| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
CELLULAR METABOLISM
John B. Pierce Laboratory and Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut
Submitted 9 August 2005 ; accepted in final form 19 October 2005
 |
ABSTRACT |
|---|
 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
50% less mitochondrial content in WG myofibers. Duplicate experiments conducted with ±exogenous superoxide dismutase revealed striking differences in the topology and/or dismutation of superoxide in WG vs. soleus and RG muscle. When normalized for mitochondrial content, overall H2O2 scavenging capacity is lower in RG and WG fibers, whereas glutathione peroxidase activity, which is largely responsible for H2O2 removal in mitochondria, is similar in all three muscle types. These findings suggest that type II myofibers, particularly type IIB, possess unique properties that potentiate mitochondrial superoxide production and/or release, providing a potential mechanism for the heterogeneous development of mitochondrial dysfunction in skeletal muscle. superoxide; reactive oxygen species; skeletal muscle; respiration; fiber type
Although the loss of mitochondrial function is thought to stem from the cumulative effects of ROS production, the mechanisms governing mitochondrial O2–· formation in the three major types of skeletal myofibers under both physiological and pathophysiological states are unknown. Both complex I and complex III have been identified as sites of O2–· production in mitochondria isolated from rat skeletal muscle (1, 6, 30). However, mitochondria in skeletal muscle are morphologically and functionally heterogeneous, reflecting differences in metabolic demand associated with each specific fiber type as well as local energy demands associated with particular subcellular regions within each myofiber (e.g., subsarcolemmal and intramyofibrillar mitochondria) (19). Moreover, skeletal muscle mitochondria in vivo are arranged in a highly organized, interconnected reticulum or "crystallike" pattern (32, 49), are dependent on cytoskeletal interactions to maintain proper function (53), and, in oxidative fibers, are thought to interact functionally with myofibrils and the sarcoplasmic reticulum, forming energy transfer systems (creatine kinase, adenylate kinase, hexokinase II) that channel ADP to the mitochondria to more efficiently couple energy utilization and oxidative phosphorylation (38).
The extent to which morphological or metabolic properties intrinsic to the three main skeletal muscle fiber types may influence mitochondrial O2–· production has not been explored previously. To this end, we developed a novel in situ approach to examine the factors governing mitochondrial O2–· production under conditions in which the cellular architecture of the myofibers and the structural integrity of the mitochondrial reticulum are maintained in their native state (35). The aim of the present study was to determine whether mitochondrial O2–· production varies among the three major types of myofibers by determining the sites and topology of mitochondrial O2–· production in permeabilized fiber bundles from rat soleus (predominantly type I oxidative fibers), red gastrocnemius (RG, predominantly type IIA oxidative/glycolytic fibers), and white gastrocnemius (WG, predominantly type IIB glycolytic fibers) muscle. Our findings reveal that the rate of mitochondrial free radical leak under both basal and elevated redox state conditions is substantially greater in type II, particularly type IIB, than in type I myofibers, providing a potential mechanistic basis for the mitochondrial dysfunction, atrophy, and loss of type II myofibers that develop with aging (28, 45, 52).
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Preparation of permeabilized muscle fibers.
The technique is partially adapted from previous methods (24, 44). Briefly, small portions (25 mg) of soleus, RG, and WG muscle were dissected and placed in ice-cold buffer X, containing (in mM) 60 K-MES, 35 KCl, 7.23 K2EGTA, 2.77 CaK2EGTA, 20 imidazole, 0.5 DTT, 20 taurine, 5.7 ATP, 15 PCr, and 6.56 MgCl2·6 H2O (pH 7.1, 295 mosmol/kgH2O). The muscle was trimmed of connective tissue and cut down to fiber bundles (2 x 7 mm, 4–8 mg wet wt). With a pair of needle-tipped forceps under a dissecting microscope, fibers were gently separated from one another to maximize surface area of the fiber bundle, leaving only small regions of contact. To permeabilize the myofibers, each fiber bundle was placed in ice-cold buffer X containing 50 µg/ml saponin and incubated on a rotator for 30 min at 4°C. The permeabilized bundles were then placed in ice-cold buffer Z containing (in mM) 110 K-MES, 35 KCl, 1 EGTA, 5 K2HPO4, and 3 MgCl2· 6 H2O, 0.05 pyruvate, and 0.02 malate with 0.5 mg/ml BSA (pH 7.1, 295 mosmol/kgH2O). Permeabilized fibers remained in buffer Z on a rotator at 4°C until analysis (<90 min) without any deterioration in mitochondrial function [i.e., respiratory control index (RCI), see below].
