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Am J Physiol Heart Circ Physiol 285: H154-H162, 2003. First published March 6, 2003; doi:10.1152/ajpheart.00955.2002
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Ischemic preconditioning inhibits mitochondrial respiration, increases H2O2 release, and enhances K+ transport

Mirian M. da Silva, Adriano Sartori, Eduardo Belisle, and Alicia J. Kowaltowski

Departamento de Bioquímica, Instituto de Química, Universidade de São Paulo, São Paulo SP 05508-900, Brazil

Submitted 7 November 2002 ; accepted in final form 5 March 2003


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Ischemic preconditioning, or the protective effect of short ischemic episodes on a longer, potentially injurious, ischemic period, is prevented by antagonists of mitochondrial ATP-sensitive K+ channels (mitoKATP) and involves changes in mitochondrial energy metabolism and reactive oxygen release after ischemia. However, the effects of ischemic preconditioning itself on mitochondria are still poorly understood. We determined the effects of ischemic preconditioning on isolated heart mitochondria and found that two brief (5 min) ischemic episodes are sufficient to induce a small but significant decrease (~25%) in mitochondrial NADH-supported respiration. Preconditioning also increased mitochondrial H2O2 release, an effect related to respiratory inhibition, because it is not observed in the presence of succinate plus rotenone and can be mimicked by chemically inhibiting complex I in the presence of NADH-linked substrates. In addition, preconditioned mitochondria presented more substantial ATP-sensitive K+ transport, indicative of higher mitoKATP activity. Thus we directly demonstrate that preconditioning leads to mitochondrial respiratory inhibition in the presence of NADH-linked substrates, increased reactive oxygen release, and activation of mitoKATP.

heart; ischemia-reperfusion; free radicals; NADH dehydrogenase; K+ channel

SINCE THE ORIGINAL DISCOVERY by Murry et al. (31) that brief ischemic periods (2–5 min) followed by reperfusion can protect heart tissue from damage after longer ischemic periods (20–50 min), many studies have focused attention on the signaling pathways involved in this process, known as ischemic preconditioning (IP). These studies have uncovered a series of intercommunicating pathways through which IP is activated, including adenosine and acetylcholine signaling (27, 48), protein kinase C activation (51), ATP-sensitive K+ channel (KATP) opening (16, 25, 48), and reactive oxygen species (ROS) generation (35, 46, 49, 50).

The most probable source of ROS generation leading to the activation of IP is the mitochondrial respiratory chain, which continuously produces low levels of ROS under physiological conditions and presents increased ROS release levels after periods of ischemia or anoxia (38, 53). Interestingly, increased ROS release also promotes at least part of the tissue damage caused by ischemia-reperfusion (46). Small and transient increases in mitochondrial ROS during IP seem to prevent the deleterious effects of ROS after long-term ischemia-reperfusion (45). Indeed, the idea that mitochondria actively participate in the IP signaling pathway by generating ROS is in line with the observation that mitochondrial function and membrane integrity can be key determinants in cellular viability (21, 39, 43).

Further evidence for the participation of mitochondria in IP was the finding that the pharmacological inhibition of KATP in mitochondria (mitoKATP) prevents IP (2, 17, 48). Also, IP can be mimicked by mitoKATP agonists (13, 16, 25). Despite these findings with the use of pharmacological mitoKATP regulators, no direct evidence for mitoKATP activation in IP has been presented to date. Unfortunately, the only technique described to follow mitoKATP activation in intact cells involves measurement of the redox state of flavin nucleotides (28), which can be affected by changes in respiratory rates and levels of reduced substrates.

Also, the mechanisms through which mitoKATP may be activated during IP are unknown. Kinase activation and phosphorylation of the channel have been suggested to participate in this process (32, 47), and kinases may have their activity increased by ROS (3, 24). In addition, ROS may directly activate mitoKATP (34, 52). Interestingly, there is also evidence that ROS act downstream of mitoKATP activation in IP (6, 10, 11, 26, 33; for discussion, see Ref. 35), although the mechanism through which these ROS are generated and the possible downstream effects of these ROS are unclear. Together, these results indicate that mitoKATP may participate together with mitochondrial ROS as an amplifying step within the IP signaling pathway. MitoKATP activation itself also presents many potentially important effects, which may directly promote ischemic protection, including preventing excessive mitochondrial contraction (9, 23), increasing the efficiency of oxidative phosphorylation during reperfusion (9), decreasing high-energy phosphate loss during ischemia (5), and preventing mitochondrial Ca2+ overload during ischemia (5, 30).

