BIOCHEMICAL
MEDICINE
AND
METABOLIC
BIOLOGY
4,
266-270 (19%))
Inhibition of Fatty Acid-Supported Mitochondrial Respiration by Cyclosporine CARLOS A. E. LEMMI, RAYMOND L. MILLER, AND JACOBRAJFER Department of Anatomy and Cell Biology, UCLA, Los Angeles, California 90024; Division of Urology, Department of Surgery, Harbor-UCLA Medical Center, Torrance, California 90509; and The Saint John’s Heart Institute, Santa Monica, California 90404
Received June 20, 1990; and in revised form July 3, 1!490 Nephrotoxic damage due to cyclosporine (CS) is still a major limiting factor in the clinical use of this immunosuppressive drug (1). Alterations in lipid metabolism, leading to the formation of thromboxane, have been correlated with the nephrotoxic event (2,3). Renal damage can be prevented, or greatly ameliorated, by inhibitors of thromboxane formation (4). In our previous studies (5), we reported inhibition of mitochondrial Complex II by CS using succinate as substrate. Since mitochondrial Complex II also participates in the oxidation of fatty acids (6), we designed a group of experiments to determine whether CS also inhibits fatty acid-supported mitochondrial respiration. METHODS Isolation of mirochondriu. A modification of the technique utilized in previous works was used (5-8). Briefly, renal cortices from adult 300-g Sprague-Dawley male rats (Simonsen Laboratories, Gilroy, CA) were homogenized in 0.25 M sucrose, 5 mu Tris-HCl, 2 mu EGTA at pH 7.5 and 4°C. Mitochondria were separated by differential centrifugation at 800 and 12,OOOg. The mitochondrial pellet was then washed three times and finally resuspended in 50 ~1 of the same solution. Protein was determined by the Biuret method (9). The integrity of mitochondria was confirmed by their coupled response to ADP. Mitochondrial function and inhibition. A modification of our previous methodology was used (5,8). A Clark oxygen electrode, in a 1.8~ml water-jacketed 37°C cell, was filled with 10 mu Mops, 7 mM Trisma base, 2.5 mu H2KP04, 2.5 mM MgClz, 15 mM KCl, 235 mM sucrose, 0.5 mu malate, 2 mu acetoacetate, 0.5 mM rotenone, pH 7.4, and 50 mu palmitoyl-L-camitine as fatty acid substrate. Rotenone, an inhibitor of Complex I, and acetoacetate, to divert NADH to the formation of p-hydroxybutyrate, were used to assure that only electrons from the flavoprotein-linked oxidation were coupled to the measured mitochondrial respiration. Malate was added to obtain a more physiological situation (8). Then, 750 mg mitochondrial preparation was added and the rate of oxygen uptake was
0885-4505190 $3.00 Copyright All rights
Q 1990 by Academic Press, Inc. of reproduction in any form reserved.
CYCLOSPORINE
FATTY
ACID MITOCHONDRIA
P(0.005 +-I
3asd
Respiration
267
!I
DNP-Stimulated
Respiration
FIG. 1. Basal nonstimulated and DNP-stimulated mitochondrial respiration in the presence (hatched bars) and absence (open bars) of 25 &ml CS. Palmitoyl-L-camitine at a concentration of 50 mM was used as substrate (n = 12 animals).
