AJP - Heart Calcium Transients and Cell-Sarcomere
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Am J Physiol Heart Circ Physiol 280: H2203-H2213, 2001;
0363-6135/01 $5.00
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Vol. 280, Issue 5, H2203-H2213, May 2001

Phenylarsine oxide induces mitochondrial permeability transition, hypercontracture, and cardiac cell death

Paavo Korge, Joshua I. Goldhaber, and James N. Weiss

Cardiovascular Research Laboratory, Departments of Medicine (Cardiology) and Physiology, University of California at Los Angeles School of Medicine, Los Angeles, California 90095-1760


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The mitochondrial permeability transition (MPT) is implicated in cardiac reperfusion/reoxygenation injury. In isolated ventricular myocytes, the sulfhydryl (SH) group modifier and MPT inducer phenylarsine oxide (PAO) caused MPT, severe hypercontracture, and irreversible membrane injury associated with increased cytoplasmic free [Ca2+]. Removal of extracellular Ca2+ or depletion of nonmitochondrial Ca2+ pools did not prevent these effects, whereas the MPT inhibitor cyclosporin A was partially protective and the SH-reducing agent dithiothreitol fully protective. In permeabilized myocytes, PAO caused hypercontracture at much lower free [Ca2+] than in its absence. Thus PAO induced hypercontracture by both increasing myofibrillar Ca2+ sensitivity and promoting mitochondrial Ca2+ efflux during MPT. Hypercontracture did not directly cause irreversible membrane injury because lactate dehydrogenase (LDH) release was not prevented by abolishing hypercontracture with 2,3-butanedione monoxime. However, loading myocytes with the membrane-permeable Ca2+ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-acetoxymethyl ester (BAPTA-AM) prevented PAO-induced LDH release, thus implicating the PAO-induced rise in cytoplasmic [Ca2+] as obligatory for irreversible membrane injury. In conclusion, PAO induces MPT and enhanced susceptibility to hypercontracture in isolated cardiac myocytes, both key features also implicated in cardiac reperfusion and reoxygenation injury.

dithiothreitol; myofibrillar Ca2+ sensitivity; mitochondrial Ca2+ efflux


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

IRREVERSIBLE REPERFUSION/REOXYGENATION injury is closely connected with the mitochondrial permeability transition (MPT), as Crompton and co-workers suggested in the late 1980s (see Refs. 5, 17, and 24 for reviews). MPT is caused by the opening of permeability transition pores (PTP) in the inner mitochondrial membrane, causing collapse of membrane potential, mitochondrial Ca2+ efflux, equilibration of solutes with molecular mass <1,500 Da across the inner membrane, and the inability of the mitochondria to generate ATP (5). Available experimental evidence suggests that PTP remain closed during ischemia but open during reperfusion (15). Hypercontracture is another hallmark of reperfusion/reoxygenation in cardiac muscle that exacerbates injury by disrupting membrane integrity. Hypercontracture requires both elevated intracellular [Ca2+] and ATP availability for cross-bridge cycling (see Ref. 30 for a review). However, relative to the elevated levels of cytoplasmic [Ca2+] during ischemia/hypoxia, the lack of a further large increase in cytoplasmic [Ca2+] on reperfusion/reoxygenation (16, 26) suggests that reperfusion/reoxygenation increases the susceptibility of cardiomyocytes to hypercontracture (30). This is supported by the observation that cells develop Ca2+-induced hypercontracture at significantly lower cytosolic Ca2+ levels after hypoxia/reoxygenation than during normoxia (23).

The hydrophobic sulfhydryl group reagent phenylarsine oxide (PAO), which specifically cross-links vicinal sulfhydryl groups (36), has been found to be a potent inducer of MPT in isolated mitochondria (2). To further investigate the role of mitochondrial permeability transition (MPT) in cardiac injury, we examined the effects of PAO in isolated ventricular myocytes. We show that PAO induces MPT and causes both severe hypercontracture and irreversible membrane injury. The mechanisms underlying these effects are explored.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Isolated Ventricular Myocyte Studies

Isolation procedure. Myocytes were isolated from adult rabbit hearts by the conventional enzymatic method described previously (25a). Cells were stored in normal Tyrode solution and used within 5-6 h.

Confocal microscopy. Single myocytes loaded with fluorescent probe(s) were imaged with an Odyssey XL laser-scanning confocal microscopy system (Noran Instruments; Middleton, WI) attached to a Ziess Axiovert TV100 fitted with a ×40 water immersion objective lens (model 40/1.2 W Corr; Apochromat). Tetramethyl rhodamine methyl ester (TMRM) and rhodamine 2 (rhod 2) were loaded at room temperature and calcein was loaded as described below. The red fluorescence of TMRM and rhod 2 were excited with the use of the 568-nm line of an argon laser, and emission was collected above 590 nm through a long-pass filter. Calcein fluorescence was excited at 488 nm and imaged through a 515-nm band-pass filter.

