AJP - Heart Fuel your research with LabChart
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 290: H1549-H1557, 2006. First published November 11, 2005; doi:10.1152/ajpheart.00913.2005
0363-6135/06 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
290/4/H1549    most recent
00913.2005v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (5)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Marcil, M.
Right arrow Articles by Burelle, Y.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Marcil, M.
Right arrow Articles by Burelle, Y.

Exercise training induces respiratory substrate-specific decrease in Ca2+-induced permeability transition pore opening in heart mitochondria

Mariannick Marcil, Karine Bourduas, Alexis Ascah, and Yan Burelle

Département de Kinésiologie, Université de Montréal, Montreal, Quebec, Canada

Submitted 24 August 2005 ; accepted in final form 9 November 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The purpose of this study was to determine whether regular exercise (treadmill running, 10 wk) alters the susceptibility of rat isolated heart mitochondria to Ca2+-induced permeability transition pore (PTP) opening and whether this could be associated with changes in the modulation of PTP opening by selected physiological effectors. Basal leak-driven and ADP-stimulated respiration in the presence of substrates for complex I, II, and IV were not affected by training. Fluorimetric studies revealed that in the control and exercise-trained groups, the amount of Ca2+ required to trigger PTP opening was greater in the presence of complex II vs. I substrates (230 ± 12 vs. 134 ± 7 nmol Ca2+/mg protein, P < 0.01; pooled average of control and trained groups). In addition, with a substrate feeding the complex II, training increased by 45% (P < 0.01) the amount of Ca2+ required to trigger PTP opening both in the presence and absence of the PTP inhibitor cyclosporin A. However, membrane potential, reactive oxygen species production, NAD(P)H ratio, and Ca2+ uptake kinetics were not different in mitochondria from both groups. Together, these results suggest the existence of a substrate-specific regulation of the PTP in heart mitochondria and suggest that regular exercise results in a reduced sensitivity to Ca2+-induced PTP opening in presence of complex II substrates.

mitochondrial function; calcium stress

MITOCHONDRIA PLAY a pivotal role in controlling cell death through their capacity to trigger both necrosis and apoptosis (16, 18, 25, 39). A number of studies have shown that an increased permeability of the mitochondrial membranes is a key event in these processes (37, 41, 59). Although several mechanisms of membrane permeation have been suggested, one of the best documented involves the opening of the permeability transition pore (PTP).

The PTP is a high-conductance nonspecific pore presumably formed by a supramolecular complex spanning the double-membrane system of the mitochondria mainly at contact sites. Although it is increasingly recognized that the molecular composition of the PTP is probably variable (61), the prevailing hypothesis is that it includes the adenylate translocator (ANT), the porin pore (voltage-dependent anion channel, VDAC), and the matrix protein foldase cyclophilin D. Opening of the PTP induces the loss of mitochondrial membrane potential ({Delta}{Psi}), uncoupling of oxidative phosphorylation, high-amplitude swelling of the matrix, and the release of several proapoptotic factors that are normally sequestered in mitochondria such as cytochrome c, apoptosis-inducing factor, Smac/Diablo, endonuclease G, and Omi/HtrA2 (5, 25).

In the heart, PTP opening was shown to occur during reperfusion after ischemia and to be involved in contractile dysfunction and tissue injury (19, 23, 24, 31, 32, 35). This phenomenon can be explained by the fact that many of the conditions required to open the PTP in vitro prevail in cardiac cells early during reperfusion. On the other hand, ischemic preconditioning was shown to decrease the sensitivity to PTP opening in isolated mitochondria (1) and intact cardiomyocytes (28) and perfused hearts (27, 31). However, whether exercise training, another physiological stress capable of inducing a cardioprotective phenotype (9, 14, 42, 51), can beneficially alter the regulatory properties of the PTP remains largely unknown. Ca2+ concentration ([Ca2+]) in the matrix is the most important determinant of PTP gating, with high [Ca2+] favoring the open conformation (60). In addition, a variety of factors modulate the sensitivity of the PTP to Ca2+, including variations in the {Delta}{Psi}, redox state of pyridine nucleotides (PNs), reactive oxygen species (ROS) production, and matrix pH as well as Pi and adenylate content (see Ref. 60 for review). In skeletal muscle mitochondria, the Ca2+ sensitivity of the PTP was also shown to depend on the type of respiratory substrate oxidized, with complex I donors acting as sensitizers compared with complex II donors (22).

In the present study we therefore determined whether exercise training is associated with changes in the sensitivity to Ca2+-induced PTP opening in isolated heart mitochondria. We also determined whether the type of substrate used for energization influences Ca2+-induced PTP opening in this organ and whether exercise training elicits changes in selected physiological modulators of PTP gating.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal care. All experiments were conducted according to the directives of the Canadian Council on Animal Care and were approved by the Université de Montréal Animal Care Committee. Female Sprague-Dawley Rats (Charles River, St-Constant, PQ, Canada) weighing ~250 g were housed in pairs and kept in a temperature-, humidity-, and light-controlled (12:12-h light-dark cycle) environment. The animals had free access to standard rat chow and water.

Exercise protocol. After a week of habituation, animals were divided into control (C) and exercise-trained (T) groups. T rats were run on a motor-driven rodent treadmill (Quinton Instruments, Seattle, WA) at 25 m/min and 16% slope on 4 days/wk for 10 wk. Running time was set at 30 min during the first week, 60 min during the second week, and 90 min during the third week. Running time was then maintained at 90 min for the remaining 7 wk.

Materials. All chemicals were purchased from Sigma (St. Louis, MO), with the exception of cyclosporin A (CsA; Tocris, Ellisville, MO), and Calcium Green-5N (Molecular Probes, Eugene, OR).

