| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
MUSCLE CELL BIOLOGY AND CELL MOTILITY
1Department of Zoology, La Trobe University, Melbourne, Australia; and 2Institute of Physiology and Biophysics, University of Aarhus, Århus, Denmark
Submitted 30 August 2006 ; accepted in final form 18 November 2006
 |
ABSTRACT |
|---|
 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
excitation-contraction coupling; mechanically skinned fiber; physiological temperature
Recently, it has been demonstrated that intracellular levels of reactive oxygen species (ROS), in particular O2·–, are increased as the temperature of rat diaphragm was raised from 25 to 43°C (41). This increase in intracellular O2·– was accompanied by an increase in extracellular O2·–. O2·– production in whole muscle tissue may arise not only from muscle fibers but also from other cellular structures such as nerve terminals, blood cells, and capillaries (38). Furthermore, in addition to mitochondria, there are a number of other sources of O2·– production involving 5-lipoxygenase, cyclooxygenase, xanthine oxidase, and NAD(P)H oxidase (41). Indeed, through the use of specific mitochondrial electron transport chain blockers, it has been hypothesized that the major site of intracellular O2·– at elevated temperatures was the sarcolemmal NAD(P)H oxidase (41). However, in that study (41), the respiratory chain was blocked with rotenone, which has been shown to increase the mitochondrial production of O2·– and not decrease it (23, 36), as these authors hypothesized (41). Moreover, recent studies on isolated muscle mitochondria have unequivocally shown that considerable amounts of O2·– produced in muscle mitochondria can be extruded to the environment surrounding the mitochondria (23, 36), challenging a long-standing view that effectively all O2·– produced by the mitochondria is dismutated by mitochondrial superoxide dismutases (SODs) (23).
ROS and in particular O2·–, have been shown to play a crucial role in skeletal muscle function (9, 20, 22, 24, 31). Application of O2·– to diaphragm muscle strips resulted in a reduced maximum Ca2+ activated force response and inhibition of sarcoplasmic reticulum (SR) Ca2+ release (5). It has also been demonstrated that, while O2·– can depress maximum force, it has no apparent effect on the sensitivity of the contractile apparatus to Ca2+ (4, 39). Similar results were obtained on cardiac muscle where O2·– production decreased force but did not affect Ca2+ sensitivity or SR function (21).
On the basis of these findings, and the fact that, at or above 37°C, there is a marked loss in function of the isolated muscle (16), we hypothesized that the loss of skeletal muscle function at or above 37°C must be connected to the increase in the amount of O2·– produced by the mitochondria in the skeletal muscle fibers. Average rat hind muscle temperature varies between 35.5 and 37.2°C when the rat is at rest and between 36.2 and 38.5°C after 15 min of level walking (6). To avoid potential sources of O2·– production outside muscle fibers, O2·– production was measured at 37°C in mechanically skinned muscle fibers devoid of the sarcolemma, split single muscle fibers, where part of the sarcolemma remained attached, and small bundles of isolated rat skeletal muscle fibers at rest, in the presence and absence of modifiers of mitochondrial function. The results clearly show that mitochondria are the major site of O2·– production in the myoplasmic environment of skeletal muscle fibers kept at 37°C and that the excitability of the muscle fibers at 37°C can be greatly improved in the presence of a potent, membrane-permeant SOD mimetic, suggesting a link between mitochondrial O2·– production and fiber excitability. Thus, the findings provide concrete evidence that the mitochondria are the principal source of O2·– radicals in muscle at normal body core temperature and that it is these O2·– radicals that interfere with aspects of excitation-contraction coupling (ECC) to cause depression of the force response to excitation.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
The dimensions of each preparation were measured in at least five points along the preparation on a television monitor connected to a CCD camera attached to a dissecting microscope to calculate fiber volume and fiber mass assuming a density of 1.03 g/ml, as previously described (39). The volume of the preparations for small intact muscle fiber bundles, mechanically skinned fibers, and split fibers ranged between 129 ± 9 (n = 7), 38 ± 3 (n = 10), and 44 ± 8 (n = 9) nl, respectively.