Selective loading of calcein into mitochondria. In isolated myocytes, MPT was evaluated by selectively loading mitochondria with the fluorescent dye calcein (molecular mass 623 Da). We modified the method developed recently by Petronilli et al. (29), who used mitochondrial calcein efflux to detect MPT in liver cells by quenching cytosolic and nuclear calcein fluorescence with Co2+. Once inside the mitochondrial matrix, calcein is trapped unless MPT occurs and allows calcein to efflux through PTP (molecular mass cutoff ~1,500 Da in the high-conductance mode of MPT). MPT is therefore detected in the confocal image of the myocyte when the discrete striped mitochondrial pattern of calcein fluoresence is lost and replaced by a diffuse fluorescence pattern.

To selectively load mitochondria with calcein, myocytes were incubated in Tyrode buffer containing 10 µM calcein-acetoxymethyl ester (AM) for 60 min at 4°C. After 60 min, cells were allowed to sediment at the bottom of the test tube at 4°C, the buffer was removed, and cells were resuspended in buffer without calcein. The cells were then incubated at 37°C for 3-4 h, during which time the buffer was changed three to four times. This incubation was necessary to selectively visualize the calcein fluorescence originating from mitochondria, leading us to hypothesize that during the warm incubation, calcein leaked out from the cytoplasm but was retained in mitochondria. Preferential loss of cytoplasmic compared with mitochondrial fluorophores during warm incubation of myocytes has been noted by others (35). The efficiency of cytoplasmic calcein removal was tested by periodically examining a sample of myocytes under the confocal microscope during warm incubation.

Preferential mitochondrial distribution of calcein was confirmed by its colocalization with TMRM (Fig. 1). Isolated myocytes were first loaded with calcein as described and imaged under the confocal microscope (Fig. 1A). TMRM (100 nm) was then added, and the same cell was imaged for the red fluorescence of TMRM (Fig. 1B). Dual fluorescence images obtained at 515 nm (for calcein) and at >590 nm (for TMRM) demonstrated nearly identical patterns. Mitochondrial localization of calcein was also supported by its fluorescence colocalization with another mitochondrial-specific probe, MitoTracker Green (data not shown). Compared with the method of Petronelli et al. (29), this technique has the advantage of not requiring intracellular Co2+ loading to quench the cytoplasmic calcein signal.


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  		Fig. 1.   Selective labeling of mitochondria in situ with calcein. A: confocal image of a myocyte loaded with calcein-acetoxymethyl ester (AM) to selectively label mitochondria (see MATERIALS AND METHODS), showing the classic striped sarcomeric fluorescence pattern of the mitochondrial network. B: same myocyte loaded with 100 nM tetramethyl rhodamine methyl ester (TMRM), a standard potentiometric mitochondrial imaging dye, gives a nearly identical pattern.

Measurement of free cytoplasmic Ca2+ and Mg2+. To measure intracellular Ca2+, isolated ventricular myocytes were loaded with 5 µM fluo 3-AM for 60 min at room temperature in normal Tyrode buffer. After the cells were washed, 100 µl of suspension were added to a fluorometer cuvette containing 2 ml of nominally Ca2+-free Tyrode buffer. This suspension was continuously stirred and excited by 470-nm blue light-emitting diode through a 40-nm band-pass filter at 470 nm. Fluorescence was measured with a fiber-optic spectrofluorometer (Ocean Optics) at 530 nm. Intracellular [Mg2+] was measured under similar conditions after isolated cells were loaded with 10 µM Mg2+ green-AM in the presence of pluronic. This method basically followed that described by Leyssens et al. (25), who demonstrated a close inverse correlation between cellular [ATP] decrease and [Mg2+] increase, validating that [Mg2+] changes during metabolic inhibition reflect [ATP] changes reasonably well. Because of uncertainties in calibration (25), changes in Mg2+ green fluorescence and fluo 3 fluorescence are presented in arbitrary fluorescence units.

Permeabilized myocyte experiments. The sarcolemma was permeabilized by treating cells with digitonin (20 µM) in a buffer containing (in mM) 130 KCl, 1 MgCl2, 3 ATP, 0.5 EGTA, and 10 HEPES (pH 7.2) for 10 min. The cells were pelleted at 50 g, washed, and resuspended in the same buffer (1). After this treatment was completed, the cells retained the rod shape morphology. Free [Ca2+] was calculated with the use of Winmax version 2.05 software. Cell length was recorded by using a video edge detector, as described previously (37).

Isolated Mitochondria Studies

Isolation of mitochondria. Mitochondria were isolated from adult rabbit hearts by enzymatic digestion of finely minced tissue with the bacterial protease nagarase (0.5 mg/ml) for 10 min on ice in a homogenization buffer composed of (in mM) 250 sucrose, 1 EGTA, and 10 3-(N-morpholino)propanesulfonic acid (pH 7.4 with Tris) and differential centrifugation, as described previously (22). Mitochondria were resuspended in the EGTA-free homogenization buffer to give ~40 mg/ml of mitochondrial protein, kept on ice, and used within 6 h after isolation. The respiratory control ratio, determined in mitochondrial incubation buffer composed of (in mM) 120 KCl, 10 HEPES (pH 7.4), and 2 potassium phosphate to which 2.5 mM of pyruvate, malate, and glutamate and 0.5 mM of ADP were consecutively added, was >5, suggesting that isolated mitochondria were well coupled.