To date, no detailed experiments evaluating whether mitochondrial alterations promoted by long-term ischemia (14, 19) can be caused by the short ischemic periods that promote IP have been performed. Also, no direct measurement of mitochondrial ROS release and K+ transport after IP has been conducted. In this study, we isolated mitochondria from preconditioned hearts and measured membrane integrity, respiration, ROS release, and mitoKATP activity to better understand the role of mitochondria in IP.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Rat heart perfusions. Heart perfusion was conducted as described previously (5). Briefly, hearts were rapidly removed from male Sprague-Dawley rats, trimmed over ice, and Langendorff perfused with 200 ml of oxygenated Krebs-Henseleit buffer containing (in mM) 118 NaCl, 25 NaHCO3, 1.2 KH2PO4, 4.7 KCl, 1.2 MgSO4, 1.25 CaCl2, 10 glucose, and 10 HEPES, pH 7.0, at 37°C. Heart beat rates were left unpaced, and perfusion was maintained at a constant pressure of 70 mmHg. During the first 5 min of perfusion, a nonrecirculating mode was used to eliminate contaminating blood. After stabilization, a recirculating perfusion mode was initiated. Control hearts were then perfused for 20 min, whereas preconditioned hearts were submitted to two 5-min periods without perfusion, as illustrated in Fig. 1.



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  		Fig. 1. Experimental protocols. All hearts were washed at 37°C for 5 min in a nonrecirculating mode, followed by 20-min continuous recirculating perfusion (control) or two 5-min periods of no-flow 37°C ischemia, followed by 5-min reperfusion (preconditioned). After the control or preconditioning period, the hearts were quickly transferred to ice-cold buffer for mitochondrial isolation. Perfusate creatine kinase activity was determined with the use of a separate group of hearts submitted to 20-min 37°C no-flow ischemia, followed by 10-min reperfusion.

 

To ascertain that IP was effective under our conditions, we submitted a group of control and preconditioned hearts to 20-min ischemia at 37°C, followed by reperfusion, and determined creatine kinase activity in aliquots of the recirculating perfusate (Fig. 2). As expected, we found that creatine kinase activity, a measurement of cellular damage, was significantly higher in control hearts compared with preconditioned hearts. All other experiments were conducted using mitochondria isolated from hearts submitted only to a 25-min perfusion or preconditioning period (Fig. 1) to determine the mitochondrial effects of IP.



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  		Fig. 2. Preconditioning decreases postischemic creatine kinase release. Rat hearts were Langendorff perfused for 25 min (control) or preconditioned through two 5-min ischemic periods and submitted to 20-min ischemia, followed by reperfusion (see Fig. 1). Aliquots of perfusate were collected after 10-min reperfusion to determine creatine kinase activity. *P < 0.05 relative to control, n = 3.

 

All studies were conducted in accordance with the Universidade de São Paulo protocol and guidelines for animal care and use. Hearts were eliminated from the study if the time between heart removal and the beginning of perfusion was >3 min.

Creatine kinase activity. Creatine kinase activity was determined in frozen perfusate aliquots (1 ml) using commercial kits (catalog no. 1070, Doles; São Paulo, Brazil), accompanying the time-dependent formation of NADH at excitation = 352 nm and emission = 464 nm on a Hitachi F4500 spectrofluorometer between 5 and 10 min after the reaction was started, when traces showed maximum linearity (5). Curves were quantified with a calibration curve prepared using lyophilized bovine heart creatine kinase.

Mitochondrial isolation. Rat heart mitochondria were isolated as described previously (5). Briefly, the Langendorff-perfused hearts were washed in ice-cold buffer containing 300 mM sucrose, 10 mM K+-HEPES buffer, pH 7.2, and 1 mM K+ EGTA. The tissue was finely minced and incubated in the presence of 1 mg protease type XXIV (Sigma Aldrich) for 10 min. Excess protease was removed by washing the heart fragments in the same buffer containing 1 mg/ml BSA, and the samples were homogenized manually. The resulting suspension was centrifuged at 600 g for 4 min, and the supernatant was recentrifuged at 9,000 g for 8 min. The mitochondrial pellet was then washed once or twice until a compact pellet was obtained. This pellet was suspended in 200–300 µl of BSA-containing buffer and kept over ice for up to 4 h.