H2O2 production in permeabilized fibers.
H2O2 production was measured with Amplex Red reagent, which reacts with H2O2 in a 1:1 stoichiometry catalyzed by HRP to yield the fluorescent compound resorufin and molar equivalent O2. Resorufin has excitation/emission characteristics of 563 nm/587 nm and is extremely stable once formed. Fluorescence was measured continuously [change in fluorescence (
F)/min] with a Spex Fluoromax 3 (Jobin Yvon) spectrofluorometer with temperature control and magnetic stirring. After baseline F (reactants only) was established, the reaction was initiated by addition of a permeabilized fiber bundle to 300 µl of buffer Z containing 5 µM Amplex Red and 0.5 U/ml HRP, with 5 mM pyruvate and 2 mM malate for complex I substrate or 3 mM succinate for complex II substrate at 37°C. For the succinate experiments, the fiber bundle was washed briefly in buffer Z without substrate to eliminate residual pyruvate and malate from the wash. Where stated, 10 µg/ml oligomycin was included in the reaction buffer to block ATP synthase and ensure state 4 respiration. To investigate the topology of O2–· production, duplicate experiments were conducted with and without Cu,Zn-SOD (40 U/ml) added to the reaction buffer. Inclusion of SOD ensures that O2–· released directly from the cytoplasmic face of the mitochondrial inner membrane is converted to H2O2, whereas in the absence of exogenous SOD, detection of O2–· is thought to be limited to that originating from the mitochondrial matrix after conversion to H2O2 by endogenous matrix SOD (41). H2O2 production rate was calculated from the slope of F/min, after subtracting background, from a standard curve established each day with the appropriate reaction conditions. At the conclusion of each experiment, fiber bundles were dried overnight on glass at 100°C. H2O2 production is expressed as picomoles per minute per milligram of dry weight.
Mitochondrial respiration in permeabilized fibers.
Respiration was measured at 37°C with a modified Clark-type O2 electrode (Hansatech Instruments). After a baseline rate of O2 consumption by the electrode itself was established, the reaction was initiated by addition of a permeabilized fiber bundle to 1 ml of buffer Z containing 20 mM creatine to saturate creatine kinase (35, 51) with either 5 mM pyruvate and 2 mM malate for complex I substrate or 3 mM succinate for complex II substrate. The rate of O2 consumption was measured in 2-min increments and expressed as nanomoles of O2 consumed per minute per milligram of dry weight. Basal respiration (state 4; 
O2) was determined in the presence of 10 µg/ml oligomycin to inhibit ATP synthesis. Maximal respiration (state 3; O2 max) was determined in parallel fiber bundles in the presence of 330 µM ADP. The RCI was calculated as O2 max/O2 (35). RCI values averaged 4.0 for all three muscle types (Table 1), indicating a similar high degree of coupling of respiration to phosphorylation. The percent free radical leak was calculated as the rate of H2O2 production divided by two times the rate of O2 consumption and multiplied by 100 (36).
|
Mitochondrial glutathione peroxidase and citrate synthase activity.
After the H2O2 scavenging experiments, permeabilized fiber bundles were placed in 300 µl of standard TE buffer (in mM: 10 Tris and 1 EDTA) + 1% Tween 20 and protease inhibitor cocktail and allowed to chill on ice for 30 min. The solutions containing the fiber bundles were then freeze-thawed in liquid N2 three times to fully fracture and liberate all remaining membrane-bound compartments. After brief centrifugation, the supernatants were collected and assayed for glutathione peroxidase (Northwest Life Science Specialties, Vancouver, WA) and citrate synthase (39) activity. The fiber bundles remaining were rinsed in H2O, dried, and weighed as described above.
Statistical analysis. Statistical analyses were performed using two-way ANOVA, with all pairwise multiple comparisons among conditions performed using the Student-Newman-Keuls method. The level of significance was set at P < 0.05.
|
RESULTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
100 µg/ml) reduced maximal ADP-stimulated respiration, likely reflecting loss of nucleotide sensitivity at the outer mitochondrial membrane. Detection of basal H2O2 production during state 4 respiration supported by pyruvate-malate also reached a maximum at 50 µg/ml saponin (Fig. 1B), confirming full permeabilization at this concentration. A low level of H2O2 production was observed in deenergized (no substrate) nonpermeabilized fiber bundles, presumably originating from metabolism of endogenous intracellular substrates.
|
Mitochondrial H2O2 production during pyruvate-malate-supported respiration.