Mitochondrial isolation. Heart mitochondria were prepared as described by Fontaine et al. (22) with slight modifications. Animals were anesthetized (pentobarbital sodium 50 mg/kg ip) 48 h after the last training session. Hearts were rapidly excised and immersed into ice-cold isolation medium (buffer A; in mM: 300 sucrose, 10 Tris·HCl, 1 EGTA, pH 7.3) and weighed. Ventricular tissue was minced with scissors in 5 ml of buffer A supplemented with 0.2% fatty acid free BSA and homogenized with a Polytron tissue tearer (~3 s at a setting of 3). The homogenate was then incubated with the protease nagarse (1.5 mg/g) for 5 min and further homogenized at the same settings. The homogenate volume was completed to 30 ml with buffer A + 0.2% BSA and centrifuged at 800 g for 10 min. The pellet was discarded, and the supernatant was decanted and centrifuged at 10 000 g for 10 min. The pellet obtained was resuspended in buffer B (in mM: 300 sucrose, 0.5 EGTA, 10 Tris·HCl, pH 7.3) and centrifuged at 10 000 g for 10 min. After this washing step was repeated twice, the final mitochondrial pellet was resuspended in 0.3 ml of buffer B to a protein concentration of ~20 mg/ml. All procedures were carried out at 4°C. Protein determinations were performed by the bicinchoninic acid method (Pierce, Rockford, IL), with BSA as a standard.

Mitochondrial respiration. Mitochondrial oxygen consumption was measured polarographically at 22°C, using Clark-type electrodes (Oxygraph, Hansatech Instruments, Kings Lynn, UK). Experiments were started with the addition of 0.30 mg of mitochondria in 1 ml of buffer C (in mM: 125 KCl, 10 KH2PO4, 0.05 EGTA, 10 Tris-MOPS, 2.5 MgCl2). Respiratory substrates feeding complex I (5 mM glutamate-2.5 mM malate), complex II (5 mM succinate) or complex IV [0.1 mM N,N,N',N'-tetramethyl-p-phenylenediamine (TMPD)-1 mM ascorbate] were added in the incubation medium. All substrates were free acids buffered to pH 7.3 with Tris. Experiments for the complex II were made in presence or absence of the complex I inhibitor rotenone (1 µM) (Fig. 1). The medium was then supplemented with 0.25 mM ADP to measure maximal rate of oxidative phosphorylation (VADP). When respiration reached state 4 after complete phosphorylation of ADP, 0.5 µM oligomycin was added to measure oligomycin-insensitive respiration (Voligo), which eliminates the contribution of slow turnover of adenylates to basal respiration due to the presence of residual ATPase activity in the mitochondrial preparation. Respiratory control ratio (RCR) was calculated as the ratio VADP/Voligo, and the amount of ATP synthesized per molecule of oxygen consumed (P/O) was calculated before state 4 was reached.


Figure 1
View larger version (20K):
[in this window]
[in a new window]
  		Fig. 1. Overview of respiratory substrates and inhibitors used: the respiratory substrates providing electrons to complex I II and IV are underlined. The various inhibitors (rotenone, antimycin A, and oligomycin) used are shown over their respective sites of action. Glut/mal, glutamate-malate; TMPD/asc, N,N,N',N'-tetramethyl-p-phenylenediamine (TMPD)-1 mM ascorbate; Q, ubiquinone; C, cytochrome c.

 
Calcium challenge. Mitochondria (0.3 mg/ml) were incubated at 22°C in 2 ml of buffer D (in mM: 250 sucrose, 10 MOPS, 0.05 EGTA, 10 KH2PO4, pH 7.2) containing glutamate-malate (5 mM-2.5 mM) or succinate (5 mM) in the presence or absence of 1 µM rotenone. Changes in extramitochondrial [Ca2+] were monitored fluorimetrically (Hitachi, F4500 spectrofluorometer), using Calcium-Green 5N (1 µM; excitation 505 nm, emission 535 nm) as described by Ichas et al. (30). Residual [Ca2+] was adjusted to the same level at the beginning of every experiment by adding a small amount of EGTA. Unless stated otherwise, Ca2+ pulses (42 nmol/mg protein) were then added at 2-min intervals until a Ca2+-induced mitochondrial Ca2+ release was observed. Calcium retention capacity (CRC) was taken as the total amount of Ca2+ accumulated by mitochondria before the Ca2+ pulse triggering Ca2+ release.

In some experiments, mitochondrial {Delta}{Psi} was measured under the same experimental conditions. For this purpose, Calcium-Green 5N was replaced with rhodamine 123 (0.2 µM; excitation 503 nm, emission 525 nm), and measurements were performed as described by Emaus et al. (21). Mitochondrial release of rhodamine 123 after uncoupling with 100 nM CCCP was taken as an index of {Delta}{Psi}. Mitochondrial swelling in response to Ca2+ pulses was measured under the same experimental conditions, using light diffraction at a setting of 545 and 545 nm (29). Each experiment was performed either in the presence or the absence of 1 µM CsA. This concentration is commonly used (4, 22, 45) and is severalfold higher than that required for full inhibition of PTP in liver (~0.15 µM; Ref. 47) and heart (~0.3 µM; Refs. 15, 17) mitochondria under standardized conditions of Ca2+ loading.

ROS production. Mitochondrial H2O2 production was measured fluorimetrically, as described by Servais et al. (53) (excitation 319 nm, emission 420 nm). Mitochondria (0.1 mg/ml) were incubated in 2 ml of buffer D containing 5 U/ml of horseradish peroxidase (HRP) and 0.1 mM homovanillic acid (HVA). At the end of each test 2 µmol of H2O2 was added as an internal standard to allow calculation of endogenous ROS production by the respiratory chain.

PN oxidation-reduction status. The oxidation-reduction status of the mitochondrial PN pool was evaluated based on endogenous NAD(P)H fluorescence (excitation 340 nm, emission 460 nm). Mitochondria (0.3 mg/ml) were incubated in 2 ml of buffer D, and redox state of PNs was measured in the presence of glutamate-malate (5 mM-2.5 mM) or succinate (5 mM) ± rotenone (1 µM). Redox state of PN was calculated as described by Arieli et al. (2): NAD(P)H ratio = (Fs – Fmin)/(Fmax – Fmin), where Fs is the fluorescence measured after addition of respiratory substrates, Fmin is the baseline fluorescence of mitochondria measured in the absence of respiratory substrates, which represents PNs in their fully oxidized form, and Fmax is the fluorescence recorded after addition of 2 µM antimycin A, which represents PNs in their fully reduced form. Preliminary experiments indicated that Fmin values measured with this method were similar to those obtained when mitochondria were uncoupled by the addition of 100 nM CCCP.