Evaluation of mitochondrial matrix swelling and membrane potential. The rate of change of mitochondrial matrix volume was estimated by measuring 90° light scattering with the use of a spectrofluorometer with excitation and emission wavelengths set at 520 nm, similar to the method described by Haworth and Hunter (18) to measure MPT by mitochondrial matrix volume changes. The membrane potential (Delta Psi ) was measured in parallel with matrix volume changes by transmembrane distribution of TMRM. Mitochondria (0.5-1.0 mg) were added to 2 ml of mitochondrial incubation buffer containing 400 nM TMRM, and continuously stirred, and the effect of PAO alone or in combination with other chemicals on light scattering or TMRM fluorescence emission (580 nm) was recorded.

Mitochondrial Ca2+ uptake and efflux determination. Mitochondria (0.5-1 mg/ml) were incubated in the buffer described above containing 1 µM Ca2+ green 5N. The suspension was continuously stirred in the fluorometer cuvette, and changes in extramitochondrial [Ca2+] were followed by recording Ca2+ green fluorescence as described above for fluo 3. Calibration of the signal was achieved by the addition of known amounts of Ca2+ to incubation buffer as described previously (21) except that the buffer also contained 1 mg/ml of bovine serum albumin.

Myosin ATPase Activity

Actin-activated heavy meromyosin (HMM) ATPase activity was determined by measuring the rate of nicotinamide adenine dinucleotide (NADH) oxidation, proportional to the rate of ADP production, in a buffer containing 1 mM MgCl2, 0.5 mM EGTA, 10 mM phosphoenolpyruvate, 10 U/ml pyruvate kinase, 15 U/ml lactate dehydrogenase (LDH), 0.5 mM NADH, and 10 mM HEPES; pH 7.4 at 25°C. HMM and F-actin (10 µM) were added first, followed by ATP. In control experiments, PAO had no effect on enzymes used to determine ATPase activity.

LDH Release

Cells were allowed to sediment on Falcon dishes, and the supernatant was changed with the fresh buffer several times. The cells, which were loosely attached to the bottom, were incubated at room temperature with or without PAO and other additions, and 150 µl of supernatant was carefully removed after the indicated time periods for LDH determination. LDH activity was determined by oxidation of NADH in potassium phosphate buffer, pH 7.65, containing 1 mM of sodium pyruvate. LDH release during metabolic inhibition was evaluated relative to the total cellular LDH content (31).

All of the experiments were conducted at room temperature (23-25°C). All of the tracings or fluorescence images are representative of experiments from at least three different isolated myocyte preparations. When appropriate, original tracings were analyzed and the results presented as means ± SD.

Chemicals

Cyclosporin A (CSA) was a gift from Ciba-Geigy. HMM and actin, isolated from the rabbit heart, were gifts from Dr. E. Homsher. Percoll was purchased from Pharmacia, and all of the fluorescent dyes were purchased from Molecular Probes. All of the other chemicals were purchased from Sigma.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

PAO Induces MPT and Hypercontracture in Isolated Ventricular Myocytes

To image MPT in intact isolated myocytes, mitochondria inside myocytes were selectively loaded with calcein (see MATERIALS AND METHODS) and imaged with confocal microscopy before and during exposure to PAO. PAO (20 µM) caused the striped pattern typical of intramitochondrial localization of calcein to be progressively replaced by a more diffuse fluorescence, indicating calcein efflux from mitochondria to cytoplasm as MPT occurred (Fig. 2; n = 5). Coincident with the diffuse fluorescence pattern, the myocyte underwent progressive shortening leading to hypercontracture. The effects of PAO were dose dependent (10-50 µM), and at 20 µM typically began after a lag period of 2-4 min (n = 5). Once shortening began, the myocyte changed from a rod shape to a rounded shape within 2-3 min (see Fig. 2), and its surface became covered with membrane blebs.


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  		Fig. 2.   Phenylarsine oxide (PAO) induces mitochondrial permeability transition (MPT) in isolated cardiac myocytes. Left: successive confocal images of a myocyte loaded with calcein to image its mitochondria during exposure to 20 µM PAO. The striped mitochondrial pattern disappeared as calcein moved from mitochondrial matrix to cytoplasm during MPT, with progressive hypercontracture occurring concomitantly. Images, from top to bottom, are before, 5, 7, and 10 min after PAO, respectively. The bath superfusate was Ca2+ free. Right: identical protocol with 2,3-butanedione monoxime (BDM; 30 mM) present to prevent hypercontracture. Images, from top to bottom, are before, 5, 9, and 20 min after PAO, respectively.

Pretreating myocytes with the cross-bridge uncoupler 2,3-butanedione monoxime (BDM; 30 mM) (33) prevented cell shortening during PAO exposure yet did not prevent the loss of the striped fluorescence pattern, although it developed more slowly (n = 3; Fig. 2). Thus the change in calcein fluorescence pattern was not an artifact of severe cell shortening.

