Heart mitochondria were isolated from ketamine-anesthetized (50 mg/kg ip) guinea pigs (250–350 g) as described previously (Gadicherla et al., 2012; Aldakkak et al., 2013; Blomeyer et al., 2013). Briefly, ventricles were excised, placed in an isolation buffer (buffer A) that contained (in mM) 200 mannitol, 50 sucrose, 5 KH₂PO₄, 5 MOPS, 1 EGTA, and 0.1% bovine serum albumin (BSA; all chemicals from Sigma, St. Louis, MO, USA), with pH adjusted to 7.15 with KOH. Ventricles were then minced into 1-mm³ pieces. The suspension was homogenized in isolation buffer containing 5U/ml protease (Bacillus licheniformis; Sigma), followed by differential centrifugation at 4°C, and the final pellet was resuspended in isolation buffer and kept on ice. Protein content was determined by the Bradford method. Mitochondrial suspension was adjusted to yield 12.5 mg protein/ml for experimental purpose. Details of the experimental approach are provided in the Supplementary Material.

Experiments were conducted at room temperature (25°C), with mitochondria (0.5 mg protein/ml) suspended in experimental buffer (buffer B) that contained (in mM) 130 KCl (EMD Chemicals, Gibbs-town, NJ, USA), 5 K₂HPO₄, 20 MOPS, 0.001 Na₄P₂O₇, and 0.1% BSA. This assured that only 40 μM EGTA was carried over from the isolation buffer (buffer A) into the experimental buffer. Based on the experimental protocol and conditions, the buffer pH was specifically adjusted upward from 6.5 to 6.9 and 7.15 by adding KOH. The respiration buffer contained 0 or 150 μM CaCl₂; concentrations of CaCl₂ between 20 and 60 μM had no significant effects on H₂O₂ production (Figure S.3) and 100 μM CaCl₂ gave inconsistent data, so these data are not reported. From the residual EGTA concentration of 40 μM, we estimated 150 μM CaCl₂ to be equivalent to ≈ 220 nmol CaCl₂/mg protein. After adding CaCl₂ (or H₂O), 10 mM Na⁺ pyruvate or Na⁺ succinate (Sigma) was added. Then either complex I blocker ROT (10 μM; Sigma) or complex III blocker AA (5 μM; Sigma) was added.

Mitochondria were suspended in buffer B in a 1 ml cuvette inside a spectrophotometer (QM-8; Photon Technology International (PTI), Birmingham, NJ, USA). The rate of H₂O₂ release was measured using Amplex red (12.5 μM; Molecular Probes, Eugene, OR, USA) and horseradish peroxidase (0.1 U/ml; Sigma) at excitation and emission wavelengths of 530 and 583 nm, respectively. H₂O₂ is the direct product of O^•−₂ when catalyzed by O^•−₂ dismutase (SOD) in the absence of nitric oxide. H₂O₂ levels were calibrated over a range of 10–200 nM H₂O₂ (Sigma) added to buffer B in the absence of mitochondria and in the presence of Amplex red and horseradish peroxidase. Mitochondrial volume change (increase/decrease) was assessed by monitoring changes in 90° light scattering at an excitation and emission wavelength of 520 nm inside the same cuvette-based PTI.

Oxygen consumption was measured polarographically using a respirometry system (System S 200A; Strathkelvin Instruments, Glasgow, Scotland). Respiration experiments using pyruvate or succinate at pH 7.15 and without Ca²⁺ were initially conducted to determine the viability of mitochondria for the rest of the experiments. Respiration was initiated by adding 10 mM complex I substrate Na⁺ pyruvate or the complex II substrate Na⁺ succinate. State 3 respiration was measured after adding 250 μM ADP (Sigma), and state 4 respiration was measured after complete phosphorylation of the added ADP. The respiratory control index (RCI) was calculated as the ratio of the rate of state 3 to state 4 respiration. Only mitochondria with an RCI of 10 or above with pyruvate or an RCI of 3 or above with succinate were used in the experiments. To assess the effects of pH and extra-matrix (e) [Ca²⁺]_e on O₂ consumption (respiration), we added either H₂O (control) or CaCl₂ for a final concentration of 150 μM to the mitochondrial suspension at three pHs (7.15, 6.9 or 6.5) before adding substrates.