Mitochondrial adenylate content. Endogenous ATP and ADP contents were measured in neutralized perchloric acid extracts with a luciferin/luciferase assay described by Drew and Leeuwenburgh (20), with modifications. Briefly, ATP content was evaluated based on the light production [Planck's constant x frequency (h{nu})] from the reactions luciferin + ATP -> luciferyl adenylate + PPi and luciferyl adenylate + O2 -> oxyluciferin + AMP + h{nu}.

In an aliquot of the same sample, ATP + ADP content was measured in a similar way after endogenous ADP was converted to ATP with excess amounts of pyruvate kinase (20 U/ml) and phosphoenolpyruvate (PEP; 5 mM) added directly in the cuvette. ADP content was calculated by subtracting the bioluminescence decay curves obtained in both samples. For every assay, baseline light emission (h{nu}) before the addition of the sample was subtracted from the luminescent decay. In addition, it was verified that the presence of pyruvate kinase or PEP did not affect the relationship between ATP concentration and h{nu}.

Statistical analyses. Results are expressed as means ± SE. Two-tailed Student's t-tests were performed to assess statistical significance. When multiple comparisons were made, differences were compared with ANOVA and Tukey post hoc tests were performed to identify the location of significant differences. The Bonferroni correction was applied to the P value obtained to correct for multiple comparisons. A corrected P value <0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Morphometric parameters. In line with our previous work (14), the exercise training regimen used in the present study resulted in myocardial hypertrophy, as indicated by a significant increase in heart weight, ventricular weight (+11%, P < 0.05) and heart weight-to-body weight ratio (+10%, P < 0.05). No significant changes in body weight and daily food intake were observed between the experimental groups. Mitochondrial yields were similar in hearts from C and T rats (~16 mg/g ventricular tissue) (Table 1).


View this table:
[in this window]
[in a new window]
  		Table 1. Morphometric data and mitochondrial isolation yield in sedentary control and exercise-trained rats

 
Mitochondrial respiration. Table 2 shows the results of experiments aimed at characterizing the effect of exercise training on basic respiratory parameters. In mitochondria respiring with substrates for complex I, II, and IV, training was not associated with any changes in maximal ADP-stimulated and oligomycin-insensitive leak-driven respiration, RCR, and P/O. These data are consistent with previous reports showing that exercise training does not significantly affect oxidative capacity (43).


View this table:
[in this window]
[in a new window]
  		Table 2. Effect of exercise on mitochondrial respiration

 
Ca2+ challenge. Figure 2 shows the typical response of mitochondria energized with the complex I substrates glutamate and malate to series of Ca2+ pulses. In these conditions, mitochondria accumulated 134 ± 7 nmol Ca2+/mg protein before abrupt release of accumulated Ca2+ occurred. The release of Ca2+ was invariably accompanied by high-amplitude swelling. When the experiments were performed in the presence of the PTP inhibitor CsA (1 µM), the amount of Ca2+ required to trigger these effects significantly increased 3.5-fold (483 ± 50 nmol/mg protein, P < 0.01), indicating that these phenomena were caused by the opening of the PTP. However, as shown in Fig. 3, in the presence of glutamate-malate, exercise training resulted only in a small increase in CRC that was not statistically significant (P = 0.15) and CsA had the same effect as in mitochondria from control hearts.


Figure 2
View larger version (22K):
[in this window]
[in a new window]
  		Fig. 2. Mitochondrial response to a train of Ca2+ pulses. Typical Calcium-Green 5N (A and B) and light scattering at 545 nm (C and D) of mitochondria energized with glutamate-malate (5 mM-2.5 mM). Tracings show progressive Ca2+ accumulation followed by release of accumulated Ca2+ and high-amplitude swelling. The presence of 1 µM cyclosporin A (CsA; B and D) doubles the amount of Ca2+ required to trigger these effects, indicating that these phenomena are related to permeability transition pore (PTP) opening. Each arrow indicates the addition of a Ca2+ pulse of 42 and 210 nmol/mg for the Calcium-Green 5N and swelling experiments, respectively. [Ca2+], Ca2+ concentration.

 

Figure 3
View larger version (13K):
[in this window]
[in a new window]
  		Fig. 3. Effect of exercise on PTP sensitivity to Ca2+ with complex I and II substrates. Ca2+ retention capacity (in nmol/mg protein) of control (C) and exercise-trained (T) mitochondria energized with glutamate-malate (GM) and succinate in absence (S) and presence (SR) of 1 µM rotenone is shown. Experiments were performed in the absence (GM, S, and SR) or presence (GM+CsA, S+CsA, and SR+CsA) of 1 µM CsA. Values are means ± SE for at least 7 separate experiments in the C and T groups, respectively. aSignificantly different from control (P < 0.05); bsignificantly different compared with –CsA (P < 0.01).

 
Figure 3 also shows the results of similar experiments performed in mitochondria respiring with the complex II substrate succinate. In both experimental groups, CRC in the presence of succinate was ~60–100% higher compared with that observed in the presence of glutamate-malate whether or not rotenone was present.

In contrast to what was observed in mitochondria energized with glutamate-malate, training significantly increased CRC by 45% in mitochondria energized with succinate substrates both in the presence (245 ± 14 vs. 327 ± 25 nmol/mg protein, P < 0.05) and the absence (215 ± 25 vs. 292 ± 26 nmol/mg protein, P < 0.05) of the complex I inhibitor rotenone. This difference remained unaltered in the presence of CsA, which increased CRC to a similar extent in both experimental groups. Importantly, training was not associated with changes in Ca2+ uptake kinetics in any of the conditions tested, as indicated by the calculated time to 50% uptake of the first two or three Ca2+ pulses (26.5 ± 1.7 s and 29.6 ± 2.0 s in C and T mitochondria, respectively; n = 6 in each group).