The preparations were first equilibrated for 5 min at 37°C with 5 µM (oxidized) cytochrome c in either a modified KRS for intact fibers or relaxing solution for mechanically skinned and split fibers, before the preparations were transferred to 1 ml of an identical solution in which they were incubated for 75 min (see Table 1 for composition of solutions).
O2·–measurements.
The cytochrome c assay (see below) was used to measure O2·– together with other molecular species that can reduce the cytochrome c (Fe3+) to cytochrome c (Fe2+) (26, 39). Cytochrome c is membrane impermeant and therefore can be reduced only by molecular species present in the same compartment. The method is based on the reaction of O2·– with oxidized cytochrome c (Fe3+) to produce reduced cytochrome c (Fe2+), which has a specific peak absorbance at 550 nm. This permits spectrophotometrical determination of reduced cytochrome c (Fe2+), and on the basis of results presented (see 
Fig. 2 and text), most cytochrome c reduction at 37°C in the resting fiber preparations was due to the reaction with O2·–. Consequently, the amount of O2·– liberated was determined spectrophotometrically from the specific change in cytochrome c absorbance at 550 nm after reduction of oxidized cytochrome c (Fe3+) to cytochrome c (Fe2+), as previously described (39). In all experiments, great care was taken to correct for any changes induced on the cytochrome c assay by factors other than the muscle preparation per se, such as addition of rotenone and succinate to the solutions. Therefore, blank tests without muscle preparations were run in parallel with all proper experiments to correct for any possible effects of such factors on the cytochrome c assay.
|
|
Modifiers of mitochondrial function. To distinguish between extracellular and intracellular O2·– production and the involvement of the mitochondria, several strategies targeting mitochondria, which remain intact in the mechanically skinned and split muscle fibers, were used. Succinate (5 mM), in the presence of ATP, but in the absence of ADP and inorganic phosphate, were added to solutions to increase O2·– production. Succinate markedly increases O2·– production in mitochondria through reverse electron transport from succinate through complex II into complex I, the major site for O2·– production in skeletal muscle mitochondria of the rat (14, 15, 36).
Complex I of the mitochondrial electron transport chain was inhibited with rotenone (50 µM), while mitochondria were fed succinate, to prevent reverse electron transport into complex I and to determine whether cytochrome c reduction decreased, as the rate of O2·– production from succinate dehydrogenase, complex III, and other downstream sites is almost negligible during reverse electron transport in rat skeletal muscle mitochondria (14, 15, 36).
To further characterize mitochondrial involvement as a source of O2·–, causing cytochrome c reduction measured in skinned fibers, we applied rotenone (50 µM) alone (in the presence of ATP) and 5 µM antimycin A (an inhibitor of complex III of electron transport chain) to the skinned-fiber preparations.
Paired experiments were performed with intact and mechanically skinned fiber preparations in the presence and in the absence of modifiers of mitochondrial function. Results were averaged for the same day and expressed as a percentage of mechanically skinned-fiber preparation on the same day and from the same muscle.
Force measurements. To determine whether increased production of O2·– in mitochondria at 37°C depresses skeletal muscle function, experiments were performed on mechanically skinned fibers, which can be activated at any step in the ECC process (29). For example, the mechanically skinned fibers can be electrically stimulated in physiological solutions mimicking the myoplasm by triggering action potentials in the sealed t-system, eliciting twitch and tetanic force responses (27, 29, 30). Mechanically skinned fibers from Long-Evans rats were attached to a force transducer (SensoNor 801, Horten, Norway) and bathed in a solution mimicking the intracellular environment (13, 27, 29, 30) (similar to that described above for skinned fibers, except that 49.95 mM EGTA was replaced by 49.95 mM HDTA and [Ca2+] = 100 nM, instead of <1 nM). Fibers were initially equilibrated at room temperature for 2 min and then transferred to equivalent solutions at 37°C with and without a O2·– scavenger. A single electrical stimulation of 2-ms duration, at 50 V/cm, was triggered every 1 min until twitch force decreased to very low values (<2% of initial twitch force).
Following the loss of the twitch response, the SR Ca2+ content was assessed by releasing all Ca2+ in the SR by placing the fiber in a solution containing very low [Mg2+] (0.015 mM), which causes opening of the SR Ca2+ release channels (17). The preparation was then maximally Ca2+ activated by placing the fiber in a solution of 30 µM [Ca2+] buffered with 50 mM EGTA.