recorded for 1 min (State 2). At this time 2,4-dinitrophenol (DNP) (final concentration 44 PM) was introduced into the chamber and the maximum rate of uncoupled electron transport respiration was recorded. Oxygen consumption calculations were based on an oxygen solubility in buffer of 390 ng atoms O/ml at 37°C. Oxygen consumption rates were expressed as nanograms atoms of oxygen per minute per milligram of mitochondrial protein (ng atoms O/min/mg protein). Cyclosporine in experimental studies. Cyclosporine A in ethanol was added to the oxygraph chamber just before the introduction of mitochondria. Control studies using solvent alone showed that ethanol (0.12 mM) had no effect on the rate of mitochondrial respiration (5). Statistical analysis. The studies were performed in paired fashion (i.e., control and CS-treated mitochondria experiments performed on the same mitochondrial preparation on the same day) and the paired Student t test was used for statistical comparison. Data were expressed as means 2 SEM. RESULTS Effect of cyclosporine on palmitoyk-carnitine-dependent mitochondrial respiration. Cyclosporine, at a concentration of 25 pg/ml, inhibited (I’ < 0.005) by 24.0 t: 13.6% the basal nonstimulated mitochondrial respiration in the presence of 50 mM palmitoyl-L-camitine (n = 12 animals) from 78.4 + 2.4 to 59.4 f 3.3 ng atom O/min/mg protein. Under the same concentrations of inhibitor and substrate, cyclosporine inhibited (P < 0.005) by 37.1 -+ 15.2% the uncoupled electron transport respiratory rate (n = 12 animals) from 133.1 -I 9.4 to 82.4 k 7.3 ng atom O/min/mg protein (Fig. 1). Effect of substrate concentration on the inhibition of mitochondrial respiratory rate by cyclosporine. The inhibition of palmitoyl-L-camitine-dependent mitochondrial respiration was repeated in a separate group of isolated mitochondria (n = 8 animals) using CS at a concentration of 25 pg/ml (2.08 x 10m5 M). The substrate concentrations used were 17, 33, 50, and 67 mM.
268
LEMMI,
MILLER,
AND RAJFER
I A I
P(O.025
/ ---.-- ..------.AT 1 1 P(O.WS
.-
..A
A
5
I
10
20
30 Palmitic
FIG.
2.
tochondrial
40 acid
50
60
70
(mu)
Effect of various concentrations of palmitoyl-L-camitine on the basal nonstimulated mirespiration in the presence (dotted lines) and absence (solid lines) of 25 &ml CS.
Figure 2 shows that the basal nonstimulated mitochondrial respiration was inhibited by CS at substrate concentrations of 33 mM (P < 0.025) or higher (P < 0.005). Figure 3 shows CS inhibition at concentrations of substrate of 50 mM (P < 0.025) or higher (P < 0.005) under conditions of uncoupled mitochondrial respiration. DISCUSSION
In previous studies we documented that CS inhibits mitochondrial respiration at the level of Complex II (5). Succinate-coenzyme Q reductase (Complex II) is a membrane-bound enzymatic complex that includes the enzyme succinate dehydrogenase, FAD, three to four iron-sulfur centers, and cytochrome bSM. Complex II transfers electrons from succinate to coenzyme Q (6). In the oxidation of fatty acids, Complex II interacts with an electron transfer flavoprotein which transfers electrons between reduced flavin and coenzyme Q (6,lO). In this study, in the absence of succinate, basal fatty acid-supported mitochondrial respiration is inhibited by CS. This inhibition also occurs with maximum respiratory rates under conditions of uncoupled electron transport. Nephrotoxic damage, a major clinical consideration in the use of this drug, has been correlated with formation of thromboxane, a lipid of the eicosanoid family (2-4). Calcium release, membrane damage, and other conditions resulting in increased levels of arachidonic acid result in formation of eicosanoids, such as prostaglandins and thromboxans (11). Thromboxane formation results in decrease renal blood flow, which is suspected to cause renal damage (12). In a separate study we reported how this ischemic condition results in a compensatory mitochondrial response involving Complex I (13). Fatty acids and their acyl-CoA forms are not permeable and they must be converted to fatty acyl carnitine derivatives to penetrate the inner mitochondrial membrane (6). We used palmitoyl+camitine, a 16-carbon saturated fatty acid which is already covalently bound to camitine and readily penetrates the mitochondrial membrane. Arachidonic acid, a 20-carbon unsaturated fatty acid, is a more direct precursor of eicosanoids. Unfortunately, arachidonic acid was not
CYCLOSPORINE
G;f F’B 0
FATTY
269
ACID MITOCHONDRIA
5-e
‘-,I
-5--157
I 0
10
20
30 Polmitic
40 Acid
50
60
I 70
(mu)
3. Effect of various concentrations of palmitoyl-L-camitine on the maximal uncoupled mitochondrial respiration in the presence (dotted lines) and absence (solid lines) of 25 pg/ml CS. FIG.