PTP modulators. To gain insights on the mechanism(s) underlying this increase in CRC with complex II substrates, several physiological modulators of PTP gating were investigated. It is well established that the PTP behaves as a voltage-gated channel and that mild depolarization favors the open conformation in the presence of Ca2+ loading (7, 22, 60). For this reason, the effect of training on {Delta}{Psi} before PTP opening was determined. At baseline, {Delta}{Psi} was similar in mitochondria from C and T animals, as indicated by a comparable release of rhodamine 123 after addition of 100 nM CCCP (553 ± 42 vs. 580 ± 64 arbitrary fluorescence units in C and T groups, respectively). In addition, the transient depolarizations associated with the uptake of single Ca2+ pulses during the Ca2+ loading phase were of similar amplitude (Fig. 4). As expected, once the pore opened, an abrupt and complete loss of {Delta}{Psi} was observed in both experimental groups. However, consistent with the CRC data, significantly more Ca2+ was required to trigger this effect in mitochondria from T compared with C animals.


Figure 4
View larger version (13K):
[in this window]
[in a new window]
  		Fig. 4. Changes in membrane potential ({Delta}{Psi}) in response to Ca2+ pulses in mitochondria from control and trained animals. Representative traces of changes in {Delta}{Psi} measured with rhodamine 123 during Ca2+ challenge experiments in the presence of succinate are shown. Each arrow indicates the addition of a calcium pulse of 42 nmol Ca2+/mg protein. AU, arbitrary units.

 
Figure 5 shows the results of experiments in which ROS production was measured. In line with previous studies (57), mitochondria generated a significant amount of ROS in presence of succinate alone (Fig. 5, A and B). Addition of rotenone decreased ROS production by half, consistent with the idea that succinate leads to ROS production at the level of the FeS center of complex I through reverse electron flow (57). Subsequent addition of antimycin A led to a large increase in ROS production, which in the presence of rotenone occurs mainly at the level of complex III. However, training did not alter steady-state ROS production with succinate alone and in the presence of both inhibitors. Similarly, training did not significantly affect ROS production when mitochondria were energized with glutamate-malate whether or not electron flow trough complex I was inhibited by rotenone (Fig. 5, C and D).


Figure 5
View larger version (21K):
[in this window]
[in a new window]
  		Fig. 5. Effect of exercise on mitochondrial H2O2 production. Representative traces of H2O2 production in heart mitochondria of control (A and C) and trained (B and D) animals when energized with succinate (A and B) or GM (C and D) are shown. Numbers below traces represent the rate of H2O2 production in nanomoles per minute per milligram of protein. Data are means ± SE for at least 7 experiments in each group. M, mitochondria; S, succinate; Rot, rotenone (1 µM); Ant A, antimycin-A (2 µM). aSignificantly different from succinate alone (P < 0.01); bsignificantly different from succinate in presence of rotenone (P < 0.01); csignificantly different from succinate alone (P < 0.01); dsignificantly different from GM alone (P < 0.01).

 
Figure 6 shows the results from experiments in which the redox state of PNs was measured in mitochondria from C and T animals. After addition of mitochondria in the absence of respiratory substrate, endogenous autofluorescence of PNs stabilized at a low level in both experimental groups (Fig. 6A). Addition of succinate led to a rapid reduction of PNs, and subsequent inhibition of electron flow with antimycin A led to the complete reduction of the PN pool. In some experiments 100 nM CCCP was added instead of antimycin A. Under this condition, fluorescence returned to the level observed in mitochondria in the absence of respiratory substrate (data not shown). Figure 6B shows the steady-state NAD(P)H ratio computed from several experiments in mitochondria respiring with succinate in the presence and absence of rotenone. No significant differences were observed between the experimental groups in any of the conditions tested.


Figure 6
View larger version (13K):
[in this window]
[in a new window]
  		Fig. 6. Effect of exercise on redox state of pyridine nucleotides (PNs). A: representative traces of PN fluorescence at 460 nm in heart mitochondria of control and trained rats when energized with succinate. B: the redox state in presence of succinate in the absence and presence of rotenone. Redox state is expressed as the ratio (FS – Fmin)/(Fmax – Fmin). Fluorescence levels corresponding to these values are shown in A (see METHODS for further details). Values are means ± SE for 6 separate experiments in each group.

 
Table 3 shows the endogenous adenine nucleotide content measured in frozen mitochondrial extracts from C and T animals. Training did not significantly alter the residual ATP, ADP, and total adenylate contents or the ATP-to-ADP ratio.


View this table:
[in this window]
[in a new window]
  		Table 3. Adenylate content of heart mitochondria from control and trained rats

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Results from the present study provide evidence that in heart mitochondria sensitivity to Ca2+-induced PTP opening is influenced by the type of respiratory substrate oxidized, with complex I donors acting as sensitizers compared with complex II substrates. In addition, the present results indicate that regular exercise training can decrease the sensitivity to Ca2+-induced PTP opening when mitochondria are energized with complex II substrates. This effect of training was observed in the absence of changes in the physiological modulators of PTP opening investigated.

Substrate-specific regulation of PTP. Fontaine et al. (22) showed that in skeletal muscle mitochondria CRC is three- to fourfold lower in mitochondria energized with glutamate-malate compared with succinate. This marked difference in CRC was shown to be due to the fact that the factors contributing to PTP opening are different in the two experimental conditions. Indeed, in the presence of complex I substrates the electron flow through this complex, independent of other regulators (i.e., redox state of PNs, {Delta}{Psi}, pH, ROS production), appeared to be the main factor regulating PTP opening by acting as a potent sensitizer (6, 22, 40). In contrast, when complex I was bypassed with succinate the mechanism was largely inactive and the contribution of other regulators to PTP opening was unmasked. Results from the present study are in line with these data. Indeed, in both experimental groups a substantially lower CRC was observed in presence of glutamate-malate compared with succinate. Our data thus indicate that, similar to what is observed in skeletal muscle, the electron flow through complex I sensitizes heart mitochondria to PTP opening.