Electrophysiological measurements.
Intracellular recordings of the membrane potential were made with microelectrodes filled with 3 M KCl (10–25 M
) in the outer layer of fibers of EDL muscles (from Wistar rats) placed in KRS solution at 25°C, as previously described (19). Some muscles were incubated at 37°C for 30 min, while others serving as controls, were kept at 25°C throughout. Action potentials (AP) were triggered indirectly via nerve stimulation using 5 mA constant current pulses, and were recorded using an Axoclamp-2a amplifier connected to a computer, using Signal 2.09 software (Cambridge Electronic Design, Cambridge, UK). The resting membrane potential (RMP) was measured from the baseline of the AP, and AP amplitude was defined as the difference between RMP and the peak potential of the AP.
Chemicals. All chemicals, unless otherwise noted, were obtained from Sigma (St. Louis, MO).
Statistical analyses. Results are expressed as means ± SE, and statistical analyses were performed using the scientific analysis program GraphPad Prism (GraphPad Software, San Diego, CA). Statistical significance was tested at P < 0.05 levels using one-way ANOVA (with Kruskal-Wallis multiple-comparison test) and Student's t-test as appropriate.
|
RESULTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
When fibers were left intact and incubated at 37°C for 75 min, the amount of reducing equivalents/mg fiber measured extracellularly with the cytochrome c assay was only 26.9 ± 6.3% (n = 7) of the amount of reducing equivalents/mg fiber detected in skinned fibers from same muscles (Fig. 1). This corresponds to an average rate of 12.8 pmol reducing equivalents·mg–1·min–1, which is close to the reported rate values measured extracellularly with different techniques in mammalian skeletal muscles at rest at 37°C (26).
Rate of cytochrome c reduction in presence of modifiers of mitochondrial O2·– production. To determine the source of O2·– production, we applied succinate, which, in purified rat mitochondria, is known to markedly increase O2·– production (36). When 5 mM succinate was present in the relaxing solution in which mechanically skinned fibers were incubated, a large increase in the rate of cytochrome c reduction was observed compared with controls [14.56 ± 7.73 nmol reducing equivalents·mg–1·h–1 (n = 5), vs. 2.84 ± 0.69 nmol reducing equivalents·mg–1·h–1, (n = 10), P < 0.05]. The large error associated with measurements in the presence of succinate was most likely related to the very high variation in succinate dehydrogenase activities in rat muscle fibers in general (25). Consistent with this, removal of one outlier reduced markedly the size of the error for these measurements (11.49 ± 3.93 nmol reducing equivalents·mg–1·h–1, n = 4) (Fig. 2A). Another strategy was to inhibit complex I with rotenone, while mitochondria were fed succinate, to prevent reverse electron transport into complex I and determine whether cytochrome c reduction decreased. Under these conditions, no net cytochrome c reduction could be measured (–0.25 ± 0.25 nmol reducing equivalents·mg–1·h–1; n = 8). Thus, when complex I was blocked with rotenone, effectively, no reduction of cytochrome c could be detected in the presence of succinate.
Because rotenone completely abolishes O2·– production in succinate respiring mitochondria (36), the results strongly suggest that effectively all measured cytochrome c reduction in our experiments is associated with O2·– production in the mitochondria. In the presence of either rotenone or antimycin A, there was a marked increase in the rate of cytochrome c reduction (5.34 ± 0.44 and 8.97 ± 1.21 nmol reducing equivalents·mg–1·h–1, respectively, see Fig. 2A), in agreement with observations reported on isolated rat skeletal muscle mitochondria (15, 36).
To find out whether NAD(P)H-dependent plasma membrane oxidase activity contributed in any significant way to O2·– release in the myoplasmic environment, experiments were performed in the presence of succinate, rotenone, and 0.1 mM NAD(P)H, which is in the upper physiological range for the rat EDL muscle (34). The rate of cytochrome c reduction in this experiment was –0.59 ± 0.35 nmol reducing equivalents·mg–1·h–1 (n = 3), which is not statistically different from the rate of cytochrome c reduction in the presence of succinate and rotenone without NAD(P)H (–0.25 ± 0.25 nmol reducing equivalents·mg–1·h–1, t-test) but significantly smaller than under control conditions (2.84 ± 0.69 nmol reducing equivalents·mg–1·h–1, n = 10), (one-way ANOVA P < 0.05). Considering that plasma membrane NAD(P)H oxidase activity (NOX) is associated with the tubular system in skeletal muscle (8), this result suggests that compared with mitochondria, the plasma membrane NAD(P)H oxidase activity is not a major source of O2·– production.