available in the arachidonyl-L-camitine form. Therefore, we used the more economical and readily available palmitoyl-L-camitine acid. It can be speculated that, if CS inhibits mitochondrial lipid oxidation, the level of fatty acids would increase leading to formation of eicosanoids. Usually, eicosanoid formation results from a sudden release of arachidonic acid by phospholipase activation (11). In our system, the observed inhibition of fatty acid oxidation may result in a slow and progressive increase in lipid levels, eventually leading to formation of thromboxane. SUMMARY
We have shown that, in addition to inhibition of the succinate-supported energy pathway (5), CS inhibition of mitochondrial Complex II activity also limits fatty acid oxidation. These results are consistent with the participation of altered lipid metabolism in CS nephrotoxicity. ACKNOWLEDGMENTS The authors thank Drs. L. LeDuc and G. Chaudhuri for their assistance in the thromboxane studies. We thank Mr. Jayendra P. Sharma and Ms. Irena Kumar for expert technical support. This project was supported in part by the Saint John’s Heart Institute Cardiac Research Center Grant SJ-5994 and the National Institutes of Health Biomedical Research Support Grant S07-RRO5551. Research and Education Institute of Harbor-UCLA Medical Center Grant GR-5917 to Dr. Lemmi.
REFERENCES 1. Kahan, B., Transplant. Proc. 17, 1 (1985). 2. Kawaguchi, A., Goldman, M. H., Shapiro, R., Foegh, M. L., Ramwell, P. W., and Lower, R., Trunspluntution 40(2), 214 (1985). 3. Coffman, T. M., Carr, D. R., Yarger, W. E., and Klotman, P. E., Transplantarion 43(2), 282 (1987). 4. Smeersters, C., Chaland, P., Giroux, L., Moutquin, J. M., Etienne, P., Douglas, F., Corman, J., St-Louis, G., and Daloze, P., Transplant. Proc. (Suppl. 2) 20(2), 663 (1988). 5. Lemmi, C. A. E., Pelikan, P. C. D., Geesaman, B., Seamon, E., Koyle, M., and Rajfer, J., Biochem. Med. Metub. Eiol. 43(3), 214 (1990). 6. Tzagoloff, A., “Mitochondria.” Plenum, New York (1982).
270 7. 8. 9. 10. 11. 12. 13.
LEMMI,
MILLER,
AND RAJFER
Junk, K., Reinholdt, C., and Scholz, D., Ziunspluntation 43, 162 (1987). Halestrap, A. P., and Dunlop, J. L., Biochem. J. 239, 559 (1986). Doumas, B., Bayse, D., Carter, D., Peters, R., and SchatTer, R., Clin. Chem. 27, 1642 (1981). Gustafson, W. G., Feinberg, B. A., and McFarland, J. T., .I. Biol. Chem. 261(17), 7733 (1986). Flower, R. J., and Blackwell, G. J., Biochem. Pharmacol. 25, 285 (1976). Coffman, T. M., Yarger, W. E., and Klotman, P. E., J. Clin. Invest. 75, 1242 (1985). Lemmi, C. A. E., Pelikan, P. C. D., Sikka, S. C., Hirschberg, R., Geesaman, B., Miller, R. L., Park, K. S., Liu, S., Koyle, M., and Rajfer, J., Amer. J. Physiol. 257 (Renal Fluid Electrolyte Physiol. 26), F837 (1989).