Training-induced alteration in Ca2+ handling. To our knowledge, the impact of exercise training on Ca2+ handling by heart mitochondria has only been investigated in two studies (50, 56), and the effect on the PTP was not directly assessed. Sordahl et al. (56) reported that the rate of Ca2+ uptake by isolated heart mitochondria energized with succinate was unchanged after training in dogs. Similar findings were reported by Penpargkul et al. (50) in rodents in response to swim training. Results from the present study are in line with these findings. Indeed, Ca2+ uptake kinetics measured during the first two or three Ca2+ pulses of the Ca2+ challenge did not differ between the two experimental groups whether mitochondria were energized with complex I or II substrates. These data thus suggest that training has little effect on the electrophoretic mechanism of Ca2+ uptake in heart mitochondria.

In the study by Sordahl et al. (56), training was found to decrease the capacity of mitochondria to accumulate Ca2+ when energized with succinate. At the time this study was published, the existence of the PTP and the inhibitory effect of cyclosporins on pore opening were unknown. However, given the experimental conditions used, this phenomenon was likely caused by a premature opening of the PTP in mitochondria from trained animals. As for the study by Penpargkul et al. (50), the experimental conditions used did not allow observation of a permeability transition.

In contrast to the results from Sordhal et al. (56), data from the present study indicate that in mitochondria from T rats the amount of Ca2+ required to trigger PTP opening was significantly 45% higher in the presence of succinate both in the presence and in the absence of rotenone. In addition, CsA increased CRC in both experimental groups but did not abolish the effect of training. These results thus indicate that, in our conditions, training was able to decrease the sensitivity to Ca2+-induced PTP opening in the presence of complex II substrates. In addition, the fact that the effect of training was not abolished by CsA used at a concentration threefold in excess of that required to fully inhibit PTP opening (15, 17, 47) suggests that the effect of training is not related to a reduction in the expression of cyclophilin D or its interaction with other PTP components (3, 4). Indeed, changes in the expression of cyclophilin D are known to translate into an altered potency of CsA at inhibiting Ca2+-induced PTP opening (3, 4, 45, 52).

Our results also indicate that, in contrast to what was observed with succinate, no significant difference between C and T groups was observed for CRC in the presence of complex I substrates. As mentioned above, PTP opening is strongly regulated by electron flow through complex I, which sensitizes mitochondria to permeability transition (6, 22, 40). Our data thus indicate that training did not protect against this sensitizing effect. In contrast, when complex I was bypassed a protective effect of training, probably through other regulators of PTP opening, was unmasked.

Effect of training on physiological modulators of PTP opening. To our knowledge, there are no data available in the literature concerning the effect of training on parameters involved in the regulation of the PTP in the heart. In the present study, we therefore determined whether the training-induced increase in the resistance to Ca2+-induced PTP opening observed with complex II substrates was accompanied by changes in selected physiological modulators of PTP gating and/or in mitochondrial respiratory activity.

Ca2+-induced permeability transition is modulated by a variety of physiological effectors. The occurrence of PTP opening at a given Ca2+ load is reduced when {Delta}{Psi} is increased because the PTP behaves as a voltage-gated channel sensitive to changes in {Delta}{Psi} over a range of 180–120 mV (7). Matrix adenine nucleotides are also potent inhibitors of PTP opening, with ADP exerting a stronger inhibitory effect than ATP (16, 60). Maintenance of the PN pool at a high reduction state is another factor known to decrease PTP opening induced by Ca2+ and Pi, probably through mechanisms involving direct interaction of PNs with the PTP and oxidation of critical SH residues of pore-forming proteins (6, 60). Low levels of ROS production (12, 26) as well as an acidic pH (8, 48) will also reduce the occurrence of PTP opening. Finally, the sensitivity of the PTP to Ca2+ was shown to be modulated by members of the Bcl-2 family of proteins, the proapoptotic Bid and Bax and the antiapoptotic Bcl-2 and Bcl-Xl acting as facilitators and repressors of pore opening, respectively (11, 38, 44, 46, 49).

However, in the present study we did not find evidence indicating that changes at the level of these PTP modulators could account for the increased resistance to Ca2+-induced PTP opening observed. Indeed, baseline {Delta}{Psi} at steady state as well as the amplitude of the transient depolarizations associated with the uptake of single Ca2+ pulses were similar in both groups, suggesting that {Delta}{Psi} before PTP opening was not affected by training. Endogenous ATP and ADP content as well as the ATP-to-ADP ratio were also similar in mitochondria from the C and T groups. In addition, the redox state of PNs measured was unchanged. As for matrix pH, it is unlikely to be a factor because the high Pi concentration used during the Ca2+ challenge experiments prevents any fluctuations in matrix pH secondary to changes in respiration (22).

In the heart, several studies have shown that training results in a significant increase in tissue content of enzymatic and nonenzymatic antioxidant systems, including some that are found in mitochondria (33, 36, 55). However, there are apparently no data available on the effect of training on the actual rate of ROS production by active mitochondria. Moreover, data available in skeletal muscle mitochondria on this question are conflicting, showing either a reduction (58) or no change (53) in ROS production after training.

In the present study, the rate of H2O2 production of mitochondria respiring with complex I or II substrates under state 4 conditions was not significantly affected by training. In addition, reduction of respiratory chain complexes with rotenone or antimycin A, which induce a large increase in ROS production, had similar effects in both experimental groups. These results would thus suggest that altered ROS production is not responsible for the increased CRC observed in mitochondria after training. However, ROS production could not be accurately measured during the Ca2+ challenge because of artifactual changes in the fluorescence of HVA in these conditions (5). Therefore, the present data cannot rule out the possibility that during the Ca2+ challenge ROS production was lower in trained mitochondria because of a reduced ability of Ca2+ to increase ROS production through its action on the TCA cycle and/or on several components of the respiratory chain (12).