Shown in Fig. 2A are representative traces from experiments with pyruvate-malate-supported state 4 respiration from permeabilized soleus, RG, and WG fiber bundles. The average basal rate of mitochondrial H2O2 production (addition of fiber bundle, no inhibitors present) was highest (P < 0.05) in WG, followed by RG and then soleus muscle (Fig. 2B). In isolated mitochondria, the rate of mitochondrial O2–· formation is directly governed by both membrane potential (
) and pH gradient across the inner membrane (5) and thus is favored only under state 4 conditions. In the present study, addition of ADP or FCCP (uncoupler; data not shown) completely eliminated the basal level H2O2 production, consistent with the fall in that occurs with transition to state 3 respiration or mild uncoupling and confirming that mitochondria are the source of H2O2 in permeabilized fibers (Fig. 2B).
|
50–75% in all three muscle types, confirming that complex III is an extremely active site of O2–· generation in permeabilized fibers when respiration is inhibited by antimycin A. In separate experiments, addition of rotenone to permeabilized fibers respiring on pyruvate-malate also induced H2O2 production, confirming complex I as a site of O2–· formation. Interestingly, the level of H2O2 production remaining after addition of stigmatellin was similar in all three muscles to the rate generated at complex I in the presence of rotenone, suggesting that inhibition of electron entry at complex III slows electron flow at complex I, elevating the redox state (i.e., similar to rotenone) and accelerating O2–· formation at complex I. Mitochondrial H2O2 production during succinate-supported respiration. Respiration supported by the complex II substrate succinate generated extremely high rates of H2O2 production under basal state 4 conditions (no inhibitors; Fig. 3). Transition to state 3 respiration by addition of ADP completely eliminated H2O2 production (Fig. 3, A, inset, and B), again confirming mitochondria as the source of H2O2. Both under basal conditions (Fig. 3B) and in the presence of inhibitors (Fig. 3C), mitochondrial H2O2 production was highest in both RG and WG muscle, whereas soleus muscle displayed the lowest overall responses. In all three muscle types, addition of rotenone reduced H2O2 production by >70% (Fig. 3, A, inset, and C), indicating that complex I is the major site of O2–· generation during respiration supported exclusively by succinate because of reverse electron flow from complex II to complex I. As anticipated, subsequent addition of antimycin A in fibers supported by succinate/rotenone elicited a marked increase in O2–· production at complex III that, in turn, was blocked by addition of stigmatellin.
|
Mitochondrial free radical leak. Depicted in Fig. 4 is the fraction of molecules of O2 consumed that give rise to H2O2 released by mitochondria (free radical leak) during either pyruvate-malate- or succinate-supported state 4 respiration in each of the three muscle types. Free radical leak was significantly (P < 0.05) higher in WG than in either RG or soleus muscle, providing evidence that type IIB myofibers possess an inherently greater propensity to generate/release O2–·-derived free radicals under state 4 conditions.
|
1.5- and 3.0-fold greater (P < 0.05) than rates obtained from RG and WG fibers, respectively (Fig. 5B, left). The H2O2 scavenging rate was also 1.5-fold greater (P < 0.05) in RG than in WG fibers. Even after normalization for mitochondrial content, overall H2O2 scavenging rate (nmol·min–1·citrate synthase activity–1) was still significantly greater in soleus than in RG and WG muscle, whereas rates for RG and WG muscle were similar (Fig. 5B, right). Activity of glutathione peroxidase, the primary enzyme responsible for H2O2 detoxification in mitochondria (2, 54), was approximately twofold higher (P < 0.05) in soleus and RG (7.02 ± 0.44 and 6.85 ± 0.68 mU·ml–1·mg dry wt–1, respectively) than in WG (3.39 ± 0.37 mU·ml–1·mg dry wt–1) muscle when expressed in absolute terms; however, once normalized for mitochondrial content, glutathione peroxidase activity was similar among the three muscle types (not shown).