Interestingly, when 5 mM succinate was added to the incubating solution for intact fibers, the amount of cytochrome c reducing equivalents measured in the extracellular solution increased by 90.6 ± 20.5%, and further addition of 50 µM rotenone to the incubating solution for intact fibers again abolished cytochrome c reduction in the extracellular solution (Fig. 2B; P < 0.05).
O2·– effect on single fiber function. After 7 min at 37°C, the amplitude of single twitches induced at 1-min intervals in single mechanically skinned fibers dropped greatly by 83 ± 10% of the initial response, compared with a drop of only 12 ± 4% at 25°C (Fig. 3, A and C, respectively). At 8 min at 37°C, there was complete loss of twitch responses in five out of the six fibers tested. To test whether this decrease in force was associated with O2·– production, a membrane-permeable SOD mimetic Tempol (1 mM), which effectively removes O2·– without being used as a substrate (12), was applied. In the presence of 1 mM Tempol, twitch responses decreased markedly less, by only 39 ± 9% (Fig. 3B, P = 0.017) after 7 min at 37°C.
|
|
10 mV (P < 0.001) compared with controls when the muscle fibers were maintained at 25°C (Table 2). Furthermore, the amplitude of the AP decreased by 20 mV (P < 0.001) after exposure to 37°C compared with controls. In the presence of 1 mM Tempol, no significant reduction of the RMP occurred after exposure to 37°C (P < 0.001), and the AP amplitude was only marginally reduced compared with controls (Table 2). |
DISCUSSION |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Mitochondria as the major source of cytosolic and extracellular O2·– in muscle. A deeper understanding of O2·– production in the skeletal rat mitochondria in the presence and absence of various mitochondrial inhibitors and different substrates was only recently achieved (14, 15, 23, 36). It has been comprehensively established that the major site for O2·– production in mitochondria is associated with complex I and that inhibitors of the quinone-binding site such as rotenone, allow rapid release of O2·– on the matrix side, particularly in the presence of ATP, which helps establish the pH and the electrical potential gradients across the inner mitochondrial membranes. Furthermore, it was demonstrated that in the presence of succinate, there is a very large increase in the rate of O2·– production by complex I on the matrix side through reverse electron transport from succinate through complex II into complex I and that this very large rate of O2·– production is effectively abolished when rotenone is also present. On this background, our cytochrome c reduction measurements in mechanically skinned fibers incubated with succinate in the presence or absence of rotenone, reflect faithfully the changes in O2·– production in the mitochondria. Moreover, the marked increase in cytochrome c reduction in the presence of antimycin A (Fig. 2A) is in full accord with the observation and the interpretation of St-Pierre and colleagues (36) and Muller and colleagues (23) that center o of the complex III is also capable of producing large amounts of O2·– when the electron transport from center o to center i on complex III is blocked by antimycin A. Thus, mitochondrial O2·– production by complex III in the intermembrane space, which can be sampled outside the mitochondria (23, 36), can fully explain the observations of Zuo and colleagues (42) and also reproduced in this study (Fig. 2A). Therefore, rather than rule out mitochondria as the source of the O2·– in rat skeletal muscle, the results of Zuo and colleagues (42) actually fully support the tight link between measured O2·– changes and mitochondrial changes in O2·– production.
The addition of succinate to the extracellular environment is known to increase the rate of respiration of intact muscle fibers from the cat diaphragm (3), indicating that succinate is transported across the plasma membrane of mammalian skeletal muscle. Thus, the results with intact fibers (Fig. 2B) follow the pattern observed in skinned fibers (Fig. 2A). The results on intact fibers further indicate that the contribution of the surface and t-system membranes to the reduction of cytochrome c measured extracellularly at 37°C must be a small fraction of that measured in the absence of succinate. This is because little O2·– could be detected outside intact fibers in the presence of succinate and rotenone (Fig. 2B), a condition that is known to abolish O2·– production in mitochondria (36). This is in agreement with recent results in which the rise in oxidant activity during contraction was shown to be of intracellular origin (20). Taken together, the data suggest that at 37°C, most molecular species that reduce cytochrome c, including O2·–, are produced intracellularly and are associated with O2·– production by mitochondria.