Together, results from the present study indicate that mitochondria isolated from trained hearts are more resistant to Ca2+-induced PTP opening when energized with succinate. This adaptation could potentially be beneficial to the heart in the setting of ischemia-reperfusion, a situation in which exercise was shown to be protective (9, 10, 13). The mechanisms underlying this increased resistance remain obscure (6, 22, 40) but are apparently not related to changes in endogenous adenylate content, alterations in respiratory chain function, redox state of PNs, or ROS production. One possibility is that this phenomenon is caused by changes in the expression of anti- and proapoptotic members of the Bcl-2 family of proteins known to modulate Ca2+ sensitivity of the PTP. Indeed, the balance between Bcl-2 and BclXL on the one hand and Bax on the other hand was recently shown to be increased after training (34, 54). However, this hypothesis remains to be tested.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by a grant from the Canadian Institute of Health Research. Y. Burelle is a Junior Investigator of the Fonds de Recherche en Santé du Québec (FRSQ). M. Marcil is supported by a PhD scholarship from FRSQ.


   ACKNOWLEDGMENTS
 
The authors thank Dr. Éric Fontaine for helpful discussions and Véronique Giroux and Sébastien Martin for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: Y. Burelle, Univ. de Montréal, Dept. of Kinesiology, PO Box 6128 Centre-Ville, Montreal, PQ, Canada, H3C 3J7 (e-mail : yan.burelle{at}umontreal.ca)