PAO-Induced Hypercontracture Requires Intracellular Ca2+ and ATP

In isolated myocytes loaded with fluo 3-AM or rhod 2-AM, PAO caused a modest increase in cytosolic Ca2+ over a similar time course as hypercontracture (Fig. 3, A and B, confirmed in five cell suspensions). When extracellular Ca2+ was replaced with EGTA, hypercontracture still developed, albeit at a modestly slower rate (Fig. 4, A and B). However, preloading myocytes with 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-AM (BAPTA-AM; a membrane-permeable, highly specific Ca2+-chelating reagent) completely prevented hypercontracture (Fig. 4A, trace c). Figure 5A shows that the sulfhydryl-reducing agent dithiothreitol (DTT) reversed the effects of PAO when applied within a few minutes of the onset of hypercontracture. These traces also show that neither the elimination of Na/Ca exchange nor the sarcoplasmic reticulum function had significant effects on PAO-induced hypercontracture. The time to 50% hypercontracture for those groups is shown in Fig. 5B. Only preincubation with BAPTA-AM prevented hypercontracture (see Figs. 4 and 6), substantiating the requirement for intracellular Ca2+ in PAO-induced hypercontracture (also confirmed in permeabilized cells, as shown later). Inhibiting other Ca2+ transport mechanisms likewise did not affect the ability of DTT to reverse PAO-induced hypercontracture (Fig. 5A), suggesting that the relaxant effect of DTT was mediated by another mechanism besides reducing cytoplasmic [Ca2+] (see below).


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  		Fig. 3.   PAO effects on free cytoplasmic [Ca2+] and ATP depletion. A: sequential confocal images of rhodamine 2-AM-loaded myocyte during exposure to 20 µM PAO (with 30 mM BDM), demonstrating a progressive rise in free cytoplasmic [Ca2+]. Tyrode buffer contained 1.8 mM Ca2+. B: increase in cytoplasmic free [Ca2+] in a suspension of fluo 3-AM-loaded myocytes on addition of 20 µM PAO or 2 µM carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), shown by arrow, or after addition of 20 µM PAO plus either thapsigargin (TG; 0.5 µM), ruthenium red (RR; 2 µM) + TG, or RR + TG + cyclosporin A (CSA; 0.5 µM). TG and RR were applied for 30 min, and CSA was applied 1 min before PAO. Tyrode buffer was nominally Ca2+ free. A.U., arbitrary units. C: effect of addition of FCCP (5 µM) or PAO (20 µM) (see arrows) on cytoplasmic MgATP, estimated from changes in free [Mg2+] in Mg2+ green-loaded myocytes in suspension in nominally Ca2+ free media. Increased fluorescence reflects decreased [MgATP] (see Ref. 25). Compared with PAO, FCCP induced much more rapid ATP depletion, which was prevented by 10 µM oligomycin (Oligo).



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  		Fig. 4.   A: cell shortening (hypercontracture) induced by addition of 50 µM PAO (arrow) in isolated myocytes during the following: trace a, control; trace b, 0.5 mM EGTA replacing Ca; trace c; preincubation with 5 mM of 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-AM (BAPTA-AM). B: means ± SD to reach half-maximal shortening (T0.5) during PAO-induced hypercontracture as a function of extracellular [Ca2+].



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  		Fig. 5.   A: hypercontracture induced by addition of 50 µM PAO (left arrow) and its reversal by addition of 5 mM dithiothreitol (DTT; right arrow), in the presence of the following: 1) no added Ca; 2) no added Ca2+ + replacement of Na+ with choline (0 Na+) to inhibit Na+-Ca2+ exchange; 3) pretreatment with 500 nM TG to deplete sarcoplasmic reticulum Ca2+ stores; 4) pretreatment with 2 µM of RR to inhibit mitochondrial Ca2+ uptake; and 5) combined interventions in 1-4 (0 Na+ + TG + RR). B: means ± SD to reach T0.5 during PAO-induced hypercontracture for conditions shown in Fig. 5A.



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  		Fig. 6.   Inhibition of PAO-induced hypercontracture by various interventions. Percentage of healthy, rod-shaped myocytes hypercontracted into rounded cells 5 min after exposure to the following: 20 µM PAO (CON) or 20 µM PAO with cyclosporin A (CSA; 0.4 µM), N-ethylmaleimide (NEM; 50 µM), BAPTA-AM (5 mM), DTT (2 mM), FCCP (4 µM), Oligo (10 µM), or Oligo + FCCP. All of the reagents were added 1-2 min before PAO except BAPTA, which was preincubated 15 min. Results are means ± SD for number of low-power fields examined (shown in parentheses).

These findings eliminate extracellular Ca2+ influx and sarcoplasmic reticulum Ca2+ release as primary causes of the PAO-induced rise in cytoplasmic Ca2+. Because PAO has been shown to increase intracellar [Ca2+] in endothelial cells in association with tyrosine phosphorylation of cytoskeletal proteins (11), we also examined the effect of peroxyvanadate (0.1 mM), a potent tyrosine phosphatase inhibitor, but it failed to induce hypercontracture over 30 min (data not shown). By exclusion, these results suggest that mitochondrial Ca2+ efflux by PAO is sufficient to raise cytoplasmic free [Ca2+] to a level causing hypercontracture. PAO-induced hypercontracture did not depend on mitochondria being highly Ca2+ loaded, because pretreatment with the mitochondrial Ca2+ uniporter inhibitor ruthenium red (2 µM) did not change the time to 50% hypercontracture in response to PAO (Fig. 5B). Even with all of these interventions (except BAPTA loading), PAO still led to hypercontracture within the same time frame (Fig. 5B). The reason why such a modest increase in cytoplasmic free Ca2+ caused hypercontracture in the presence of PAO is addressed later.