|
|
DISCUSSION |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
It is generally thought that basal respiration (state 4) supported by substrates feeding exclusively to complex I in the absence of respiratory chain inhibitors elicits little to no H2O2 production (41). However, the recent development of a highly sensitive fluorescent H2O2 indicator (Amplex Red) has enabled characterization of ROS production in isolated mitochondria respiring on NADH-linked substrates at levels previously considered insignificant (40). In the present study, low but consistent rates of H2O2 production were observed in all three types of permeabilized fiber bundles during state 4 respiration supported by the complex I substrate combination of pyruvate plus malate (Fig. 2). This basal level of H2O2 production occurred only in the presence of substrate (i.e., no H2O2 production observed in deenergized fiber bundles; Fig. 2B, inset) and was completely eliminated by addition of ADP (lowers by increasing H+ flow through ATP synthase; Fig. 2B) or FCCP (protonophore that lowers by increasing H+ conductance; data not shown), confirming mitochondria as the source of H2O2 in respiring permeabilized fiber bundles. Surprisingly, basal rates of H2O2 production were considerably different among the three types of muscles (WG > RG > soleus; Fig. 2B). H2O2 production in isolated mitochondria from skeletal muscle, heart, and brain is particularly sensitive to , increasing exponentially at the upper range of values (i.e., 170–185 mV; Refs. 17, 22, 40, 50). Whether differences in basal rates of H2O2 production among different skeletal muscle fiber types reflect corresponding differences in requires further study. However, the percentage of O2 giving rise to H2O2 release during complex I-supported state 4 respiration was 3.5-fold higher in WG than in either RG or soleus muscle (Fig. 4), consistent with the notion that mitochondrial function (i.e., inherent proton leak, respiratory control, electron transport chain stoichiometry) differs among mitochondria found within the three major types of skeletal muscle (20, 23).
In the absence of inhibitors, succinate-supported state 4 respiration generated extremely high basal rates of mitochondrial H2O2 production in permeabilized fiber bundles from all three types of skeletal muscle (Fig. 3). Succinate donates its electrons directly via complex II (succinate dehydrogenase) of the respiratory chain. The high rate of H2O2 production from this source was virtually eliminated by addition of the complex I inhibitor rotenone in permeabilized soleus and RG muscle and partially inhibited in permeabilized WG muscle. These findings largely confirm those of previous studies on isolated mitochondria from rat brain, heart, and skeletal muscle (14, 16, 17, 26, 43, 48, 50), demonstrating that complex I is the major site of O2–· generation during succinate-supported respiration due to reverse electron flow from complex II to complex I. The remaining H2O2 production evident in WG fibers likely stems from O2–· generation at complex III (see below).
Addition of antimycin A, a complex III inhibitor, to permeabilized fibers supported by either pyruvate-malate or succinate/rotenone (blocking reverse electron flow) also generated extremely high rates of H2O2 production (Figs. 2 and 3). Complex III catalyzes the oxidation of reduced coenzyme Q (QH2, quinol) by using a bifurcated series of single electron transfers, with cytochrome c serving as the final electron acceptor (31). Antimycin A, which blocks electron transfer from the cytochrome b proteins to the Qi site ("inside" or matrix side of inner membrane) of complex III, dramatically increases O2–· production during respiration supported by pyruvate-malate or succinate/rotenone (Figs. 2 and 3), presumably due to accumulation of an unstable semiquinone at the Qo site ("outside" or cytosolic side of inner membrane) (47). Subsequent addition of stigmatellin, which prevents the transfer of the first electron from QH2 to the Rieske protein, completely abrogated (succinate/rotenone) or significantly reduced (pyruvate-malate) H2O2 production induced by antimycin A, confirming complex III as the site of O2–· generation. O2–· production at complex III has recently been implicated in the cellular response to hypoxia, providing some hint of the potential physiological relevance of O2–· production at this site (13).
Of considerable relevance to the potential mechanisms and consequences of ROS production is the topology or sidedness of O2–· production, a matter that has been somewhat controversial with respect to complex III. Initial studies on submitochondrial particles indicated that O2–· is released exclusively to the matrix (4, 48), whereas more recent studies using isolated mitochondria (±SOD) have provided evidence that O2–· is released almost exclusively to the intermembrane space (15, 29, 41). Using both indirect (H2O2) and direct measures of O2–· production in mitochondria isolated from mouse whole hindlimb muscles (containing a mixed population of myofiber types), Muller et al. (30) found that the Qo site of complex III releases O2–· in approximately equal portions to both the cytoplasmic and matrix sides of the inner membrane, offering two potential mechanisms by which this may occur. The data presented in Figs. 2 and 3 of the present study, however, provide evidence that the topology of O2–· release from complex III in skeletal muscle may be fiber type specific. In permeabilized fiber bundles from soleus and RG muscle, antimycin A-stimulated H2O2 production was similar in the presence or absence of SOD, whereas in WG permeabilized fibers, H2O2 detection was reduced >65% when SOD was not included in the reaction buffer. The simplest interpretation of these data is that virtually all of the O2–· produced at complex III in types I and IIA fibers is released to the mitochondrial matrix, whereas in type IIB fibers, the majority is released to the intermembrane space. An alternative possibility is that mitochondria present in oxidative fibers may possess the means of rapidly dismutating O2–· released directly into the intermembrane space, which would explain the apparent insensitivity to exogenous SOD in RG and soleus fibers observed in the present study. Indeed, the recent confirmation of Cu,Zn-SOD in the intermembrane space of mitochondria from rat liver and yeast (33, 42) lends credence to this possibility. However, complex III-mediated H2O2 production is similar in skeletal muscle mitochondria isolated from mice lacking Cu,Zn-SOD and wild-type mice (30), implying that Cu,Zn-SOD may not be localized to the intermembrane space of mitochondria in skeletal muscle or that activity of the enzyme is lost during the isolation of mitochondria but retained in permeabilized fibers. Further work is required to determine the fiber type-specific topology of O2–· release by complex III in skeletal muscle.