The plasma membrane of the t-system in mechanically skinned fibers represents the major interface between the intracellular and extracellular environments of an intact muscle fiber (37), and it has been recently shown that the t-system has NOX activity (8). Without NAD(P)H in solutions there is no NOX activity in the mechanically skinned and split muscle fibers. Therefore, it was of interest to measure NOX activity in mechanically skinned fibers in the presence of 0.1 mM NADH, which is in the upper physiological range for the rat EDL muscle (34). Results show that when mitochondrial O2·– production was blocked in the presence of succinate and rotenone, the addition of 0.1 mM NADH to the solutions did not produce any measurable amount of O2·–, indicating that NOX activity was under the detection limit of our method, which could otherwise reliably detect mitochondrial dependent O2·– in the myoplasmic environment. The NOX activity (3.6 nmol·mg triad protein–1·min–1) measured in skeletal muscle triads at 37°C in the presence of 0.1 mM NADH (8) can also be used to estimate the contribution of t-system dependent NOX activity in muscle fibers. Considering a ratio of 2,000 between the wet muscle mass and the triad protein mass (11), the t-system NOX activity for O2·– production amounts to 0.11 nmol·mg fiber–1·h–1, which is only about 4% of the measured rate in the skinned muscle fiber at rest or 1% of the rate measured in the presence of succinate. These two sets of results are fully consistent with each other and indicate that NOX activity produces only a small fraction of O2·– produced by the muscle fiber and, therefore, compared with mitochondria, the plasma membrane NOX activity is not a major source of O2·– production.
Although the mechanism by which some O2·– crosses the mitochondrial and plasma membranes in skeletal muscles is not clear, O2·– is in rapid equilibrium with its protonated and uncharged form (·OOH) (about 1% of total O2·– at pH 7) that can readily cross membranes (10). Considering that the pH in the intermembrane space is more acidic than that in the surrounding environment, this would increase the fraction of protonated O2·–, facilitating its diffusion across the outer mitochondrial membrane. Furthermore, some O2·– from the intermembrane space may be carried across the outer mitochondrial membrane to the myoplasmic environment via voltage-dependent anion channels (7). Considering now the spatial arrangement of mitochondria in the fast-twitch rat EDL muscle, they are situated in very close proximity of the t-system (27), which opens to the extracellular space, with the membranes of the two systems running in parallel. This arrangement may provide a pathway for O2·– release to the extracellular environment, such that a small fraction of the mitochondrial O2·– could be sampled in the extracellular environment even in the presence of high mitochondrial SOD activity.
O2·– and muscle excitability. It is important to note that all diffusible O2·– scavengers and SOD found initially in the myoplasm of intact muscle fibers are lost after the fiber is mechanically skinned and placed in an effectively infinite volume of solution. This renders the skinned fiber preparation more sensitive than the intact fiber to the influence of exogenously added O2·– scavengers. Here, we show that the use of 1 mM Tempol, which is a SOD mimetic, is able to markedly attenuate the decrease of the twitch response in mechanically skinned fibers electrically stimulated at 37°C (Fig. 3C). This provides strong evidence that at least part of the observed decrease in the twitch force response at 37°C (Fig. 3B) is due to mitochondrial O2·– production.
Furthermore, as shown in Fig. 3C, reducing O2·– production in the muscle fiber by reducing the temperature from 37°C to 25°C markedly attenuated the decline in the twitch response in electrically stimulated mechanically skinned fibers. A marked reduction in cytochrome c reduction when temperature decreased from 37°C to 23°C was reported for rat diaphragm muscle (2), and other studies have shown that most of the temperature-induced increase in cytochrome c reduction measured extracellularly in muscle is due to O2·– production (42). Also, the rate of cytochrome c reduction measured extracellularly in rat EDL muscle at rest was much smaller at 25°C (0.0033 ± 0.0005 nmol reducing equivalents·mg–1·h–1) (39) than that measured here at 37°C (0.76 nmol reducing equivalents·mg–1·h–1).