Hypercontracture was not equivalent to rigor, because severe ATP depletion with carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) caused cell shortening only to a brick, rigor form (to ~50-70% initial cell length), without progressing further to rounded shapes (to <30% initial cell length) as in Fig. 2. FCCP, which causes fast Ca2+ efflux from mitochondria via the uniporter (28), also caused cytoplasmic free [Ca2+] to increase, albeit to lesser extent than PAO (Fig. 3B). In contrast, ATP depletion was much more rapid with FCCP than PAO (Fig. 3C). In fact, if cellular ATP levels were depressed drastically by FCCP before exposure to PAO, PAO no longer induced hypercontracture (n = 10; Fig. 6). If ATP depletion by FCCP was slowed with oligomycin, which prevents F0F1-ATPase from consuming ATP when the mitochondrial proton gradient is dissipated (25), the ability of PAO to induce hypercontracture was restored (n = 6; Fig. 6).

The MPT inhbitor CSA (0.4 µM) was only partially effective at preventing PAO-induced hypercontracture (n = 8; Fig. 6), but DTT and N-ethylmaleimide (NEM) were completely effective (n = 12 and 5, respectively; Fig. 6). These compounds prevent cross-linking by reducing or modifying sulfhydryl groups, confirming that the effects of PAO were selectively mediated by cross-linking of vicinal sulfhydryl groups. In isolated cardiac mitochondria, PAO-induced MPT was also reversed by DTT, provided substrate was present to allow Delta Psi recovery (n = 4 preparations; Fig. 7A). The rapid time course of DTT-induced Delta Psi recovery in isolated mitochondria was comparable to the rapid reversal of PAO-induced hypercontracture by DTT in intact myocytes (Fig. 5A). In contrast, renormalization of cytoplasmic ATP levels after Delta Psi recovery occurs over a time course of many minutes (8), making it unlikely that an increased ATP-to-ADP ratio (17) was responsible for the rapid reversal of hypercontracture by DTT. This suggests that PAO may have a direct effect on myofilament Ca2+ sensitivity, which can be rapidly reversed by DTT. This possibility was further tested in permeabilized cardiac myocytes, as described below.


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  		Fig. 7.   Effects of PAO in isolated mitochondria. Shown are the reversibility of 25 µM of PAO-induced mitochondrial depolarization (top trace, using TMRM) and swelling (bottom trace) by 2 mM DTT + 5 mM succinate (Suc). Mitochondrial recovery depended on both DTT and Suc to reenergize mitochondria. Neither DTT nor Suc added alone was efficient. FCCP (0.5 µM) and alamethicin (Alam, 2.5 µg/ml) were added for calibration. This experiment was repeated and gave similar results in 4 preparations. Delta Psi , Membrane potential.

PAO Increases Myocyte Susceptibility to Hypercontracture by Increasing Myofibrilar Ca2+ Sensitivity

To examine the effects of PAO on susceptibility to Ca2+-induced hypercontracture, we permeabilized the sarcolemma of isolated myocytes with digitonin (20 µM). In the presence of 3 mM ATP and 0.5 mM EGTA, permeabilized myocytes retained a rod-shape morphology and typically tolerated additions of Ca2+ until the buffered free [Ca2+] in the bath reached >10 µM, at which point they rapidly underwent hypercontracture (Fig. 8A). PAO, at 50 µM, had no effect on cell length in the absence of Ca2+, confirming a Ca2+ requirement for PAO-induced hypercontracture. After PAO, however, hypercontracture was induced at much lower free [Ca2+] (Fig. 8A, trace b) than in its absence (0.7 ± 0.8 µM, n = 29, vs. 19 ± 10 µM, n = 15, respectively; Fig. 8B). Thus PAO increased myofilament sensitivity to Ca2+. Although we cannot exclude the possibility that this effect was mediated by MPT-related alterations in local myofibrillar ATP-to-ADP ratios, a well-recognized determinant of the myofibrillar Ca2+ sensitivity (17), the rapid reversal by DTT, suggests that cross-linking of vicinal sulfhydryl groups of the contractile elements may be involved. We examined whether PAO affected myofibrillar ATP hydrolysis, but 50 µM PAO had no effect on actin-activated HMM activity (Fig. 8C; n = 3).