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
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Argaud L, Gateau-Roesch O, Chalabreysse L, Gomez L, Loufouat J, Thivolet-Bejui F, Robert D, and Ovize M. Preconditioning delays Ca2+-induced mitochondrial permeability transition. Cardiovasc Res 61: 115–122, 2004.[Abstract/Free Full Text]
  2. Arieli Y, Gursahani H, Eaton MM, Hernandez LA, and Schaefer S. Gender modulation of Ca2+ uptake in cardiac mitochondria. J Mol Cell Cardiol 37: 507–513, 2004.[CrossRef][ISI][Medline]
  3. Baines CP, Kaiser RA, Purcell NH, Blair NS, Osinska H, Hambleton MA, Brunskill EW, Sayen MR, Gottlieb RA, Dorn GW, Robbins J, and Molkentin JD. Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature 434: 658, 2005.[CrossRef][Medline]
  4. Basso E, Fante L, Fowlkes J, Petronilli V, Forte MA, and Bernardi P. Properties of the permeability transition pore in mitochondria devoid of cyclophilin D. J Biol Chem 280: 18558–18561, 2005.[Abstract/Free Full Text]
  5. Batandier C, Leverve X, and Fontaine E. Opening of the mitochondrial permeability transition pore induces reactive oxygen species production at the level of the respiratory chain complex I. J Biol Chem 279: 17197–17204, 2004.[Abstract/Free Full Text]
  6. Bernardi P. Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev 79: 1127–1155, 1999.[Abstract/Free Full Text]
  7. Bernardi P. Modulation of the mitochondrial cyclosporin A-sensitive permeability transition pore by the proton electrochemical gradient. Evidence that the pore can be opened by membrane depolarization. J Biol Chem 267: 8834–8839, 1992.[Abstract/Free Full Text]
  8. Bernardi P, Vassanelli S, Veronese P, Colonna R, Szabo I, and Zoratti M. Modulation of the mitochondrial permeability transition pore. Effect of protons and divalent cations. J Biol Chem 267: 2934–2939, 1992.[Abstract/Free Full Text]
  9. Bowles DK, Farrar RP, and Starnes JW. Exercise training improves cardiac function after ischemia in the isolated, working rat heart. Am J Physiol Heart Circ Physiol 263: H804–H809, 1992.[Abstract/Free Full Text]
  10. Bowles DK and Starnes JW. Exercise training improves metabolic response after ischemia in isolated working rat heart. J Appl Physiol 76: 1608–1614, 1994.[Abstract/Free Full Text]
  11. Brenner C, Cadiou H, Vieira HL, Zamzami N, Marzo I, Xie Z, Leber B, Andrews D, Duclohier H, Reed JC, and Kroemer G. Bcl-2 and Bax regulate the channel activity of the mitochondrial adenine nucleotide translocator. Oncogene 19: 329–336, 2000.[CrossRef][ISI][Medline]
  12. Brookes PS, Yoon Y, Robotham JL, Anders MW, and Sheu SS. Calcium, ATP, and ROS: a mitochondrial love-hate triangle. Am J Physiol Cell Physiol 287: C817–C833, 2004.[Abstract/Free Full Text]
  13. Brunner M, Moeslinger T, and Spieckermann PG. Regulation of cyclosporin A sensitive mitochondrial permeability transition by the redox state of pyridine nucleotides. Comp Biochem Physiol B Biochem Mol Biol 128: 31–41, 2001.[CrossRef][Medline]
  14. Burelle Y, Wambolt RB, Grist M, Parsons HL, Chow JC, Antler C, Bonen A, Keller A, Dunaway GA, Popov KM, Hochachka PW, and Allard MF. Regular exercise is associated with a protective metabolic phenotype in the rat heart. Am J Physiol Heart Circ Physiol 287: H1055–H1063, 2004.[Abstract/Free Full Text]
  15. Clarke SJ, McStay GP, and Halestrap AP. Sanglifehrin A acts as a potent inhibitor of the mitochondrial permeability transition and reperfusion injury of the heart by binding to cyclophilin-D at a different site from cyclosporin A. J Biol Chem 277: 34793–34799, 2002.[Abstract/Free Full Text]
  16. Crompton M. The mitochondrial permeability transition pore and its role in cell death. Biochem J 341: 233–249, 1999.
  17. Crompton M, Ellinger H, and Costi A. Inhibition by cyclosporin A of a Ca2+-dependent pore in heart mitochondria activated by inorganic phosphate and oxidative stress. Biochem J 255: 357–360, 1988.[ISI][Medline]
  18. Di Lisa F and Bernardi P. Mitochondrial function as a determinant of recovery or death in cell response to injury. Mol Cell Biochem 184: 379–391, 1998.[CrossRef][ISI][Medline]
  19. Di Lisa F, Menabo R, Canton M, Barile M, and Bernardi P. Opening of the mitochondrial permeability transition pore causes depletion of mitochondrial and cytosolic NAD+ and is a causative event in the death of myocytes in postischemic reperfusion of the heart. J Biol Chem 276: 2571–2575, 2001.[Abstract/Free Full Text]
  20. Drew B and Leeuwenburgh C. Method for measuring ATP production in isolated mitochondria: ATP production in brain and liver mitochondria of Fischer-344 rats with age and caloric restriction. Am J Physiol Regul Integr Comp Physiol 285: R1259–R1267, 2003.[Abstract/Free Full Text]
  21. Emaus RK, Grunwald R, and Lemasters JJ. Rhodamine 123 as a probe of transmembrane potential in isolated rat-liver mitochondria: spectral and metabolic properties. Biochim Biophys Acta 850: 436–448, 1986.[Medline]
  22. Fontaine E, Eriksson O, Ichas F, and Bernardi P. Regulation of the permeability transition pore in skeletal muscle mitochondria. Modulation by electron flow through the respiratory chain complex I. J Biol Chem 273: 12662–12668, 1998.[Abstract/Free Full Text]
  23. Griffiths EJ and Halestrap AP. Mitochondrial non-specific pores remain closed during cardiac ischaemia, but open upon reperfusion. Biochem J 307: 93–98, 1995.
  24. Griffiths EJ and Halestrap AP. Protection by Cyclosporin A of ischemia/reperfusion-induced damage in isolated rat hearts. J Mol Cell Cardiol 25: 1461–1469, 1993.[CrossRef][ISI][Medline]
  25. Halestrap AP, Doran E, Gillespie JP, and O'Toole A. Mitochondria and cell death. Biochem Soc Trans 28: 170–177, 2000.[ISI][Medline]
  26. Harris EJ, Booth R, and Cooper MB. The effect of superoxide generation on the ability of mitochondria to take up and retain Ca2+. FEBS Lett 146: 267–272, 1982.[CrossRef][ISI][Medline]
  27. Hausenloy DJ, Maddock HL, Baxter GF, and Yellon DM. Inhibiting mitochondrial permeability transition pore opening: a new paradigm for myocardial preconditioning? Cardiovasc Res 55: 534–543, 2002.[Abstract/Free Full Text]
  28. Hausenloy DJ, Yellon DM, Mani-Babu S, and Duchen MR. Preconditioning protects by inhibiting the mitochondrial permeability transition. Am J Physiol Heart Circ Physiol 287: H841–H849, 2004.[Abstract/Free Full Text]
  29. Hunter DR and Haworth RA. The Ca2+-induced membrane transition in mitochondria. I. The protective mechanisms. Arch Biochem Biophys 195: 453–459, 1979.[CrossRef][ISI][Medline]
  30. Ichas F, Jouaville LS, and Mazat JP. Mitochondria are excitable organelles capable of generating and conveying electrical and calcium signals. Cell 89: 1145–1153, 1997.[CrossRef][ISI][Medline]
  31. Javadov SA, Clarke S, Das M, Griffiths EJ, Lim KH, and Halestrap AP. Ischaemic preconditioning inhibits opening of mitochondrial permeability transition pores in the reperfused rat heart. J Physiol 549: 513–524, 2003.[Abstract/Free Full Text]
  32. Javadov SA, Lim KH, Kerr PM, Suleiman MS, Angelini GD, and Halestrap AP. Protection of hearts from reperfusion injury by propofol is associated with inhibition of the mitochondrial permeability transition. Cardiovasc Res 45: 360–369, 2000.[Abstract/Free Full Text]