We first evaluated H₂O₂ release rates resulting from ROT-inhibited complex I in pyruvate-energized mitochondria at pHs 7.15, 6.9, and 6.5, each with H₂O (control) or with added 150 μM CaCl₂ (Figures 1A–C, 2). In the absence of CaCl₂, adding ROT caused a modest increase in H₂O₂ release rate at each pH. In the presence of 150 μM CaCl₂, adding ROT caused a marked increase in H₂O₂ release rate at pH 6.9 (1.38 ± 0.12 pmol/mg/s) compared to pH 7.15 (0.37 ± 0.02 pmol/mg/s) or to pH 6.5 (0.28 ± 0.07 pmol/mg/s). Since high Ca²⁺ is known to induce mPTP opening (Camara et al., 2010), we tested the hypothesis that the Ca²⁺-induced H₂O₂ increase is a result of mPTP opening. Addition of cyclosporine A (CsA; 0.5 μM; Sigma) prevented the large increase in H₂O₂ release, which was more apparent at pH 6.9 (0.39 ± 0.04 pmol/mg/s). To further evaluate the role of mPTP, we measured the corresponding mitochondrial volume changes in pyruvate-energized mitochondria in the same combinations of buffer pH and CaCl₂ with later addition of ROT (Figures 3A–C). Adding CaCl₂ alone did not significantly alter mitochondrial volume before addition of pyruvate. However, at pH 6.9 (Figure 3B) and 6.5 (Figure 3C), mitochondrial volume significantly increased with added 150 μM CaCl₂ after adding pyruvate, but volume was not significantly affected at pH 7.15 (Figure 3A). Additionally, mitochondrial volume did not change significantly in experiments without added CaCl₂. Adding CsA prevented the large increases in mitochondrial volume; adding ROT stopped any increases in volume (Figures 3A–C). The effect of adding superoxide SOD on H₂O₂ release under these conditions is given in Supplementary Materials.

We evaluated H₂O₂ release rates resulting from adding AA to succinate-energized mitochondria at pH 7.15, 6.9, or 6.5, without (control) or with added 150 μM CaCl₂ (Figures 4A–C, 5). In the absence of CaCl₂ at all pHs tested, adding succinate prior to adding AA caused an increase in H₂O₂ release rate while later addition of AA reduced succinate-induced H₂O₂ release rate (Figures 4A–C). In the presence of 150 μM CaCl₂ without AA, adding succinate did not significantly increase H₂O₂ release. However, later addition of AA with added CaCl₂ (150 μM) caused a large increase in H₂O₂ release rate at all pHs with pH 7.15 showing the highest rate (35.2 ± 1.0 pmol/mg/s), followed by pH 6.9 (32.6 ± 0.8 pmol/mg/s), and then by pH 6.5 (23.7 ± 1.6 pmol/mg/s) (Figures 4A–C, 5). Adding CsA prevented the Ca²⁺-induced increase in H₂O₂ release rate resulting from AA treatment at each pH with the least effect on mPTP at pH 7.15. Because enzyme activity is pH dependent, we examined the effect of adding SOD to the buffer under the same experimental conditions. The effect of adding SOD on H₂O₂release was minimal (Figures S.1, S.2). In parallel, we measured the corresponding mitochondrial volume changes in succinate-energized mitochondria in the same combinations of pH and extra-matrix CaCl₂ with later addition of AA (Figures 6A–C). Adding CaCl₂ alone did not significantly alter mitochondrial volume before adding succinate (Figures 6A–C). Later addition of succinate induced a significant increase in mitochondrial volume at all pHs which was prevented by CsA. Adding AA led to an attenuation of mitochondrial volume (Figures 6A–C).

Because O^•−₂ generation is dependent on electron flux through the ETC and is a product of electron leak at several complexes, we evaluated the effects of substrate, pH, and [Ca²⁺]_e on mitochondrial respiration during states 2, 3, and 4 respiration (all of the above experiments were conducted during state 2). In pyruvate-energized mitochondria (Figure 7A), and before adding CaCl₂, there was no difference in state 2 respiration among all pH groups. Adding 150 μM CaCl₂ led to an increase in state 2 in the pH 7.15 group only. In succinate-energized mitochondria (Figure 7B), and before adding CaCl₂, there was no difference in state 2 respiration among all pH groups. However, adding 150 μM CaCl₂ led to similar decreases in state 2 respiration for all pH groups. States 3 and 4 respiration rates and the respective RCI values for each pH group without (control) or with addition of 150 μM CaCl₂ are summarized (Table 1). Adding 150 μM CaCl₂ decreased states 3 and 4 respiration and RCI under each of the two substrate and three pH conditions, except at pH 7.15 for state 4 respiration with pyruvate where state 4 respiration increased slightly but significantly. State 3 respiration and RCI were unaffected by pH except in two conditions: (1) pyruvate and 0 μM CaCl₂, in which they were reduced at pH 6.5, and (2) pyruvate and 150 μM CaCl₂, in which they were reduced at pH 6.9 and 6.5. State 4 respiration was unaffected by pH except in the condition pyruvate and 150 μM CaCl₂, in which it was reduced at pH 6.9 and 6.5.