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  		Fig. 8.   Effect of PAO on susceptibility to hypercontracture in digitonin-permeabilized cardiac myocytes. Bath solution contained (in mM) 130 KCl, 1 MgCl2, 0.5 EGTA, 10 HEPES (pH 7.2), and 3 ATP. A: trace a, in the absence of PAO; multiple Ca2+ additions (arrows, raising free [Ca2+] to 10-20 µM) were required to induce hypercontracture. Trace b, PAO (50 µM) induced hypercontracture after fewer Ca2+ additions (arrows, raising free [Ca2+] to ~0.7 µM). Preincubation with 500 nM TG and 2 µM RR in traces a and b had no significant effect on hypercontracture (not shown). B: means ± SD of added [Ca2+] required to induce hypercontracture in permeabilized cells with or without PAO (50 µM). C: PAO had no effect on actin-activated heavy meromyosin ATPase activity. At the top arrow 1 or 5 µM ATP was added, followed by 50 µM PAO at the bottom arrow. This experiment was repeated three times with equivalent results. Contr, control.

PAO-Induced Mitochondrial Ca2+ Release

The above results indicate that both increased myofilament Ca2+ sensitivity and a modest increase in cytoplasmic free [Ca2+] are required for PAO to induce hypercontracture. Furthermore, Fig. 5 demonstrates that even when extracellular Ca2+ was removed and sarcoplasmic reticulum Ca2+ was depleted from resting aerobically superfused myocytes, their mitochondria retained sufficient releasable Ca2+ to induce hypercontracture in response to PAO. To examine whether PAO could elevate extramitochondrial Ca2+ to a significant extent in non-Ca2+-loaded mitochondria, heart mitochondria were isolated and incubated in KCl buffer, which contains several micromolars of contaminant Ca2+. In this setting, nonenergized mitochondria were not expected to accumulate a significant amount of contaminant Ca2+, but some accumulation cannot be excluded. The addition of PAO-induced Delta Psi depolarization and rapid matrix swelling indicates MPT and concomitantly produced a significant rise in extramitochondrial [Ca2+] due to Ca2+ efflux (Fig. 9A). Although we did not measure matrix free [Ca2+], experiments under comparable conditions in beef heart mitochondria in KCl-HEPES-low-Ca2+ buffer determined this value to be 0.25 µM, similar to the values (0.1-0.2 µM) reported for mitochondrial matrix free [Ca2+] in resting isolated myocytes (6, 26). Paradoxically, this is equivalent to the resting cytoplasmic free [Ca2+] in isolated cardiac myocytes. Therefore, to generate a driving force for Ca2+ efflux, PAO would have to release Ca2+ from matrix buffering sites [the estimated matrix total-to-free Ca2+ ratio is ~6,000 under physiological conditions (20), similar to the effects of intracellular acidosis (12)].


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  		Fig. 9.   A: PAO-induced mitochondrial Ca2+ efflux. Mitochondria (1 mg/ml) were incubated in KCl-based buffer (see MATERIALS AND METHODS) with 1 µM Ca2+ green 5N or 0.4 µM TMRM. Arrow shows when 20 µM PAO was added. Delta Psi depolarization was accompanied by slight transient matrix shrinkage (see also Fig. 7), followed by Ca2+ efflux and matrix swelling indicating MPT. This experiment was repeated in 4 preparations with similar results. Addition of FCCP (1 µM), shown by second arrow, produced no further Ca2+ efflux or Delta Psi depolarization. B: PAO causes Ca2+ release from mitochondrial matrix Ca2+ buffers. Mitochondria were incubated in KCl-based buffer containing 2 mM Pi and 0.5 µM RR, permeabilized with 250 nM Alam (first arrow), and then exposed to 40 µM PAO (second arrow). Extramitochondrial free [Ca2+] (using Ca2+ green 5N) increased, indicating release of Ca2+ from mitochondrial binding sites. Experiment was repeated in 2 other preparations with similar results.

To test this possibility, we permeabilized isolated mitochondria with alamethicin (Fig. 9B). Control experiments using TMRM showed that alamethicin completely dissipated Delta Psi (n = 6 preparations), similar to PAO and FCCP. In the presence of a Ca2+ indicator, alamethicin caused a small increase in extramitochondrial free [Ca2+], reflecting equilibration with matrix [Ca2+]. On addition of PAO, a further large increase in extramitochondrial free [Ca2+] occurred (average increase in extramitochondrial Ca2+ 1.06 ± 0.2 µM, n = 3 preparations). These findings suggest that PAO directly released Ca2+ from mitochondrial matrix Ca2+ buffers, providing the source of Ca2+ causing the rise in cytoplasmic free [Ca2+]. Most importantly, the PAO-induced myoplasmic [Ca2+] increase, even though relatively small, promoted hypercontracture because of increased myofibrilar Ca2+ sensitivity. Under those conditions, the rate of hypercontracture was most likely regulated by the extent of mitochondrial Ca2+ efflux and subsequent myoplasmic [Ca2+] increase.

PAO Causes Irreversible Sarcolemmal Damage and Cell Death

Figure 10A shows that PAO led to irreversible membrane injury, as judged by cellular LDH release. Similar to hypercontracture, PAO-induced LDH release did not require extracellular [Ca2+] (Fig. 10A). However, the time course of LDH release was much slower than hypercontracture, suggesting that irreversible membrane injury was not mediated mechanically. In addition, BDM totally prevented hypercontracture yet did not prevent PAO-induced LDH release (Fig. 10B). Adding CSA with BDM to the incubation medium before PAO exposure markedly inhibited LDH release (Fig. 10B), suggesting that sarcolemmal membrane damage was directly related to MPT. CSA also decreased LDH release without BDM but was less effective. Coupled with the observation that BATPA prevented hypercontracture (Fig. 6) and LDH release (Fig. 10B), these findings implicate the rise in intracellular Ca2+ during MPT as a primary cause of sarcolemmal membrane injury in isolated myocytes, with hypercontracture playing at most a secondary role.