Surprisingly, ROS production at complex I also appeared to be partially sensitive to SOD, because rotenone-stimulated H2O2 production decreased by 25% in RG muscle and 50% in WG muscle during pyruvate-malate-supported respiration when SOD was not included in the reaction buffer (Fig. 2). These findings suggest that at least a fraction of complex I-generated O2–· may be released to the intermembrane space, an interpretation that is at odds with the widely held notion that complex I releases O2–· exclusively to the matrix (30, 41). Complex I consists of 46 distinct subunits, the majority of which have been identified but remain uncharacterized in terms of function (8, 18). Whether mitochondria preserved in their native reticulum configuration in permeabilized fibers retain a O2–·-producing site in complex I that is otherwise lost or masked in preparations of isolated mitochondria requires further study.
The fact that mitochondrial H2O2 production rates observed among the three types of muscles did not mirror differences in respiratory capacity/mitochondrial content suggests that mitochondria housed within each myofiber type possess intrinsic features that affect ROS production and/or ROS removal. Measurements of H2O2 scavenging rate in permeabilized fibers indicated that ROS removal capacity is significantly greater in soleus > RG > WG muscle when expressed relative to milligrams of dry weight (Fig. 5B, left). When normalized to citrate synthase activity (index of mitochondrial content), H2O2 scavenging rate was still highest in soleus muscle, whereas rates were similar in RG and WG muscle (Fig. 5B, right). H2O2 detoxification in mitochondria is largely a function of glutathione peroxidase activity (2, 54). Although higher in soleus and RG muscle when expressed in absolute terms, glutathione peroxidase activity was similar in all three types of muscle when normalized for mitochondrial content. Thus it appears that soleus myofibers may possess a slightly greater capacity to scavenge mitochondrial H2O2, limiting overall ROS emission. By contrast, the similar mitochondrion-specific scavenging capacity evident in RG and WG fibers suggests that the higher state 4 rate of free radical leak observed in WG muscle (Fig. 4) may be due to an elevated sensitivity to conditions that favor mitochondrial ROS production in type IIB fibers, perhaps due to differences in the stoichiometry-activity ratios of the respiratory complexes (25), susceptibility to proton pump "slip" at complex IV (21), or other mechanisms. Alternatively, the much thinner ultrastructure of the mitochondrial reticulum in type IIB fibers (32) may limit the efficiency of intramitochondrial scavenging and thus contribute to an overall greater rate of ROS emission.
In summary, with a novel in situ approach to examine the control of mitochondrial ROS production in skeletal muscle, the findings of the present study suggest that both basal and maximal rates of mitochondrial H2O2 production, as well as the topology of O2–· release/dismutation, differ markedly among the three major skeletal muscle fiber types. Type IIB fibers, which possess fewer mitochondria and rely heavily on glycolytic metabolism, appear to possess unique properties that potentiate mitochondrial ROS production/emission. Considered in the context of the mitochondrial theory of aging (10), these findings are consistent with the hypothesis that the mitochondrial dysfunction, atrophy, and selective loss of type II myofibers that occur with aging (28, 45, 52) may stem from an inherent susceptibility of type II (particularly type IIB) myofibers to conditions that favor ROS generation and oxidative stress over the course of a lifetime.
|
ACKNOWLEDGMENTS |
|---|
|
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 METHODSRESULTSDISCUSSIONREFERENCES |
|---|
2. Antunes F, Han D, and Cadenas E. Relative contributions of heart mitochondria glutathione peroxidase and catalase to H2O2 detoxification in in vivo conditions. Free Radic Biol Med 33: 1260–1267, 2002.[CrossRef][ISI][Medline]
3. Beal MF, Howell N, and Bodis-Wollner I. Mitochondria and Free Radicals in Neurodegenerative Disease. New York: Wiley-Liss, 1997.