Electrophysiological measurements obtained from this study provide strong evidence that O2·– production in skeletal muscle at 37°C reduces the excitability of the muscle fibers by causing membrane depolarization, which, in turn, causes slow inactivation of the Na+ channels (33), reduction in the amplitude of the action potential, and impaired AP propagation along the t-system (35), as these effects could be largely prevented in the presence of 1 mM Tempol. O2·– altering a step in ECC before SR Ca2+ release is further evidenced by results showing that fibers that can no longer produce any twitch force at 37°C, retain a normal SR Ca2+ content and ability to produce force (Table 2). Separate experiments further showed that neither the SR Ca2+ handling properties nor the sensitivity to Ca2+ of the contractile apparatus were affected by exposure to 40°C for up to 10 min (40). Considering that previous results from this laboratory have already provided evidence that the functional state of mitochondria in rat fibers is linked to the excitability of the t-system (27), these observations may suggest that O2·– may be involved in the feedback mechanism between mitochondria and plasma membranes. Thus, if mitochondria increase the rate of O2·– production in response to different stressors, such as a rise in temperature, this then causes depolarization of the t-system and consequently a decrease of excitability, which ultimately causes a reduction in the force response and a lower rate of ATP utilization, thereby reducing the demand for ATP production placed on mitochondria.
General remarks. In conclusion, this study shows that mitochondria are the principal source of O2·– radicals in muscle fibers at normal body core temperature and that it is these O2·– radicals that interfere with aspects of ECC in mechanically skinned fibers to cause depression of the force response to excitation at sites that lie upstream of the myofilaments and SR. Although the presence of various cytosolic scavengers and antioxidants in the intact fiber would attenuate the effects of O2·– radicals produced by mitochondria on ECC seen in mechanically skinned fibers, the results nevertheless indicate that mitochondria can be involved in modulating muscle function by processes that are not directly related to their main function of producing ATP.
The results are also physiologically relevant in explaining various conditions associated with increased O2·– production in muscle. For example, O2·– released by muscles during fever or exercise and entering the bloodstream could also help the phagocytes in eliciting their immune response (1). Furthermore, reperfusion injury can be related to the increased O2·– production in muscle under such conditions (28).
|
GRANTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
FOOTNOTES |
|---|
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 |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
2. Arbogast S, Reid MB. Oxidant activity in skeletal muscle fibers is influenced by temperature, CO2 level, and muscle-derived nitric oxide. Am J Physiol Regul Integr Comp Physiol 287: R698–R705, 2004.
3. Baum H, El-Khangary HA, Moore RE. Catecholamine-stimulated thermogenesis: some factors influencing the pattern of respiration of isolated kitten diaphragms. J Physiol 169: 237–248, 1963.
4. Callahan LA, She ZW, Nosek TM. Superoxide, hydroxyl radical, and hydrogen peroxide effects on single-diaphragm fiber contractile apparatus. J Appl Physiol 90: 45–54, 2001.
5. Darnley GM, Duke AD, Steele DS, MacFarlane NG. Effects of reactive oxygen species on aspects of excitation-contraction coupling in chemically skinned rabbit diaphragm muscle fibres. Exp Physiol 86: 161–168, 2001.[Abstract]
6. Delp MD, Duan C, Ray CA, Armstrong RB. Rat hindlimb muscle blood flow during level and downhill locomotion. J Appl Physiol 86: 546–568, 1999.
7. Han D, Antunes F, Canali R, Rettori D, Cadenas E. Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol. J Biol Chem 278: 5557–5563, 2003.
8. Hidalgo C, Sanchez G, Barrientos P, Aracena-Parks G. A transverse tubule NADPH oxidase activity stimulates calcium release from isolated triads via RYR1 S-glutathionylation. J Biol Chem 281: 26473–26482, 2006.
9. Kolbeck RC, She Z, Callahan LA, Nosek TN. Increased superoxide production during fatigue in the perfused rat diaphragm. Am J Respir Crit Care Med 156: 140–145, 1997.