  33. Ji LL, Fu RG, Mitchell EW, Griffiths M, Waldrop TG, and Swartz HM. Cardiac hypertrophy alters myocardial response to ischaemia and reperfusion in vivo. Acta Physiol Scand 151: 279–290, 1994.[ISI][Medline]
  34. Kang PM, Yue P, Liu Z, Tarnavski O, Bodyak N, and Izumo S. Alterations in apoptosis regulatory factors during hypertrophy and heart failure. Am J Physiol Heart Circ Physiol 287: H72–H80, 2004.[Abstract/Free Full Text]
  35. Kerr PM, Suleiman MS, and Halestrap AP. Reversal of permeability transition during recovery of hearts from ischemia and its enhancement by pyruvate. Am J Physiol Heart Circ Physiol 276: H496–H502, 1999.[Abstract/Free Full Text]
  36. Kihlstrom M. Protection effect of endurance training against reoxygenation-induced injuries in rat heart. J Appl Physiol 68: 1672–1678, 1990.[Abstract/Free Full Text]
  37. Kim JS, He L, and Lemasters JJ. Mitochondrial permeability transition: a common pathway to necrosis and apoptosis. Biochem Biophys Res Commun 304: 463, 2003.[CrossRef][ISI][Medline]
  38. Kowaltowski AJ, Vercesi AE, and Fiskum G. Bcl-2 prevents mitochondrial permeability transition and cytochrome c release via maintenance of reduced pyridine nucleotides. Cell Death Differ 7: 903–910, 2000.[CrossRef][ISI][Medline]
  39. Lemasters J, Qian T, Bradham C, Brenner D, Cascio W, Trost L, Nishimura Y, Nieminen AL, and Herman B. Mitochondrial dysfunction in the pathogenesis of necrotic and apoptotic cell death. J Bioenerg Biomembr 31: 305–319, 1999.[CrossRef][ISI][Medline]
  40. Leverve XM and Fontaine E. Role of substrates in the regulation of mitochondrial function in situ. IUBMB Life 52: 221–229, 2001.[ISI][Medline]
  41. Mattson MP and Kroemer G. Mitochondria in cell death: novel targets for neuroprotection and cardioprotection. Trends Mol Med 9: 196–205, 2003.[CrossRef][ISI][Medline]
  42. McElroy CL, Gissen SA, and Fishbein MC. Exercise-induced reduction in myocardial infarct size after coronary artery occlusion in the rat. Circulation 57: 958–962, 1978.[Abstract/Free Full Text]
  43. Moore RL and Palmer BM. Exercise training and cellular adaptations of normal and diseased hearts. Exerc Sport Sci Rev 27: 285–315, 1999.[Medline]
  44. Murphy RC, Schneider E, and Kinnally KW. Overexpression of Bcl-2 suppresses the calcium activation of a mitochondrial megachannel. FEBS Lett 497: 73–76, 2001.[CrossRef][ISI][Medline]
  45. Nakagawa T, Shimizu S, Watanabe T, Yamaguchi O, Otsu K, Yamagata H, Inohara H, Kubo T, and Tsujimoto Y. Cyclophilin D-dependent mitochondrial permeability transition regulates some necrotic but not apoptotic cell death. Nature 434: 652–658, 2005.[CrossRef][Medline]
  46. Narita M, Shimizu S, Ito T, Chittenden T, Lutz RJ, Matsuda H, and Tsujimoto Y. Bax interacts with the permeability transition pore to induce permeability transition and cytochrome c release in isolated mitochondria. Proc Natl Acad Sci USA 95: 14681–14686, 1998.[Abstract/Free Full Text]
  47. Nicolli A, Basso E, Petronilli V, Wenger RM, and Bernardi P. Interactions of cyclophilin with the mitochondrial inner membrane and regulation of the permeability transition pore, and cyclosporin A-sensitive channel. J Biol Chem 271: 2185–2192, 1996.[Abstract/Free Full Text]
  48. Nicolli A, Petronilli V, and Bernardi P. Modulation of the mitochondrial cyclosporin A-sensitive permeability transition pore by matrix pH. Evidence that the pore open-closed probability is regulated by reversible histidine protonation. Biochemistry 32: 4461–4465, 1993.[CrossRef][Medline]
  49. Pastorino JG, Tafani M, Rothman RJ, Marcinkeviciute A, Hoek JB, Farber JL, and Marcineviciute A. Functional consequences of the sustained or transient activation by Bax of the mitochondrial permeability transition pore. J Biol Chem 274: 31734–31739, 1999.[Abstract/Free Full Text]
  50. Penpargkul S, Schwartz A, and Scheuer J. Effect of physical conditioning on cardiac mitochondrial function. J Appl Physiol 45: 978–986, 1978.[Free Full Text]
  51. Powers SK, Demirel HA, Vincent HK, Coombes JS, Naito H, Hamilton KL, Shanely RA, and Jessup J. Exercise training improves myocardial tolerance to in vivo ischemia-reperfusion in the rat. Am J Physiol Regul Integr Comp Physiol 275: R1468–R1477, 1998.[Abstract/Free Full Text]
  52. Schinzel AC, Takeuchi O, Huang Z, Fisher JK, Zhou Z, Rubens J, Hetz C, Danial NN, Moskowitz MA, and Korsmeyer SJ. Cyclophilin D is a component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia. Proc Natl Acad Sci USA 102: 12005–12010, 2005.[Abstract/Free Full Text]
  53. Servais S, Couturier K, Koubi H, Rouanet JL, Desplanches D, Sornay-Mayet MH, Sempore B, Lavoie JM, and Favier R. Effect of voluntary exercise on H2O2 release by subsarcolemmal and intermyofibrillar mitochondria. Free Radic Biol Med 35: 24–32, 2003.[CrossRef][ISI][Medline]
  54. Siu PM, Bryner RW, Martyn JK, and Alway SE. Apoptotic adaptations from exercise training in skeletal and cardiac muscles. FASEB J 18: 1150–1152, 2004.[Abstract/Free Full Text]
  55. Somani SM, Frank S, and Rybak LP. Responses of antioxidant system to acute and trained exercise in rat heart subcellular fractions. Pharmacol Biochem Behav 51: 627–634, 1995.[CrossRef][ISI][Medline]
  56. Sordahl LA, Asimakis GK, Dowell RT, and Stone HL. Functions of selected biochemical systems from the exercised-trained dog heart. J Appl Physiol 42: 426–431, 1977.[Abstract/Free Full Text]
  57. Turrens JF. Superoxide production by the mitochondrial respiratory chain. Biosci Rep 17: 3–8, 1997.[CrossRef][ISI][Medline]
  58. Venditti P, Masullo P, and Di Meo S. Effect of training on H2O2 release by mitochondria from rat skeletal muscle. Arch Biochem Biophys 372: 315–320, 1999.[CrossRef][ISI][Medline]
  59. Zamzami N and Kroemer G. Apoptosis: mitochondrial membrane permeabilization—the (w)hole story? Curr Biol 13: R71–R73, 2003.[CrossRef][ISI][Medline]
  60. Zoratti M and Szabo I. The mitochondrial permeability transition. Biochim Biophys Acta 1241: 139–176, 1995.[Medline]
  61. Zoratti M, Szabo I, and De Marchi U. Mitochondrial permeability transitions: how many doors to the house? Biochim Biophys Acta 1706: 40–52, 2005.[Medline]



This article has been cited by other articles:


Home page
 Am. J. Physiol. Regul. Integr. Comp. Physiol.Home page
M. Picard, K. Csukly, M.-E. Robillard, R. Godin, A. Ascah, C. Bourcier-Lucas, and Y. Burelle
Resistance to Ca2+-induced opening of the permeability transition pore differs in mitochondria from glycolytic and oxidative muscles
Am J Physiol Regulatory Integrative Comp Physiol, August 1, 2008; 295(2): R659 - R668.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
J. W. Starnes, B. D. Barnes, and M. E. Olsen
Exercise training decreases rat heart mitochondria free radical generation but does not prevent Ca2+-induced dysfunction
J Appl Physiol, May 1, 2007; 102(5): 1793 - 1798.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
290/4/H1549    most recent
00913.2005v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (5)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Marcil, M.
Right arrow Articles by Burelle, Y.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Marcil, M.
Right arrow Articles by Burelle, Y.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2006 by the American Physiological Society.