4. Boveris A and Cadenas E. Mitochondrial production of superoxide anions and its relationship to the antimycin insensitive respiration. FEBS Lett 54: 311–314, 1975.[CrossRef][ISI][Medline]
5. Brand MD, Affourtit C, Esteves TC, Green K, Lambert AJ, Miwa S, Pakay JL, and Parker N. Mitochondrial superoxide: production, biological effects, and activation of uncoupling proteins. Free Radic Biol Med 37: 755–767, 2004.[CrossRef][ISI][Medline]
6. Brand MD, Buckingham JA, Esteves TC, Green K, Lambert AJ, Miwa S, Murphy MP, Pakay JL, Talbot DA, and Echtay KS. Mitochondrial superoxide and aging: uncoupling-protein activity and superoxide production. Biochem Soc Symp 71: 203–213, 2004.[Medline]
7. Bua EA, McKiernan SH, Wanagat J, McKenzie D, and Aiken JM. Mitochondrial abnormalities are more frequent in muscles undergoing sarcopenia. J Appl Physiol 92: 2617–2624, 2002.
8. Carroll J, Fearnley IM, Shannon RJ, Hirst J, and Walker JE. Analysis of the subunit composition of complex I from bovine heart mitochondria. Mol Cell Proteomics 2: 117–126, 2003.
9. Delp MD and Duan C. Composition and size of type I, IIA, IID/X, and IIB fibers and citrate synthase activity of rat muscle. J Appl Physiol 80: 261–270, 1996.
10. Dufour E and Larsson NG. Understanding aging: revealing order out of chaos. Biochim Biophys Acta 1658: 122–132, 2004.[Medline]
11. Evans JL, Goldfine ID, Maddux BA, and Grodsky GM. Oxidative stress and stress-activated signaling pathways: a unifying hypothesis of type 2 diabetes. Endocr Rev 23: 599–622, 2002.
12. Fiskum G. Mitochondrial participation in ischemic and traumatic neural cell death. J Neurotrauma 17: 843–855, 2000.[ISI][Medline]
13. Guzy RD, Hoyos B, Robin E, Chen H, Liu L, Mansfield KD, Simon MC, Hammerling U, and Schumacker PT. Mitochondrial complex III is required for hypoxia-induced ROS production and cellular oxygen sensing. Cell Metab 1: 401–408, 2005.[CrossRef][ISI][Medline]
14. Gyulkhandanyan AV and Pennefather PS. Shift in the localization of sites of hydrogen peroxide production in brain mitochondria by mitochondrial stress. J Neurochem 90: 405–421, 2004.[CrossRef][ISI]
15. Han D, Antunes F, Canali R, Rettori D, and Cadenas E. Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol. J Biol Chem 278: 5557–5563, 2003.
16. Han D, Canali R, Rettori D, and Kaplowitz N. Effect of glutathione depletion on sites and topology of superoxide and hydrogen peroxide production in mitochondria. Mol Pharmacol 64: 1136–1144, 2003.
17. Hansford RG, Hogue BA, and Mildaziene V. Dependence of H2O2 formation by rat heart mitochondria on substrate availability and donor age. J Bioenerg Biomembr 29: 89–95, 1997.[CrossRef][ISI][Medline]
18. Hirst J, Carroll J, Fearnley IM, Shannon RJ, and Walker JE. The nuclear encoded subunits of complex I from bovine heart mitochondria. Biochim Biophys Acta 1604: 135–150, 2003.[Medline]
19. Hood DA. Contractile activity-induced mitochondrial biogenesis in skeletal muscle. J Appl Physiol 90: 1137–1157, 2001.
20. Jackman MR and Willis WT. Characteristics of mitochondria isolated from type I and type IIb skeletal muscle. Am J Physiol Cell Physiol 270: C673–C678, 1996.
21. Kadenbach B. Intrinsic and extrinsic uncoupling of oxidative phosphorylation. Biochim Biophys Acta 1604: 77–94, 2003.[Medline]
22. Korshunov SS, Skulachev VP, and Starkov AA. High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria. FEBS Lett 416: 15–18, 1997.[CrossRef][ISI][Medline]
23. Kushmerick MJ, Meyer RA, and Brown TR. Regulation of oxygen consumption in fast- and slow-twitch muscle. Am J Physiol Cell Physiol 263: C598–C606, 1992.