It is well-documented that reperfusion after ischemia is associated with considerable ROS emission (Vanden Hoek et al., 1997; Becker et al., 1999; Kevin et al., 2003). It appears paradoxical that ROS are also produced during ischemia; but total mitochondrial anoxia is unlikely to exist even with extensive cardiac ischemia (Becker, 2004). It has been reported that isolated mitochondria produce H₂O₂ when O₂ is as low as 0.5 mM (pO₂ of about 10 mmHg) (Saborido et al., 2005). Mitochondrial pO₂ in vivo is only about 1–5 mmHg with normoxia, so mitochondria normally thrive in a low O₂ environment. The emission of ROS is due to excessive O^•−₂ generation (likely during early mild ischemia when the redox potential is high) and to diminished ROS scavenging (likely during later severe ischemia when the redox potential is low).

In previous studies of IR injury in guinea pig isolated hearts, we observed a modest increase in O^•−₂ generation during early ischemia (10–15 min) followed by a larger increase in O^•−₂ generation during late ischemia (20–30 min) and a surge during early reperfusion (Kevin et al., 2003; Riess et al., 2004; Aldakkak et al., 2008a,b, 2011). Recently, we demonstrated in isolated mitochondria, under conditions simulating ischemia, that CaCl₂ addition in the presence of succinate resulted in enhanced H₂O₂ release when complex III was blocked (Aldakkak et al., 2013).

It is widely recognized that both mitochondrial complexes I and III play a crucial role in producing ROS during cardiac IR injury, particularly with induction of mPTP. “Triggering” amounts of ROS can be generated in isolated cardiomyocytes by photoactivation of tetramethylrhodamine derivatives that leads to membrane depolarization (mPTP induction) and a burst of ROS, which is coined “ROS-induced ROS release (RIRR)” (Zorov et al., 2000). However, inducing mPTP opening with excess CaCl₂ in isolated liver mitochondria did not increase H₂O₂ release unless exogenous NADH was added to the buffer (Batandier et al., 2004). Because mPTP opening resulted in a ROT-sensitive impairment of complex I activity, they concluded that mPTP opening is associated with defective electron transfer within complex I, leading to O^•−₂ release at that site. We did not add NADH to the buffer but we observed an increase in H₂O₂ release under either substrate condition when complex I and III inhibitors were used to mimic some conditions of IR injury. Despite these advances, the prevailing metabolic conditions during mild vs. severe IR injury that promote differential dysfunction of the complexes to cause excess O^•−₂ generation and ROS emission in a vicious cycle of RIRR remain unclear.

In ischemia, decreasing O₂ levels and concomitant activation of glycolysis caused a decrease in pH, increased extra-matrix and matrix [Ca²⁺], and eventually damage to complexes I and III due to oxidative stress (Rouslin, 1983). O^•−₂ generated under these conditions is derived in part from complex I through forward electron transfer (FET) (Starkov et al., 2004). In the study above (Rouslin, 1983), it was reported that the activity of complex I decreased markedly after 20 min ischemia and that this decrease closely paralleled the decrease in mitochondrial O₂ uptake with NADH-linked substrates; it was also reported that the activity of complex III decreased at a more gradual rate during ischemia and that its rate of decrease paralleled that of succinate-supported O₂ uptake. As ischemia progresses, pH drops further and extra-matrix and matrix Ca²⁺ levels rise, while substrate utilization switches from primarily pyruvate to mostly succinate, which accumulates from 0.2 to 0.4 mM during normoxia to 4–7 mM during ischemia or hypoxia (Kakinuma et al., 1994; Starkov et al., 2004). Based on these studies, we used pyruvate + ROT to approximate the condition of impaired electron transfer via complex I to complex III, and succinate + AA to approximate the condition of enhanced utilization of succinate at complex II and impaired electron transfer through complex I (by RET) and III. Our protocols were also based on a summary of data (Lukyanova, 2013) stating that the switch from normoxia to hypoxia increased succinate utilization by complex II from about 25–35% to 65–85% while complex I activity was mostly inhibited. Under this condition, complex III appears to be damaged by ROS due to the initial O^•−₂ generated at complex I, which is analogous to AA inhibition of complex III (Musatov and Robinson, 2012). Given these conditions and the diminished ΔΨ_m, ROS production during severe IR injury likely results from FET including complex II and III. Therefore, our experiments with succinate + AA may mimic the conditions of O^•−₂ generated at both complex I and III during IR injury.