10. Korshunov SS, Imlay JA. A potential role for periplasmic superoxide dismutase in blocking the penetration of external superoxide into the cytosol of Gram-negative bacteria. Mol Microbiol 43: 95–106, 2002.[CrossRef][Web of Science][Medline]
11. Kramer JW, Ferguson DG, Corbett AM. Enrichment of triadic and terminal cisternae vesicles from rabbit skeletal muscle. J Membr Biol 195: 9–20, 2003.[CrossRef][Web of Science][Medline]
12. Krishna MC, Russo A, Mitchell JB, Goldstein S, Dafni H, Samuni A. Do nitroxide antioxidants act as scavengers of O2·–. or as SOD mimics? J Biol Chem 271: 26026–26030, 1996.
13. Lamb GD, Stephenson DG. Effects of intracellular pH and [Mg2+] on excitation-contraction coupling in skeletal muscle fibres of the rat. J Physiol 478: 331–339, 1994.
14. Lambert AJ, Brand MD. Superoxide production by NADH: ubiquinone oxidoreductase (complex I) depends on the pH gradient across the mitochondrial inner membrane. Biochem J 382: 511–517, 2004.[CrossRef][Web of Science][Medline]
15. Lambert AJ, Brand MD. Inhibitors of the quinone-binding site allow rapid superoxide production from mitochondrial NADH: ubiquinone oxidoreductase (complex I). J Biol Chem 279: 39414–39420, 2004.
16. Lännergren J, Westerblad H. The temperature dependence of isometric contractions of single, intact fibres dissected from a mouse foot muscle. J Physiol 390: 285–293, 1987.
17. Launikonis BS, Stephenson DG. Effects of Mg2+ on Ca2+ release from the sarcoplasmic reticulum of skeletal muscle fibres from yabby (crustacean) and rat. J Physiol 256: 299–312, 2000.
18. Launikonis BS, Stephenson DG. Osmotic properties of the sealed tubular system of toad and rat skeletal muscle. J Gen Physiol 123: 231–247, 2004.
19. Macdonald WA, Pedersen TH, Clausen T, Nielsen OB. N-Benzyl-p-toluene sulphonamide allows the recording of trains of intracellular action potentials from nerve-stimulated intact fast-twitch skeletal muscle of the rat. Exp Physiol 90: 815–825, 2005.
20. McArdle F, Pattwell DM, Vasilaki A, McArdle A, Jackson MJ. Intracellular generation of reactive oxygen species by contracting skeletal muscle cells. Free Radic Biol Med 39: 651–657, 2005.[CrossRef][Web of Science][Medline]
21. MacFarlane NG, Miller DJ. Depression of peak force without altering calcium sensitivity by the superoxide anion in chemically skinned cardiac muscle of rat. Circ Res 70: 1217–1224, 1992.
22. Moopanar TR, Allen DG. Reactive oxygen species reduce myofibrillar Ca2+ sensitivity in fatiguing mouse skeletal muscle at 37°C. J Physiol 564: 189–199, 2005.
23. Muller FL, Liu Y, Van Remmen H. Complex III releases superoxide to both sides of the inner mitochondrial membrane. J Biol Chem 279: 49064–49073, 2004.
24. Murrant CL, Reid MB. Detection of reactive oxygen and reactive nitrogen species in skeletal muscle. Microsc Res Tech 55: 236–248, 2001.[CrossRef][Web of Science][Medline]
25. Nemeth P, Pette D. Succinate degydrogenase activity in fibres classified by myosin ATPas in three hind limb muscles of the rat. J Physiol 320: 73–80, 1981.
26. Nethery D, Stofan D, Callahan L, Di Marco A, Supinski G. Formation of reactive oxygen species by the contracting diaphragm is PLA(2) dependent. J Appl Physiol 87: 792–800, 1999.
27. Ørtenblad N, Stephenson DG. A novel signalling pathway originating in mitochondria modulates rat skeletal muscle membrane excitability. J Physiol 548: 139–145, 2003.
28. Park JW, Qi WN, Cai Y, Zelko I, Liu JQ, Chen LE, Urbaniak JR, Folz RJ. Skeletal muscle reperfusion injury is enhanced in extracellular superoxide dismutase knockout mouse. Am J Physiol Heart Circ Physiol 289: H181–H187, 2005.
