion of {Delta}{psi} and {Delta}pH could change, for instance by opening of ATP-dependent potassium channels. These potassium channels exist in the inner mitochondrial membrane and could be opened after hypoxia and during reoxygenation, when cellular ATP levels are low [11]. This would increase potassium permeability in the inner mitochondrial membrane, resulting in a decrease of mitochondrial {Delta}{psi}, an increase of mitochondrial {Delta}pH, leaving mitochondrial {Delta}p unchanged [10]. Under these circumstances, {Delta}{psi} would not be sufficient to determine the ability of mitochondria to produce ATP, if mitochondrial ATP production is not measured directly. The present studies clearly show that mitochondrial {Delta}pH is abrogated after H/R, making mitochondrial {Delta}{psi} the significant portion of mitochondrial {Delta}p. Thus, it is sufficient to measure {Delta}{psi} to interpret the ability of mitochondria to produce ATP during mitochondrial damage induced by H/R.

The alteration of lipid metabolism that occurs after renal injury was first described in 1983, when Tannenbaum and colleagues reported accumulation of NEFA after ureteral obstruction [28]. Since then numerous studies have been shown that hypoxia induced NEFA accumulation in renal tissue and proximal tubules (for a review, see [9]). In our study, NEFA were significantly increased in proximal tubules subjected to H/R compared to proximal tubules subjected to normoxia and this increase was well in the range of what others have reported and what we have reported in the rabbit tubules [9]. The mechanism of hypoxia-induced NEFA accumulation is complex. Probably, a combination of different pathways is involved: among them are increase of fatty acid transport, inhibition of mitochondrial and peroxisomal β-oxidation and phospholipid hydrolysis mediated by normal and increased phospholipase activity unopposed by ATP-requiring re-esterification [29]. The resulting impairment of mitochondrial energy metabolism by NEFA is a well-known phenomenon and has been recognized for over 50 years now [30]. In the present study, NEFA impaired mitochondrial ATP production by decreasing mitochondrial {Delta}{psi} and abrogating mitochondrial {Delta}pH, which is in the line with the favoured concept for the mechanism of NEFA-induced mitochondrial uncoupling [13]. According to this concept, the capability of NEFA to decrease ATP production 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. This is followed by dissociation of the proton and transport of the deprotonated fatty acid ion out of the matrix by different anion carriers (Figure 7) [13]. If NEFA act primarily by increasing the permeability for protons (protonophoric effect), {Delta}pH should be abrogated. This is indeed shown in this study, when the exogenous NEFA oleate added during the measurement of safranin O uptake in mitochondria from proximal tubules, reduced {Delta}pH to zero (Figure 4). The abrogation of {Delta}pH is also detected in mitochondria from permeabilized proximal tubules subjected to H/R (Figure 4). This supports the hypothesis that NEFA are indeed responsible for the mitochondrial de-energization seen after H/R. We have shown that addition of exogenous NEFA and subjecting proximal tubules to H/R have the same impact on mitochondrial {Delta}{psi}, {Delta}pH and {Delta}p. This strengthens our hypothesis that NEFA-induced protonophoric uncoupling is responsible for the energetic deficit, which develops in proximal tubules after H/R.


Figure 7
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  		Fig. 7 Proposed mechanism of the protonophoric effect of nonesterified fatty acids (NEFA) during hypoxia and reoxygenation (H/R). NEFA act by attracting a proton on the cytosolic side of the inner mitochondrial membrane [1], followed by entering of the mitochondrial matrix in their protonated, nonpolar form [2]. After dissociation of the proton [3], NEFA are transported out of the matrix in their deprotonated polar form by the anion tricaroboxylate carrier (TCC) [4]. This results in a sustained proton entry into the mitochondrial matrix diminishing mitochondrial membrane potential ({Delta}{psi}).