Mitochondrial respiration. Respiration was measured using a computer-interfaced Clark-type oxygen electrode from Hansatech Instruments equipped with magnetic stirring. Oxygen solubility at 37°C was taken to be 220 nmol/ml.

Mitochondrial membrane potential estimation. Mitochondrial membrane potential ({Delta}{Psi}) was estimated by following safranine O (1, 22) fluorescence at excitation = 495 nm and emission = 586 nm on a Hitachi F4500 spectrofluorometer.

Mitochondrial H2O2 release. H2O2 was measured in the mitochondrial suspension by following the oxidation of Amplex red (Molecular Probes) in the presence of horseradish peroxidase (HRP) recorded on a temperature-controlled Hitachi F4500 fluorescence spectrophotometer equipped with continuous stirring at excitation and emission wavelengths of 563 and 587 nm, respectively (29). Because Amplex red presents a slow rate of spontaneous oxidation in the presence of HRP, all traces were subtracted from a baseline trace recorded in the same media devoid of mitochondria. The data were quantified by adding known quantities of a freshly prepared H2O2 stock calibrated by its absorbance at 240 nm (E = 43.6 M/cm).

NAD(P)H fluorescence. Mitochondrial NAD(P)H fluorescence was measured at excitation = 352 nm and emission = 464 nm on a spectrofluorometer. Rotenone and carbonyl cyanide 3-chlorophenylhydrazone (CCCP) were used to obtain maximal pyridine nucleotide reduction and oxidation levels, respectively.

Mitochondrial swelling. Changes in mitochondrial volume, which accompany net salt transport into mitochondria (4), were followed with the use of a Hitachi F4500 fluorescence spectrophotometer operating at excitation and emission wavelengths of 520 nm, with 2.5-nm slits.

Reagents. Amplex red was purchased from Molecular Probes. HRP (P8125), safranine O, EGTA, malate, glutamate, pyruvate, BSA, CCCP, rotenone, and valinomycin were from Sigma-Aldrich.

Data analysis. Data shown illustrate either representative traces or means ± SE for 3–6 repetitions with the use of different mitochondrial preparations. Multiple pairwise Tukeys tests conducted using SigmaStat software were used for comparisons between experimental groups.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
The effects of IP on isolated heart mitochondria were determined to uncover mitochondrial alterations that may be protective during ischemia. Figure 3A shows measurements of maximal (ADP stimulated) respiratory rates supported by different substrates. We found that preconditioned mitochondria presented smaller respiratory rates compared with control mitochondria when respiring on pyruvate or malate plus glutamate. With either succinate or N,N,N'N'-tetramethyl-1,3-propandiamine (TMPD)/ascorbate, which donate electrons to complex II and cytochrome c, respectively, respiratory rates in control and preconditioned mitochondria were equal. Because the difference in respiration was only present using NADH-linked substrates, these data indicate that IP partially inhibits mitochondrial complex I but not complexes II, III, or IV. Despite the small respiratory inhibition observed with NADH-linked substrates, preconditioned mitochondria maintained respiratory control ratios (Fig. 3B) similar to control mitochondria, with the use of either pyruvate or succinate as substrates, suggesting that IP does not significantly hamper ATP synthesis.



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  		Fig. 3. Preconditioning decreases complex I activity but does not affect respiratory control ratios, outer membrane integrity, or change in membrane potential ({Delta}{Psi}). Preconditioned or control mitochondria (0.5 mg/ml) were added to reaction medium at 37°C containing 125 mM KCl, 0.1 mM EGTA, 1 mg/ml BSA, 1 mM MgCl2, 10 mM HEPES, 2 mM inorganic phosphate (Pi), and 1 mM ADP, pH 7.2 (KOH). A: respiratory rates were measured in the presence of 2 mM pyruvate, 2 mM malate and 2 mM glutamate, 10 mM succinate and 1 µM rotenone, or 2 mM N,N,N',N'-tetramethyl-1,3-propandiamine (TMPD), 2 mM ascorbate (asc), and 1 µM antimycin A (AA), as shown. B: ADP-stimulated respiratory rates obtained in the presence of 2 mM pyruvate or 10 mM succinate were divided by respiratory rates in the presence of the same substrates with added oligomycin (1 µg/ml) to obtain the respiratory control ratio. C: O2 consumption supported by 10 mM succinate was measured, and 5 µM cytochrome c was added where indicated. Numbers in parenthesis indicate respiratory rates (in nmol·min-1·mg-1). D: mitochondria (1 mg/ml) were incubated in 250 mM sucrose, 0.1 mM EGTA, 10 mM succinate, 1 mg/ml BSA, 10 mM HEPES, and 2 mM Pi, pH 7.2 (NaOH), containing 0.5 µg/ml valinomycin and 5 µM safranine O at 37°C. Safranine fluorescence, which is inversely proportional to {Delta}{Psi}, was measured in the presence of increasing KCl concentrations (0.15, 0.3, 0.45, 0.75, 1.05, 1.35, and 1.95 mM). Carbonyl cyanide 3-chlorophenylhydrazone (CCCP; 5 µM) was added where indicated. A.U., arbitrary units. *P < 0.05 relative to control, n = 3.