24. Kuznetsov AV, Tiivel T, Sikk P, Kaambre T, Kay L, Daneshrad Z, Rossi A, Kadaja L, Peet N, Seppet E, and Saks VA. Striking differences between the kinetics of regulation of respiration by ADP in slow-twitch and fast-twitch muscles in vivo. Eur J Biochem 241: 909–915, 1996.[ISI][Medline]
25. Kwong LK and Sohal RS. Substrate and site specificity of hydrogen peroxide generation in mouse mitochondria. Arch Biochem Biophys 350: 118–126, 1998.[CrossRef][ISI][Medline]
26. Liu Y, Fiskum G, and Schubert D. Generation of reactive oxygen species by the mitochondrial electron transport chain. J Neurochem 80: 780–787, 2002.[CrossRef][ISI][Medline]
27. Lopez ME, Van Zeeland NL, Dahl DB, Weindruch R, and Aiken JM. Cellular phenotypes of age-associated skeletal muscle mitochondrial abnormalities in rhesus monkeys. Mutat Res 452: 123–138, 2000.[ISI][Medline]
28. Lopez ME, Zainal TA, Chung SS, Aiken JM, and Weindruch R. Oxidative stress and the pathogenesis of sarcopenia. In: Exercise and Oxygen Toxicity: A Handbook, edited by Sen CK, Packer L, and Haenninen O. New York: Elsevier Science, 1999, p. 831–880.
29. Miwa S, St-Pierre J, Partridge L, and Brand MD. Superoxide and hydrogen peroxide production by Drosophila mitochondria. Free Radic Biol Med 35: 938–948, 2003.[CrossRef][ISI][Medline]
30. Muller FL, Liu Y, and Van Remmen H. Complex III releases superoxide to both sides of the inner mitochondrial membrane. J Biol Chem 279: 49064–49073, 2004.
31. Nicholls DG and Ferguson SJ. Bioenergetics 3. London: Academic, 2002.
32. Ogata T and Yamasaki Y. Ultra-high-resolution scanning electron microscopy of mitochondria and sarcoplasmic reticulum arrangement in human red, white, and intermediate muscle fibers. Anat Rec 248: 214–223, 1997.[CrossRef][Medline]
33. Okado-Matsumoto A and Fridovich I. Subcellular distribution of superoxide dismutases (SOD) in rat liver. Cu,Zn-SOD in mitochondria. J Biol Chem 276: 38388–38393, 2001.
34. Ran Q, Liang H, Gu M, Qi W, Walter CA, Roberts LJ II, Herman B, Richardson A, and Van Remmen H. Transgenic mice overexpressing glutathione peroxidase 4 are protected against oxidative stress-induced apoptosis. J Biol Chem 279: 55137–55146, 2004.
35. Saks VA, Veksler VI, Kuznetsov AV, Kay L, Sikk P, Tiivel T, Tranqui L, Olivares J, Winkler K, Wiedemann F, and Kunz WS. Permeabilized cell and skinned fiber techniques in studies of mitochondrial function in vivo. Mol Cell Biochem 184: 81–100, 1998.[CrossRef][ISI][Medline]
36. Sanz A, Caro P, Ibanez J, Gomez J, Gredilla R, and Barja G. Dietary restriction at old age lowers mitochondrial oxygen radical production and leak at complex I and oxidative DNA damage in rat brain. J Bioenerg Biomembr 37: 83–90, 2005.[CrossRef][ISI][Medline]
37. Schriner SE, Linford NJ, Martin GM, Treuting P, Ogburn CE, Emond M, Coskun PE, Ladiges W, Wolf N, Van Remmen H, Wallace DC, and Rabinovitch PS. Extension of murine lifespan by overexpression of catalase targeted to mitochondria. Science 308: 1901–1911, 2005.
38. Seppet EK, Kaambre T, Sikk P, Tiivel T, Vija H, Tonkonogi M, Sahlin K, Kay L, Appaix F, Braun U, Eimre M, and Saks VA. Functional complexes of mitochondria with Ca,MgATPases of myofibrils and sarcoplasmic reticulum in muscle cells. Biochim Biophys Acta 1504: 379–395, 2001.[Medline]
39. Srere PA. Citrate synthase. Methods Enzymol 13: 3–5, 1969.
40. Starkov AA and Fiskum G. Regulation of brain mitochondrial H2O2 production by membrane potential and NAD(P)H redox state. J Neurochem 86: 1101–1107, 2003.[ISI][Medline]
41. St-Pierre J, Buckingham JA, Roebuck SJ, and Brand MD. Topology of superoxide production from different sites in the mitochondrial electron transport chain. J Biol Chem 277: 44784–44790, 2002.
42. Sturtz LA, Diekert K, Jensen LT, Lill R, and Culotta VC. A fraction of yeast Cu,Zn-superoxide dismutase and its metallochaperone, CCS, localize to the intermembrane space of mitochondria. A physiological role for SOD1 in guarding against mitochondrial oxidative damage. J Biol Chem 276: 38084–38089, 2001.