29. Pedersen TH, Nielsen OB, Lamb GD, Stephenson DG. Intracellular acidosis enhances the excitability of working muscle. Science 305: 1144–1147, 2004.
30. Posterino GS, Lamb GD, Stephenson DG. Twitch and tetanic force responses and longitudinal propagation of action potentials in skinned skeletal muscle fibres of the rat. J Physiol 527: 131–137, 2000.
31. Reid MB, Shoji T, Moody MR, Entman ML. Reactive oxygen in skeletal muscle. II. Extracellular release of free radicals. J Appl Physiol 73: 1805–1809, 1992.
32. Reid MB, Durham WJ. Generation of reactive oxygen and nitrogen species in contracting skeletal muscle: potential impact on aging. Ann NY Acad Sci 959: 108–116, 2002.[Web of Science][Medline]
33. Ruff RL. Sodium channel slow inactivation and the distribution of sodium channels on skeletal muscle fibres enable the performance properties of different muscle fibre types. Acta Physiol Scand 156: 159–168, 1996.[CrossRef][Web of Science][Medline]
34. Sahlin K, Katz A. The content of NADH in rat skeletal muscle at rest and after cyanide poisoning. Biochem J 239: 245–248, 1986.[Web of Science][Medline]
35. Sejersted OM, Sjøgaard G. Dynamics and consequences of potassium shifts in skeletal muscle and heart during exercise. Physiol Rev 80: 1411–1481, 2000.
36. St-Pierre J, Buckingham JA, Roebuck SJ, Brand MD. Topology of superoxide production from different sites in the mitochondrial electron transport chain. J Biol Chem 277: 44784–44790, 2002.
37. Stephenson DG. Tubular system excitability: an essential component of excitation-contraction coupling in fast-twitch fibres of vertebrate skeletal muscle. J Muscle Res Cell Motil 27: 259–274, 2006.[CrossRef][Web of Science][Medline]
38. Turrens JF. Mitochondrial formation of reactive oxygen species. J Physiol 552: 335–344, 2003.
39. van der Poel C, Stephenson DG. Reversible changes in Ca2+-activation properties of rat skeletal muscle exposed to elevated physiological temperatures. J Physiol 544: 765–776, 2002.
40. van der Poel C, Stephenson DG. Elevated temperature effects on sarcoplasmic reticulum function in mammalian skeletal muscle fibres. Biophys J Suppl 84: 573A–573A, 2003.
41. Zuo L, Christofi FL, Wright VP, Liu CY, Merola AJ, Berliner LJ, Clanton TL. Intra- and extracellular measurement of reactive oxygen species produced during heat stress in diaphragm muscle. Am J Physiol Cell Physiol 279: C1058–C1066, 2000.
42. Zuo L, Pasniciuc S, Wright VP, Merola AJ, Clanton TL. Sources for superoxide release: lessons from blockade of electron transport, NADPH oxidase, and anion channels in diaphragm. Antioxid Redox Signal 5: 667–675, 2003.[CrossRef][Web of Science][Medline]
This article has been cited by other articles:
|
|
 |
|
  S. P. Cairns and M. I. Lindinger Do multiple ionic interactions contribute to skeletal muscle fatigue? J. Physiol., September 1, 2008; 586(17): 4039 - 4054. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. H. Fitts The cross-bridge cycle and skeletal muscle fatigue J Appl Physiol, February 1, 2008; 104(2): 551 - 558. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. G. Allen, G. D. Lamb, and H. Westerblad Skeletal Muscle Fatigue: Cellular Mechanisms Physiol Rev, January 1, 2008; 88(1): 287 - 332. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. N. Edwards, W. A. Macdonald, C. van der Poel, and D. G. Stephenson O2bullet production at 37{degrees}C plays a critical role in depressing tetanic force of isolated rat and mouse skeletal muscle Am J Physiol Cell Physiol, August 1, 2007; 293(2): C650 - C660. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. van der Poel and D. G. Stephenson Effects of elevated physiological temperatures on sarcoplasmic reticulum function in mechanically skinned muscle fibers of the rat Am J Physiol Cell Physiol, July 1, 2007; 293(1): C133 - C141. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
P < 0.001 compared with controls; ¤P < 0.001 compared with controls; and +P < 0.05 compared with controls.