 

We also measured mitochondrial outer membrane permeability to cytochrome c after IP because outer membrane permeabilization resulting in cytochrome c loss is an early indicator of tissue damage in ischemic hearts (19). In Fig. 3C, ADP-stimulated respiratory rates supported by succinate oxidation were compared before and after the addition of exogenous cytochrome c. A small increase in respiration was observed after this addition in both preconditioned and control mitochondria, indicative of an essentially intact outer membrane under both conditions. These data are in accordance to the finding that maximal respiratory rates supported by succinate or TMPD/ascorbate oxidation are equal in control and preconditioned mitochondria (Fig. 3A), suggesting no significant loss of cytochrome c occurs during IP.

To determine whether inner membrane permeability was affected by IP, we measured the change in {Delta}{Psi} in control and preconditioned mitochondria by using the fluorescent {Delta}{Psi} probe safranine O. We found that both control and preconditioned mitochondria decreased the fluorescence of the {Delta}{Psi} probe safranine to very similar levels both in the absence (not shown) and presence (Fig. 3D) of valinomycin. Because safranine fluorescence quenching can be affected by parameters other than {Delta}{Psi}, such as mitochondrial volume (22), {Delta}{Psi} traces were calibrated using a K+ gradient in the presence of valinomycin (1, 22). We found that control and preconditioned mitochondria presented equal fluorescence increments in the presence of added K+, confirming that {Delta}{Psi} is equivalent in these mitochondria and that IP does no affect inner membrane integrity and proton permeability.

Next, we measured mitochondrial H2O2 release to evaluate whether changes in ROS release observed previously in preconditioned cells (3, 35, 46) can be measured directly in isolated mitochondria from preconditioned hearts. Indeed, we found that H2O2 release levels in mitochondria from preconditioned hearts were constantly higher than H2O2 release levels in control mitochondria (Fig. 4A). On average (Fig. 4B), H2O2 release was significantly enhanced in preconditioned mitochondria in oligomycin-induced state 4 (in which the lack of ATP synthesis results in high {Delta}{Psi}, low respiratory rates, and augmented H2O2 release) and in the presence of CCCP, which increases respiration and decreases {Delta}{Psi} and ROS generation (20). H2O2 release also tended to be larger in preconditioned mitochondria in the presence of ADP. Because H2O2 release was increased even in the presence of CCCP, this increase is not due to changes in mitochondrial ion transport, such as K+ transport through the mitoKATP or H+ transport through uncoupling proteins or the proton leak (18). The increased H2O2 release rates in preconditioned mitochondria must therefore be related to changes in respiratory chain function. Indeed, respiratory chain inhibition promoted by myxothiazol or antimycin resulted in equal H2O2 release levels in control and preconditioned mitochondria. In addition, we were not able to measure any difference in H2O2 release when mitochondria were incubated in the presence of succinate plus rotenone (a chemical inhibitor of complex I), suggesting that the increase in H2O2 production measured is due to changes in NADH oxidation.