MEDICINE
AND
METABOLIC
BIOLOGY
4,
266-270 (19%))
Inhibition of Fatty Acid-Supported Mitochondrial Respiration by Cyclosporine CARLOS A. E. LEMMI, RAYMOND L. MILLER, AND JACOBRAJFER Department of Anatomy and Cell Biology, UCLA, Los Angeles, California 90024; Division of Urology, Department of Surgery, Harbor-UCLA Medical Center, Torrance, California 90509; and The Saint John’s Heart Institute, Santa Monica, California 90404
Received June 20, 1990; and in revised form July 3, 1!490 Nephrotoxic damage due to cyclosporine (CS) is still a major limiting factor in the clinical use of this immunosuppressive drug (1). Alterations in lipid metabolism, leading to the formation of thromboxane, have been correlated with the nephrotoxic event (2,3). Renal damage can be prevented, or greatly ameliorated, by inhibitors of thromboxane formation (4). In our previous studies (5), we reported inhibition of mitochondrial Complex II by CS using succinate as substrate. Since mitochondrial Complex II also participates in the oxidation of fatty acids (6), we designed a group of experiments to determine whether CS also inhibits fatty acid-supported mitochondrial respiration. METHODS Isolation of mirochondriu. A modification of the technique utilized in previous works was used (5-8). Briefly, renal cortices from adult 300-g Sprague-Dawley male rats (Simonsen Laboratories, Gilroy, CA) were homogenized in 0.25 M sucrose, 5 mu Tris-HCl, 2 mu EGTA at pH 7.5 and 4°C. Mitochondria were separated by differential centrifugation at 800 and 12,OOOg. The mitochondrial pellet was then washed three times and finally resuspended in 50 ~1 of the same solution. Protein was determined by the Biuret method (9). The integrity of mitochondria was confirmed by their coupled response to ADP. Mitochondrial function and inhibition. A modification of our previous methodology was used (5,8). A Clark oxygen electrode, in a 1.8~ml water-jacketed 37°C cell, was filled with 10 mu Mops, 7 mM Trisma base, 2.5 mu H2KP04, 2.5 mM MgClz, 15 mM KCl, 235 mM sucrose, 0.5 mu malate, 2 mu acetoacetate, 0.5 mM rotenone, pH 7.4, and 50 mu palmitoyl-L-camitine as fatty acid substrate. Rotenone, an inhibitor of Complex I, and acetoacetate, to divert NADH to the formation of p-hydroxybutyrate, were used to assure that only electrons from the flavoprotein-linked oxidation were coupled to the measured mitochondrial respiration. Malate was added to obtain a more physiological situation (8). Then, 750 mg mitochondrial preparation was added and the rate of oxygen uptake was
0885-4505190 $3.00 Copyright All rights
Q 1990 by Academic Press, Inc. of reproduction in any form reserved.
CYCLOSPORINE
FATTY
ACID MITOCHONDRIA
P(0.005 +-I
3asd
Respiration
267
!I
DNP-Stimulated
Respiration
FIG. 1. Basal nonstimulated and DNP-stimulated mitochondrial respiration in the presence (hatched bars) and absence (open bars) of 25 &ml CS. Palmitoyl-L-camitine at a concentration of 50 mM was used as substrate (n = 12 animals).