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  		Fig. 10.   PAO induces lactic dehydrogenase (LDH) release. A: with 20 µM PAO (at time 0), LDH release was similar with (n = 7) or without (n = 5) 1.8 mM of extracellular Ca2+ present and accelerated by 40 µM PAO + Ca2+ (n = 5). B: inhibition of PAO-induced LDH release by CSA or BAPTA-AM. Cells were incubated in nominally Ca2+-free Tyrode buffer containing 30 mM BDM (n = 6) or BDM + 400 nM CSA (n = 7), or pretreated with BAPTA-AM for 15 min, as shown in Fig. 6.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

This study shows that, in isolated cardiac myocytes, the chemical MPT inducer PAO caused severe hypercontracture and irreversible sarcolemmal injury. PAO increased myofibrillar Ca2+ sensitivity, explaining why hypercontracture occurred with only a modest increase in cytoplasmic free [Ca2+] associated with MPT-induced mitochondrial Ca2+ efflux. Irreversible membrane injury, attributed to MPT-induced changes in cytoplasmic [Ca2+] and [ATP], developed independently of hypercontracture.

PAO-Induced Hypercontracture

In isolated cardiac myocytes, PAO was very effective at inducing MPT (see Fig. 2) but also markedly increased susceptibility to hypercontracture at cytoplasmic free [Ca2+] normally well tolerated by permeabilized myocytes (see Fig. 8). In this setting, mitochondrial Ca2+ efflux due to PAO-induced MPT provided sufficient [Ca2+] to induce hypercontracture, because removal of extracellular Ca2+ efflux and depletion of sarcoplasmic reticulum Ca2+ stores before PAO treatment only modestly slowed the rate at which hypercontracture developed (see Fig. 5). On the other hand, CSA, which inhibits PAO-induced Ca2+ efflux via PTP, slowed the rate of hypercontracture development more effectively (see Fig. 6). The delay rather than prevention of hypercontracture by CSA is probably related to the PAO-induced increased myofibrillar Ca2+ sensitivity. Thus even if CSA was able to inhibit major part of PAO-induced mitochondrial Ca2+ efflux, hypercontracture still developed slowly due to increased myofibrillar Ca2+ sensitivity.

The ability of DTT and NEM to inhibit both increased myofilament Ca2+ sensitivity and PAO-induced MPT accounts for their complete effectiveness at preventing PAO-induced hypercontracture. An important finding is the rapidity (within seconds) with which DTT application reversed PAO-induced hypercontracture (Fig. 5A). The most likely explanation for this rapid effect is that DTT reversed PAO-induced sulfhydryl group cross-linking, which we postulate was the cause of the increased myofibrillar Ca2+ sensitivity (by an unknown mechanism). An alternative possibility is that increased myofibrillar Ca2+ sensitivity was due to reduction in the local ATP-to-ADP ratio as a result of PAO-induced MPT, because a low ATP-to-ADP ratio favors strongly bound cross-bridge states, thereby sensitizing the myofilaments to Ca2+ (19). The rapid restoration of Delta Psi by DTT in isolated mitochondria after exposure to PAO (Fig. 7) could potentially restore a normal ATP-to-ADP ratio and reverse the increased myofibrillar Ca2+ sensitivity. However, the nearly instantaneous relaxing effect of DTT on PAO-induced hypercontracture seems inconsistent with this mechanism, because in rat cardiomyocytes exposed to anoxia/reoxygenation, mitochondrial Delta Psi recovery was fast, but ATP regeneration occurred slowly, taking >10 min to restore prehypoxic levels (8). PAO also had no effect on actomyosin ATPase activity in vitro (Fig. 8C), suggesting that increased Ca2+ sensitivity was not related to enhanced ATP hydrolysis by this enzyme, generating reduced local ATP-to-ADP ratios in the vicinity of myofilaments. Thus, although the mechanism of the effect of PAO on myofibrillar Ca2+ sensitivity remains unclear, it is most consistent with a direct effect on contractile elements of sulfhydryl group cross-linking. The specific protein targets of this agent, however, remain unknown, with respect to the subunits of the multiprotein PTP complex that must be cross-linked via vicinal sulfhydryl groups to induce MPT.

PAO, as a general sulfhydryl group cross-linking agent, is likely to affect a variety of cellular proteins. Although we cannot exclude the possibility that PAO affected other cellular processes that regulate cytoplasmic free Ca2+, mitochondrial Ca2+ efflux coupled with increased myofilament Ca2+ sensitivity provides a sufficient explanation for hypercontracture. In addition to inducing MPT, PAO has been shown to activate neutrophil NADPH oxidase (9) receptor-mediated endocytosis (32), cause tumor necrosis factor-dependent activation of nuclear factor-kappa B (34), and stimulate 2-deoxyglucose transport (4). None of these effects can explain the rapid onset of hypercontracture and irreversible cell injury. Furthermore, PAO has been shown to increase intracellular [Ca2+] in endothelial cells in association with tyrosine phosphorylation of cytoskeletal proteins (11). However, the potent tyrosine phosphatase inhibitor peroxyvanadate did not induce hypercontracture in cardiac myocytes.