 
The strong effect of dBSA to enhance mitochondrial safranin O uptake after H/R is explained by the strong binding capacity of dBSA for NEFA, preventing the interaction of NEFA with the mitochondria [9]. Albumin has seven binding sites for fatty acids [31] of which three share the highest affinity [32]. We have shown that in the presence of 7.57 µM dBSA, addition of 22 µM oleate was required to begin to dissipate {Delta}{psi} in normal tubules, 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. Tubules subjected to normoxia also showed increased mitochondrial membrane potential when dBSA was present during measurement of safranin O uptake. Our NEFA measurements on tubules sampled immediately at the end of normoxic incubation indicated background levels of NEFA sufficient to account for this, plus there could have been some additional accumulation during the cold holding period between sampling and the safranin O uptake assays. Spontaneous deterioration of mitochondrial function in isolated mitochondria due to NEFA and its restoration by dBSA is a well-known phenomenon and has been termed ‘mitochondrial aging’ [33–35]. The isolated tubules used in the present studies are relatively stable for several hours held in the cold prior to digitonin permeabilization for safranin uptake, but there is some dBSA-reversible deterioration of maximal safranin uptakethat occurs.

We have provided evidence using isolated rabbit tubules that the anion carriers, adenine nucleotide translocase and the glutamate/aspartate carrier, in the inner mitochondrial membrane are involved in the mechanism of NEFA-induced uncoupling effect and that this is relevant for the development of the energetic deficit that develops in proximal tubules after H/R [15]. In isolated mitochondria, it has been reported that the tricarboxylate carrier is also involved in the mechanism of NEFA-induced uncoupling [36]. The tricarboxylate carrier is a member of the SLC25 family of mitochondrial anion carriers that includes the aspartate/glutamate carrier, the dicarboxylate carrier, the phosphate carrier and the uncoupling proteins. It catalyses the electroneutral exchange of a tricarboxylate (e.g. citrate, isocitrate) for either another tricarboxylate, a dicarboxylate (e.g. malate) or PEP [16]. In the present studies, addition of citrate to compete for the tricarboxylate carrier attenuated the mitochondrial de-energization in proximal tubules induced by the NEFA oleate or H/R (Figures 5 and 6). This suggests that the tricarboxylate carrier is also involved in the mechanism of NEFA-induced uncoupling effect after H/R in proximal tubules.

Previous studies of NEFA effects on tubules during H/R have all used rabbit tubules [9,15]. The present studies importantly show that the findings are applicable to a rodent model as well and, therefore, represent a general, fundamental mechanism for mitochondrial damage during H/R in the proximal tubule. The freshly isolated rat tubules developed the same impairment of ATP production after H/R (Figure 1) and partial but incomplete recovery of mitochondrial {Delta}{psi} even after full reoxygenation (Figure 2). As with the rabbit tubules, dBSA, added during the measurement of mitochondrial safranin O uptake, enhanced mitochondrial safranin O uptake in proximal rat tubules, showing that NEFA are responsible for the decrease of mitochondrial {Delta}{psi}.

In summary, we have shown that the energetic deficit that develops in freshly isolated rat proximal tubules during H/R is characterized by impaired ATP production after full reoxygenation, accumulation of NEFA, impaired recovery of {Delta}{psi} and abrogation of {Delta}pH, and is ameliorated by citrate. All of the mitochondrial changes were also mimicked by exogenous addition of the NEFA oleate. {Delta}p was reduced in mitochondria from freshly isolated rat proximal tubules after H/R, and the proportion of {Delta}{psi} and {Delta}pH did not change even under injury conditions. Mitochondrial {Delta}{psi} is still the predominant component of {Delta}p, so measurement of {Delta}{psi} is sufficient to determine the ability of mitochondria to produce ATP during H/R. These data support the concept that the energetic deficit, which develops in proximal tubules after H/R can be explained by NEFA-induced protonophoric uncoupling and that this pathomechanism of tubule injury is a fundamental cellular response to mitochondrial damage during H/R.



   Acknowledgments
 
This study was supported by a grant from the German Research Foundation to T.F. and A.K. (DFG; Fe-929). J.M.W. acknowledges funding from the National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-34275 and the Department of Veterans’ Affairs. J.N. acknowledges funding from the German Research Foundation (DFG, Nu 118). We thank Barbara Nilewski-Kühl and Simone Leyting for excellent technical assistance.

Conflict of interest statement. None declared.



   References
Top
 Abstract
 Introduction
 Subjects and methods
 Results
 Discussion
 References
 

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Received for publication: 2. 3.08
Accepted in revised form: 8. 7.08


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