recorded for 1 min (State 2). At this time 2,4-dinitrophenol (DNP) (final concentration 44 PM) was introduced into the chamber and the maximum rate of uncoupled electron transport respiration was recorded. Oxygen consumption calculations were based on an oxygen solubility in buffer of 390 ng atoms O/ml at 37°C. Oxygen consumption rates were expressed as nanograms atoms of oxygen per minute per milligram of mitochondrial protein (ng atoms O/min/mg protein). Cyclosporine in experimental studies. Cyclosporine A in ethanol was added to the oxygraph chamber just before the introduction of mitochondria. Control studies using solvent alone showed that ethanol (0.12 mM) had no effect on the rate of mitochondrial respiration (5). Statistical analysis. The studies were performed in paired fashion (i.e., control and CS-treated mitochondria experiments performed on the same mitochondrial preparation on the same day) and the paired Student t test was used for statistical comparison. Data were expressed as means 2 SEM. RESULTS Effect of cyclosporine on palmitoyk-carnitine-dependent mitochondrial respiration. Cyclosporine, at a concentration of 25 pg/ml, inhibited (I’ < 0.005) by 24.0 t: 13.6% the basal nonstimulated mitochondrial respiration in the presence of 50 mM palmitoyl-L-camitine (n = 12 animals) from 78.4 + 2.4 to 59.4 f 3.3 ng atom O/min/mg protein. Under the same concentrations of inhibitor and substrate, cyclosporine inhibited (P < 0.005) by 37.1 -+ 15.2% the uncoupled electron transport respiratory rate (n = 12 animals) from 133.1 -I 9.4 to 82.4 k 7.3 ng atom O/min/mg protein (Fig. 1). Effect of substrate concentration on the inhibition of mitochondrial respiratory rate by cyclosporine. The inhibition of palmitoyl-L-camitine-dependent mitochondrial respiration was repeated in a separate group of isolated mitochondria (n = 8 animals) using CS at a concentration of 25 pg/ml (2.08 x 10m5 M). The substrate concentrations used were 17, 33, 50, and 67 mM.
268
LEMMI,
MILLER,
AND RAJFER
I A I
P(O.025
/ ---.-- ..------.AT 1 1 P(O.WS
.-
..A
A
5
I
10
20
30 Palmitic
FIG.
2.
tochondrial
40 acid
50
60
70
(mu)
Effect of various concentrations of palmitoyl-L-camitine on the basal nonstimulated mirespiration in the presence (dotted lines) and absence (solid lines) of 25 &ml CS.
Figure 2 shows that the basal nonstimulated mitochondrial respiration was inhibited by CS at substrate concentrations of 33 mM (P < 0.025) or higher (P < 0.005). Figure 3 shows CS inhibition at concentrations of substrate of 50 mM (P < 0.025) or higher (P < 0.005) under conditions of uncoupled mitochondrial respiration. DISCUSSION
In previous studies we documented that CS inhibits mitochondrial respiration at the level of Complex II (5). Succinate-coenzyme Q reductase (Complex II) is a membrane-bound enzymatic complex that includes the enzyme succinate dehydrogenase, FAD, three to four iron-sulfur centers, and cytochrome bSM. Complex II transfers electrons from succinate to coenzyme Q (6). In the oxidation of fatty acids, Complex II interacts with an electron transfer flavoprotein which transfers electrons between reduced flavin and coenzyme Q (6,lO). In this study, in the absence of succinate, basal fatty acid-supported mitochondrial respiration is inhibited by CS. This inhibition also occurs with maximum respiratory rates under conditions of uncoupled electron transport. Nephrotoxic damage, a major clinical consideration in the use of this drug, has been correlated with formation of thromboxane, a lipid of the eicosanoid family (2-4). Calcium release, membrane damage, and other conditions resulting in increased levels of arachidonic acid result in formation of eicosanoids, such as prostaglandins and thromboxans (11). Thromboxane formation results in decrease renal blood flow, which is suspected to cause renal damage (12). In a separate study we reported how this ischemic condition results in a compensatory mitochondrial response involving Complex I (13). Fatty acids and their acyl-CoA forms are not permeable and they must be converted to fatty acyl carnitine derivatives to penetrate the inner mitochondrial membrane (6). We used palmitoyl+camitine, a 16-carbon saturated fatty acid which is already covalently bound to camitine and readily penetrates the mitochondrial membrane. Arachidonic acid, a 20-carbon unsaturated fatty acid, is a more direct precursor of eicosanoids. Unfortunately, arachidonic acid was not
CYCLOSPORINE
G;f F’B 0
FATTY
269
ACID MITOCHONDRIA
5-e
‘-,I
-5--157
I 0
10
20
30 Polmitic
40 Acid
50
60
I 70
(mu)
3. Effect of various concentrations of palmitoyl-L-camitine on the maximal uncoupled mitochondrial respiration in the presence (dotted lines) and absence (solid lines) of 25 pg/ml CS. FIG.