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  		Fig. 4. Preconditioning increases mitochondrial H2O2 release. Preconditioned or control mitochondria (0.5 mg/ml) were added to a reaction medium at 37°C containing 125 mM KCl, 0.1 mM EGTA, 1 mg/ml BSA, 1 mM MgCl2, 50 µM Amplex red, 1 U/ml horseradish peroxidase (HRP), 10 mM HEPES, and 2 mM Pi, pH 7.2 (KOH), in the presence of 2 mM pyruvate and 1 µg/ml oligomycin (A) or 2 mM pyruvate, 0.5 mM ADP, 2 mM succinate, 1 µM rotenone, 1 µg/ml oligomycin, 1 µM CCCP, 1.25 µg/ml myxothiazol (myx), and/or 0.5 µg/ml AA, as indicated (B). H2O2 release was measured as described in MATERIALS AND METHODS. A: representative traces obtained under each experimental condition are shown. B: means ± SE of 3–6 repetitions using different mitochondrial preparations under each experimental condition are shown. *P < 0.05 relative to control.

 

We then determined the levels of levels of pyridine nucleotides in our preparations in addition to the NAD(P)H-to-NAD(P)+ ratios to verify whether the changes in H2O2 release measured were linked to increased pyridine nucleotide levels or reduction. As expected, we found pyridine nucleotides were almost completely reduced during state 4 respiration and were rapidly oxidized when CCCP was added to both control and preconditioned mitochondria (results not shown). No significant change in mitochondrial pyridine nucleotide redox state or content was observed in preconditioned hearts.

To investigate whether the partial decrease in complex I activity observed in Fig. 3A was responsible for the increase in H2O2 observed in Fig. 4, we used low doses of rotenone on control mitochondria to mimic the respiratory inhibition effect of IP. We found that the addition of rotenone (5 nM) to control mitochondria promoted a respiratory inhibition similar to that observed in preconditioned mitochondria (~25%, Fig. 5A). The same rotenone concentration also increased H2O2 release to an extent very similar to that observed in preconditioned mitochondria (~30%, Fig. 5B).



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  		Fig. 5. Complex I inhibition increases H2O2 release. Control mitochondria (0.5 mg/ml) were added to reaction medium at 37°C containing 125 mM KCl, 0.1 mM EGTA, 10 mM HEPES, 1 mg/ml BSA, 2 mM pyruvate, 2 mM Pi [pH 7.2 (KOH)], 0.5 mM ADP, 50 µM Amplex red, and 1 U/ml HRP. Rotenone (5 nM) was added where indicated, and mitochondrial respiration (A) and H2O2 release (B) were measured as described in MATERIALS AND METHODS. Numbers in parenthesis represent respiratory and H2O2 release rates (both in nmol·min-1·mg-1). The traces are representative of three similar repetitions.

 

MitoKATP activity has a regulatory role in IP (15, 40) and can be increased by oxidation of the mitoKATP channel (34, 52). On the basis of these findings, we hypothesized that mitoKATP activity may be altered in preconditioned mitochondria due to their increased ROS release. To evaluate this possibility, we measured K+ transport in our preparations. Figure 6, A and B, shows typical light-scattering measurements of isolated mitochondria added to hypotonic media containing K+ salts. Because mitochondria take up K+, their matrix volume increases due to the concomitant uptake of water, and light scattering of the mitochondrial suspension decreases. In the presence of ATP (Fig. 6, A and B, traces a and b), both control and preconditioned mitochondria presented similar swelling rates, demonstrating that inner membrane integrity and K+ leak is equivalent in these preparations. However, we found that the swelling rate and extent in the absence of ATP (Fig. 6, A and B, traces c) or in the presence of ATP and the mitoKATP agonist diazoxide (Fig.6, A and B, traces b) was larger in preconditioned mitochondria, indicating higher ATP-sensitive K+ transport in these preparations. In a series of five repetitions under each experimental condition, we found that the ATP-sensitive change in light scattering for preconditioned mitochondria was on average more than double that of control mitochondria (Fig. 6C).



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  		Fig. 6. Preconditioning increases mitochondrial ATP-sensitive K+ (mitoKATP) transport. A and B: control and preconditioned mitochondria (0.5 mg/ml) were added to 37°C medium containing 100 mM KCl, 25 mM Pi, 100 µM EGTA, 1 mM Mg2+, 5 mM HEPES, 2 mM succinate [pH 7.4 (KOH)], 1 µg/ml oligomycin, and 5 µM cytochrome c in the presence of 0.2 mM ATP (traces a), 0.2 mM ATP and 30 mM diazoxide (traces b), or no further additions (traces c). The decrease in light scattering due to mitochondrial K+ uptake and swelling was measured as described in MATERIALS AND METHODS. C: means ± SE light-scattering decrease sensitive to ATP measured 5 min after mitochondrial addition using 5 different isolations under each experimental condition. D: Michaelis-Menten constant (Km) for K+ in control and preconditioned mitochondria was measured in six mitochondrial preparations under each experimental condition by gradually substituting K+ for Na+ in a reaction medium similar to A–C. E: Km for ATP in control and preconditioned mitochondria was measured by following light scattering in the presence of different ATP concentrations (2–2,000 µM). The results represent means ± SE of five determinations under each experimental condition using different preparations. *P < 0.05 relative to control.