Like PAO, FCCP induces mitochondrial Ca2+ efflux and MPT in Ca2+-loaded mitochondria (3) but did not cause hypercontracture in isolated myocytes. FCCP by itself induced rigor (shortened square cells) rather than hypercontracture (rounded cells with extensive blebs), as has been reported previously (7). In addition, pretreatment with FCCP prevented hypercontracture (but not MPT) by PAO. From Fig. 3, B and C, we suggest that FCCP decreased [ATP] so extensively that myofilaments went into a rigor state in which further cross-bridge cycling was prevented (27) despite the rise in free cytoplasmic [Ca2+]. In contrast, PAO decreased the [ATP] more slowly while increasing mitochondrial Ca2+ efflux rate more rapidly and concomitantly increasing myofibrillar Ca2+ sensitivity. If ATP was severely depleted by FCCP pretreatment before PAO exposure, any further cross-bridge cycling was prevented, so that the cell remained in rigor and did not hypercontract after PAO exposure, despite increases in cytoplasmic free [Ca2+] and myofibrillar Ca2+ sensitivity.

PAO-Induced Membrane Injury and Cell Death

Hypercontracture did not directly cause irreversible membrane injury and cell death in this model, because BDM prevented hypercontracture yet did not prevent the latter (Fig. 10). In contrast, preincubation with BAPTA-AM prevented both hypercontracture and membrane injury (Fig. 10), implicating the MPT-induced increase in cytoplasmic Ca2+, rather than hypercontracture per se, as the primary cause of irreversible injury. It is not surprising that a nonmechanically mediated mode of necrotic cell death exists in cardiac tissue, because widespread MPT is known to promote necrotic cell death in noncontractile cells that are incapable of hypercontracture (24). This nonmechanical mode may therefore represent a general necrotic cell death mechanism triggered by high cytoplasmic free Ca2+ and low [ATP], both consequences of MPT (7).

Implications

Although no direct link can be established on the basis of our findings, the effects of PAO bear some striking similarities to reperfusion/reoxygenation injury, including MPT (5), hypercontracture, and enhanced susceptibility to hypercontracture (23), irreversible membrane injury, prevention of hypercontracture by pretreatment with metabolic inhibitors (13), and partial protection by CSA (17). However, there are also significant differences. In contrast to the normal state of mitochondrial Ca2+ loading before PAO exposure, mitochondria become heavily Ca2+ loaded during ischemia/anoxia (10), a key factor directly promoting MPT (5, 17), hypercontracture, and irreversible injury on reoxygenation (16, 26). Also, as discussed above, PAO is a potent sulfhydryl group cross-linker, which is likely to affect other proteins besides those affected by reperfusion/reoxygenation, and vice versa. Despite these differences, however, the similar effects on cardiac function are intriguing and potentially illuminating. Both hypercontracture (30) and MPT (5, 17) have been proposed to play central roles in reperfusion/reoxygenation injury, but the relationship between the two, if any, has not been well documented. Our findings suggest that Ca2+ release from mitochondria during MPT can markedly enhance hypercontracture, provided that myofibrillar Ca2+ sensitivity is increased. In the setting of reperfusion/reoxygenation, in which mitochondria are highly Ca2+ loaded (to a much greater extent than mitochondria in this study), mitochondrial Ca2+ release during MPT is likely to make an even greater contribution to hypercontracture, with rapid generation of uncontrolled excessive force directly disrupting sarcolemmal membrane integrity and causing immediate enzyme release (30). In addition, the rise in free cytoplasmic [Ca2+] may trigger MPT-associated necrotic cell death independent of hypercontracture and associated with delayed enzyme release.

In conclusion, both of these MPT-related necrotic cell death mechanisms, in addition to apoptosis (17), are likely to play important roles in reperfusion/reoxygenation injury in intact cardiac muscle. In view of its efficacy at rapidly inducing MPT and increasing myofibrilar Ca2+ sensitivity, PAO may be a useful tool for investigating the underlying mechanisms. Our findings suggest that prevention of MPT during reperfusion/reoxygenation has promise as a therapeutic target.


    ACKNOWLEDGEMENTS

This study was supported by National Heart, Lung, and Blood Institute Specialized Center of Research in Sudden Cardiac Death Grant P50-HL-52319 and R29-HL-51129, the Laubisch Cardiovascular Research Fund, and the Kawata Endowment.


    FOOTNOTES

Address for reprint requests and other correspondence: J. N. Weiss, Div. of Cardiology, 3641 MRL Bldg., University of California at Los Angeles School of Medicine, Los Angeles, CA 90095-1760 (E-mail: jweiss{at}mednet.ucla.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 27 September 2000; accepted in final form 9 January 2001.


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