available in the arachidonyl-L-camitine form. Therefore, we used the more economical and readily available palmitoyl-L-camitine acid. It can be speculated that, if CS inhibits mitochondrial lipid oxidation, the level of fatty acids would increase leading to formation of eicosanoids. Usually, eicosanoid formation results from a sudden release of arachidonic acid by phospholipase activation (11). In our system, the observed inhibition of fatty acid oxidation may result in a slow and progressive increase in lipid levels, eventually leading to formation of thromboxane. SUMMARY
We have shown that, in addition to inhibition of the succinate-supported energy pathway (5), CS inhibition of mitochondrial Complex II activity also limits fatty acid oxidation. These results are consistent with the participation of altered lipid metabolism in CS nephrotoxicity. ACKNOWLEDGMENTS The authors thank Drs. L. LeDuc and G. Chaudhuri for their assistance in the thromboxane studies. We thank Mr. Jayendra P. Sharma and Ms. Irena Kumar for expert technical support. This project was supported in part by the Saint John’s Heart Institute Cardiac Research Center Grant SJ-5994 and the National Institutes of Health Biomedical Research Support Grant S07-RRO5551. Research and Education Institute of Harbor-UCLA Medical Center Grant GR-5917 to Dr. Lemmi.
REFERENCES 1. Kahan, B., Transplant. Proc. 17, 1 (1985). 2. Kawaguchi, A., Goldman, M. H., Shapiro, R., Foegh, M. L., Ramwell, P. W., and Lower, R., Trunspluntution 40(2), 214 (1985). 3. Coffman, T. M., Carr, D. R., Yarger, W. E., and Klotman, P. E., Transplantarion 43(2), 282 (1987). 4. Smeersters, C., Chaland, P., Giroux, L., Moutquin, J. M., Etienne, P., Douglas, F., Corman, J., St-Louis, G., and Daloze, P., Transplant. Proc. (Suppl. 2) 20(2), 663 (1988). 5. Lemmi, C. A. E., Pelikan, P. C. D., Geesaman, B., Seamon, E., Koyle, M., and Rajfer, J., Biochem. Med. Metub. Eiol. 43(3), 214 (1990). 6. Tzagoloff, A., “Mitochondria.” Plenum, New York (1982).
270 7. 8. 9. 10. 11. 12. 13.
LEMMI,
MILLER,
AND RAJFER
Junk, K., Reinholdt, C., and Scholz, D., Ziunspluntation 43, 162 (1987). Halestrap, A. P., and Dunlop, J. L., Biochem. J. 239, 559 (1986). Doumas, B., Bayse, D., Carter, D., Peters, R., and SchatTer, R., Clin. Chem. 27, 1642 (1981). Gustafson, W. G., Feinberg, B. A., and McFarland, J. T., .I. Biol. Chem. 261(17), 7733 (1986). Flower, R. J., and Blackwell, G. J., Biochem. Pharmacol. 25, 285 (1976). Coffman, T. M., Yarger, W. E., and Klotman, P. E., J. Clin. Invest. 75, 1242 (1985). Lemmi, C. A. E., Pelikan, P. C. D., Sikka, S. C., Hirschberg, R., Geesaman, B., Miller, R. L., Park, K. S., Liu, S., Koyle, M., and Rajfer, J., Amer. J. Physiol. 257 (Renal Fluid Electrolyte Physiol. 26), F837 (1989).