 

In addition to testing the rate and extent of K+ uptake in these mitochondria, we determined transport rates in the presence of increasing K+ and ATP concentrations in preconditioned and control mitochondria, to establish whether IP changes the regulatory characteristics of this channel. By measuring light scattering rates in media in which K+ was gradually replaced by Na+ (which is not transported by mitoKATP), we established that under our conditions, both control and preconditioned mitochondria exhibit a Michaelis-Menten constant (Km) of ~60 mM for K+ transport (Fig. 6D), a value slightly higher but not considerably different from that reported previously with the use of (32 mM) reconstituted mitoKATP (36). We also determined the Km for inhibition by ATP in these mitochondria and found a similar Km of ~10–15 µM in both preconditioned and control mitochondria (Fig. 6E). Previously, the Km for ATP determined using mitoKATP reconstituted into proteolipossomes was in the 20–40 µM range (37), quite similar to our values. Together, the data shown in Fig. 6 indicate that IP enhances mitoKATP transport activity, whereas both the K+ concentration necessary for transport and the inhibitory effect of ATP remain unchanged.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we evaluated the effect of IP on mitochondrial respiration, inner and outer membrane integrity, ROS generation, and K+ transport. Our aim was to uncover mitochondrial alterations promoted by IP that could be responsible for the protective properties of this procedure on tissue damage induced by ischemia-reperfusion.

Unlike tissue-damaging ischemia (19), we found that the short ischemic periods that promote IP do not lead to inner or outer mitochondrial membrane permeabilization, as determined by measuring {Delta}{Psi} and outer membrane permeability to cytochrome c, respectively (Fig. 3, C and D). Impermeability to cytochrome c indicates that apoptotic cell death signals initiated by mitochondrial intermembrane space components are not activated during IP (21, 39). In addition, mitochondrial function involving ATP synthesis is preserved because respiratory control ratios are equal (Fig. 3B). These findings are compatible with the protective, rather than detrimental, effects of IP.

However, IP did affect mitochondrial function in a less-damaging manner. One interesting finding in our study is that complex I activity in preconditioned mitochondria was decreased by ~25% (Fig. 3A), an amount probably insufficient to lead to deleterious respiratory impairment. Previous studies (7, 8) have measured respiration supported by the NADH-linked substrate {infty}-ketoglutarate in preconditioned mitochondria and found a tendency toward lower rates compared with control mitochondria, but the differences were not significant. This lack of significance probably occurred because the measurements were conducted in the presence of low (213 µM) ADP concentrations, which are insufficient to achieve maximal respiratory rates (19). Indeed, we found that under conditions in which respiration was not maximized by high ADP concentrations (<800 µM) or the presence of 1 µM CCCP, no difference in respiratory rates could be detected between preconditioned and control mitochondria (results not shown).

A decrease in the activity of complex I in preconditioned mitochondria is not unexpected because complex I has been shown to be very susceptible to inactivation during ischemia and is an early marker of ischemic damage (40, 44). The cause for the reduction of complex I activity during ischemia is not fully understood, but may involve loss of flavin mononucleotide (42) and damage due to acidosis (41). We also found that preconditioned mitochondria generate between 30% and 130% more H2O2 than control mitochondria, depending on the respiratory state (Fig. 4). This increased H2O2 release is caused by respiratory inhibition in preconditioned mitochondria, because the presence of respiratory inhibitors myxothiazol and antimycin A in the presence of NADH-linked substrates completely eliminates the differences in H2O2 release levels. Indeed, the higher H2O2 release rates due to preconditioning are not observed when mitochondria are energized with succinate plus rotenone and can be reproduced in control mitochondria by a partial inhibition of complex I promoted by rotenone (Fig. 5), suggesting that this effect is linked to the complex I inhibition observed in Fig. 3A. Complex I is an important respiratory chain site for the generation of superoxide radicals, and the inhibition of this complex increases mitochondrial ROS release because electrons accumulated within the NADH dehydrogenase are less capable of being transferred to coenzyme Q and have a higher probability of reducing oxygen to superoxide radical anions (44). Interestingly, we found no difference in ROS release levels at complex III, a major site for ROS production (see Fig. 4B), in our preconditioned mitochondria.