Adding CaCl₂ and lowering pH both modulated H₂O₂ production in pyruvate-energized mitochondria after adding ROT (Figures 1A–C, 2). Based on previous studies showing a distinct early phase of H₂O₂ production during mild ischemia correlated with the timing of complex I damage, we hypothesized that pyruvate-energized mitochondria with inhibited complex I and added CaCl₂ would show an increased H₂O₂ release rate as buffer pH decreased. Indeed, a sharp rise in H₂O₂ production was observed at pH 6.9 on addition of CaCl₂, which was not seen at pH 7.15. However, as pH decreased further to 6.5, a rise in H₂O₂ did not occur. It is unclear what the exact mechanism is for the large increase in H₂O₂ release in pyruvate-supported mitochondria at pH 6.9 with added CaCl₂. It is important to note that under these conditions, there was a significant increase in mitochondrial volume, probably indicating mPTP opening. Indeed, addition of CsA to desensitize mPTP prevented the increase in mitochondrial volume and reduced H₂O₂ release to levels similar to those observed at pH 7.15. Nonetheless, mPTP opening occurred also at pH 6.5 with high CaCl₂ as indicated by the increase in mitochondrial volume that was prevented with CsA; but this was not associated with a large increase in H₂O₂ release.

mPTP opening can occur during mitochondrial Ca²⁺ overload (Orrenius et al., 2003), whereas a low mitochondrial pH is associated with a reduced probability of mPTP opening. For example, reoxygenation or reperfusion under acidic conditions is associated with much lower ROS emission (Haworth and Hunter, 1979; Halestrap, 1991; Bernardi et al., 1992). However, a previous study (Halestrap, 1991), with glutamate/malate-energized mitochondria isolated from rat hearts, showed a graded effect of pH on the probability of mPTP opening with the least likelihood of opening at pH 6.0 and below. At pHs above 6.0 there was a significant increase in the probability of mPTP opening. Indeed, at pH 6.5, a Ca²⁺ -induced increase in volume, presumably through mPTP opening, was demonstrated to be greater than 40% of the increase seen at pH 7.4; and pH 6.9 had approximately 75% of the volume increase seen at pH 7.4 (Halestrap, 1991).

mPTP opening is proposed to increase ROS emission through three mechanisms: a loss of glutathione leading to decreased ROS scavenging, a loss of cytochrome c leading to increased reduction of upstream ETC complexes and subsequent electron loss and diminished scavenging, and an increase in ROS derived from the α-ketoglutarate dehydrogenase complex due to loss of NAD⁺ (Camara et al., 2010, 2011; Toledo et al., 2014). In our study different probabilities of mPTP opening at pH 6.9 and 6.5 may be responsible for the disparity in H₂O₂ levels generated at these two pHs. Limited opening of mPTP at pH 6.5 may be responsible for H₂O₂ generation at levels not sufficient to induce RIRR, whereas at pH 6.9 the increased opening of mPTP may induce RIRR. Additionally, the lack of increased H₂O₂ generation and mitochondrial volume at pH 7.15, when compared to pH 6.9, might be related to an increase in inhibition of complex I with high CaCl₂ as pH increases (Sadek et al., 2004; Chen et al., 2010). The decrease in complex I activity leading to decreased ROS production might in turn prevent RIRR and subsequent opening of the mPTP (Zorov et al., 2006). Thus, the combined effects of inhibited complex I activity and mPTP opening at pH 6.9 might explain the elevation in H₂O₂ release rates observed at pH 6.9, but not at pHs 7.15 and 6.5.

Adding CaCl₂ and altering pH also modulated H₂O₂ generation in succinate-energized mitochondria with added AA (Figures 4A–C). With AA, mitochondria in buffer with high CaCl₂ showed a dramatic rise in H₂O₂ release at all pHs. The increase in H₂O₂ corresponded to an increase in pH. In addition, an increase in mitochondrial volume occurred at each pH in the presence of high CaCl₂, and these conditions led to increased H₂O₂ release, suggesting a role for mPTP opening. Indeed, both the increases in H₂O₂ release and volume were inhibited by adding CsA at each pH. H₂O₂ generation under these conditions is likely caused by FET from complex II through complex III, because adding AA decreased ΔΨ_m, which would prevent RET from occurring. In this case, the primary source of H₂O₂ is that derived from O^•−₂ generated at the Q_o site of complex III.