43. Talbot DA, Lambert AJ, and Brand MD. Production of endogenous matrix superoxide from mitochondrial complex I leads to activation of uncoupling protein 3. FEBS Lett 556: 111–115, 2004.[CrossRef][ISI][Medline]
44. Tonkonogi M, Fernstrom M, Walsh B, Ji LL, Rooyackers O, Hammarqvist F, Wernerman J, and Sahlin K. Reduced oxidative power but unchanged antioxidative capacity in skeletal muscle from aged humans. Pflügers Arch 446: 261–269, 2003.[ISI][Medline]
45. Trappe S, Gallagher P, Harber M, Carrithers J, Fluckey J, and Trappe T. Single muscle fibre contractile properties in young and old men and women. J Physiol 552: 47–58, 2003.
46. Trifunovic A, Wredenberg A, Falkenberg M, Spelbrink JN, Rovio AT, Bruder CE, Bohlooly YM, Gidlof S, Oldfors A, Wibom R, Tornell J, Jacobs HT, and Larsson NG. Premature ageing in mice expressing defective mitochondrial DNA polymerase. Nature 429: 417–423, 2004.[CrossRef][Medline]
47. Turrens JF. Mitochondrial formation of reactive oxygen species. J Physiol 552: 335–344, 2003.
48. Turrens JF and Boveris A. Generation of superoxide anion by the NADH dehydrogenase of bovine heart mitochondria. Biochem J 191: 421–427, 1980.[ISI][Medline]
49. Vendelin M, Beraud N, Guerrero K, Andrienko T, Kuznetsov AV, Olivares J, Kay L, and Saks VA. Mitochondrial regular arrangement in muscle cells: a "crystal-like" pattern. Am J Physiol Cell Physiol 288: C757–C767, 2005.
50. Votyakova TV and Reynolds IJ. 
m-Dependent and -independent production of reactive oxygen species by rat brain mitochondria. J Neurochem 79: 266–277, 2001.[CrossRef][ISI][Medline]
51. Walsh B, Tonkonogi M, Soderlund K, Hultman E, Saks V, and Sahlin K. The role of phosphorylcreatine and creatine in the regulation of mitochondrial respiration in human skeletal muscle. J Physiol 537: 971–978, 2001.
52. Wanagat J, Cao Z, Pathare P, and Aiken JM. Mitochondrial DNA deletion mutations colocalize with segmental electron transport system abnormalities, muscle fiber atrophy, fiber splitting, and oxidative damage in sarcopenia. FASEB J 15: 322–332, 2001.
53. Yaffe MP. The machinery of mitochondrial inheritance and behavior. Science 283: 1493–1497, 1999.
54. Zoccarato F, Cavallini L, and Alexandre A. Respiration-dependent removal of exogenous H2O2 in brain mitochondria: inhibition by Ca2+. J Biol Chem 279: 4166–4174, 2004.
This article has been cited by other articles:
|
|
 |
|
  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]
|
|
|
|
 |
|
 V. Ljubicic and D. A. Hood Kinase-specific responsiveness to incremental contractile activity in skeletal muscle with low and high mitochondrial content Am J Physiol Endocrinol Metab, July 1, 2008; 295(1): E195 - E204. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. L. Clanton Hypoxia-induced reactive oxygen species formation in skeletal muscle J Appl Physiol, June 1, 2007; 102(6): 2379 - 2388. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Mogensen, K. Sahlin, M. Fernstrom, D. Glintborg, B. F. Vind, H. Beck-Nielsen, and K. Hojlund Mitochondrial Respiration Is Decreased in Skeletal Muscle of Patients With Type 2 Diabetes Diabetes, June 1, 2007; 56(6): 1592 - 1599. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. E. Conley, C. E. Amara, S. A. Jubrias, and D. J. Marcinek Muscle-energetic and cardio-pulmonary determinants of exercise tolerance in humans: Mitochondrial function, fibre types and ageing: new insights from human muscle in vivo Exp Physiol, March 1, 2007; 92(2): 333 - 339. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. Freyssenet Energy sensing and regulation of gene expression in skeletal muscle J Appl Physiol, February 1, 2007; 102(2): 529 - 540. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||
significantly (P < 0.001) different from +SOD condition.
), RG (
), WG (
), and no-fiber bundle control with 40 µM H2O2 (
). B: quantified H2O2 scavenging rates (means ± SE; n = 5 or 6) expressed both in absolute terms (left; pmol·min–1·mg dry wt–1) and relative to citrate synthase (CS) activity (right; pmol·min–1·CS activity–1). *Significantly (P < 0.001) different from soleus;