Our finding that preconditioned mitochondria generate higher levels of H2O2 is in agreement with literature data showing an increase in ROS release during IP (3, 46). This mild state of oxidative stress is an important signal in IP because it prevents the large increase in ROS release observed after ischemia-reperfusion (3, 45). Indeed, antioxidants can prevent the beneficial effects of IP (26, 46).

ROS produced during IP can enhance mitoKATP activity either through direct oxidation (34, 52) or by increasing kinase activity and mitoKATP phosphorylation (13, 15). In fact, our finding that preconditioned mitochondria exhibit an increased rate of ATP-sensitive, diazoxide-stimulated, K+ uptake (Fig. 6, A–C) provides direct experimental evidence for mitoKATP activation during IP. Thus mitochondria in preconditioned hearts are capable of undergoing faster and more extensive volume changes, the main effect of mitoKATP opening (23). Despite an increased ability to transport K+, we found that IP does not alter mitoKATP K+ transport affinity or its inhibition by ATP (Fig. 6, D and E).

It was originally suggested that mitoKATP activation was responsible for the increase in ROS release observed during IP (6, 11, 26, 33; for a discussion, see Ref. 35). Under our conditions, mitoKATP could not be the cause of increased ROS because we observed an increased H2O2 generation when mitoKATP was closed by ADP, which has an inhibitory effect on the channel (36), or open, in the presence of high or low {Delta}{Psi} (oligomycin and CCCP, respectively), and did not observe this effect when succinate was used as a respiratory substrate (Fig. 4). In addition, we (10) recently found that mitoKATP activation in isolated mitochondria decreases, rather than increases mitochondrial H2O2 release. This was an expected result because mitoKATP activity promoted slight mitochondrial uncoupling (23), and mild mitochondrial uncoupling significantly prevents electron leakage at the respiratory chain by decreasing the lifetime of electron transport chain intermediates capable of donating electrons to oxygen (20). On the basis of these new findings, we propose that ROS release promoted by respiratory complex I inhibition (Figs. 3A and 4) occurs upstream of mitoKATP activation in IP (26, 35), promoting the opening of this channel (see Fig. 7). MitoKATP activity then leads to more efficient oxidative phosphorylation (9), lower mitochondrial ATP consumption (5, 9), the prevention of mitochondrial Ca2+ accumulation (9, 30), attenuated reperfusion ROS release (45), and other cardioprotective effects, which may include increased reactive nitrogen species generation (26).



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  		Fig. 7. Proposed sequence of events leading to ischemic protection in preconditioned hearts. Ischemic preconditioning promotes a small decrease in the efficiency of NADH oxidation by the respiratory chain, which increases mitochondrial reactive oxygen species (ROS) release. Increased ROS in preconditioned hearts promotes the activation of mitoKATP channels, which decrease reperfusion ROS. In addition, mitoKATP activation improves high-energy phosphate levels during ischemia and reperfusion. As a result of lower reperfusion, ROS levels, and larger high-energy phosphate contents, the tissue is protected against ischemic damage.

 

In summary, our study indicates that IP does not change inner or outer mitochondrial membrane integrity, but promotes a partial inhibition of NADH-supported respiration, an increase in H2O2 release, and activation of mitoKATP channels. These are the first direct measurements demonstrating that mitochondrial ROS and mitoKATP activity are enhanced after IP and are in line with experiments in cells and organs suggesting these effects. In addition, our studies uncover the inhibition of NADH-supported respiration as a novel mechanism through which ROS generation is increased in preconditioned hearts.


   ACKNOWLEDGMENTS
 
The authors thank Edson Alves Gomes for excellent technical assistance, and Prof. Roger F. Castilho for the critical reading of the paper.

This project was supported by Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) and Conselho Nacional de Desenvolvimento Científico e Tecnológico. M. M. da Silva, A. Sartori, and E. Belisle are students supported by FAPESP scholarships.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. J. Kowaltowski, Departamento de Bioquímica, IQ, USP, Av. Prof. Lineu Prestes, 748, 05508-900, São Paulo SP, Brazil (E-mail: alicia{at}iq.usp.br).

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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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