Mitochondrial pH may have a significant role in moderating O^•−₂ generation by complex III. Matrix alkalinization (higher pH) tended to stabilize the semiquinone radical at the Q_o site (Selivanov et al., 2008). This was proposed to result from decreased binding of H⁺ ions necessary to drive the Q cycle forward (Selivanov et al., 2008). Stability of the semiquinone radical leads to increased likelihood of direct transfer of an electron to an O₂ molecule, leading to the formation of O^•−₂ (Selivanov et al., 2008). Additionally, because AA blocks the oxidation of semiquinone at Q_i and the transfer of an electron from the Q_o site, this can lead to increased O^•−₂ generation from the Q_o site, which may be analogous to impaired complex III function during ischemia (Chen et al., 2008; Musatov and Robinson, 2012). Consequently, in our experiments, the increased H₂O₂ release rate at a high pH is possibly related to the increased stability of semiquinone leading to increased direct electron donation to O₂ to generate O^•−₂.

Mitochondrial uncouplers like dinitrophenol tend to increase respiration to counteract a decline in ΔΨ_m due to H⁺ leak. But others have reported that it is not unusual for mitochondrial uncouplers or uncoupling events such as mPTP opening to inhibit succinate-supported state 2 respiration. Mitochondrial uncouplers can retard succinate oxidation under some conditions (Papa et al., 1969). In the absence of ROT, and with succinate in high concentrations (conditions similar to those used in our study), mitochondrial uncouplers have been found to inhibit succinate oxidation due to the formation of oxaloacetate (Wojtczak et al., 1969; Vik and Hatefi, 1981; Kotlyar and Vinogradov, 1984; Drose, 2013).

In our study, the lower state 2 respiration after adding CaCl₂ with succinate (Figure 7B) at each pH, may be due to a greater collapse in ΔΨ_m due to the excess influx of Ca²⁺. In contrast, the higher state 2 respiration after adding CaCl₂ with pyruvate (Figure 7A) at pH 7.15 may be a result of enhanced H⁺ pumping at complex I by this NADH-linked substrate; however, as the trans-membrane pH potential is increased (and thus the proton motive force), a faster respiration might not be needed to maintain ΔΨ_m. The RCI for pyruvate was reduced at pH 6.5 likely because of an uncoupling effect due to H⁺ leak with slower ATP production (Table 1). The RCI for succinate, which is much lower than that for pyruvate, appears to stem from the much higher basal respiratory rate for succinate vs. pyruvate (Figures 7B vs. 7A). In the absence of CaCl₂, mitochondria showed well-coupled oxidative phosphorylation with both substrates at each pH, except in pyruvate-energized mitochondria at pH 6.5, in which case the RCI was lower, indicating relatively less coupling. The effect of added CaCl₂ on states 3 and 4 respirations and RCI with either substrate or at any pH is likely a result of marked uncoupling due to mPTP opening (increased mitochondrial volume) because this was sensitive to CsA (Figures 3, 6).

In conclusion, in our previous studies using the isolated, beating heart model of 30 min global ischemia, we demonstrated two phases of increased O^•−₂ generation, an early phase (10–20 min) that emits low to moderate O^•−₂ levels and a late phase (20–30 min) that emits higher O^•−₂ levels just before a surge in O^•−₂ release at the beginning of reperfusion. The present study sheds novel insights into the modulatory effect of matrix pH in Ca²⁺-induced mitochondrial H₂O₂ release. The early or mild phase of H₂O₂ release due to O^•−₂ generation at complex I could be related to differential effects of pH on the mPTP, which allows H₂O₂ production at the pH observed during early or mild ischemia (pH 6.9) but not at the lower or higher pHs. The late or severe phase of H₂O₂ release due to O^•−₂ generation primarily at complex III, but also at complex I, may also be dependent on mPTP opening, but H₂O₂ production is intensified with increasing pH. Therefore, it is possible that the surge in H₂O₂ production commonly observed on reperfusion results from O^•−₂ generated from complex III as the pH rises gradually with mPTP opening. Although both complex I and III contribute simultaneously to H₂O₂ production during IR, our results suggest that the role of each respiratory complex is not static but rather changes dynamically as the pH changes. Thus, each complex may play a more prominent role during a certain period of IR. Being cognizant of this information is important as it can be used to reduce ROS emission as ischemia progresses by targeting each complex separately, or possibly by manipulating the pH using Na⁺/Ca²⁺ and/or Na⁺/H⁺ exchange inhibitors, e.g., by maintaining a more alkaline environment during early ischemia and a more acidic environment during late ischemia/early reperfusion, to reduce O^•−₂ generation at complex I and III.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.