| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
MUSCLE CELL BIOLOGY AND CELL MOTILITY
1Department of Physiology and Pharmacology, Karolinska Institutet and 2Center for Surgical Sciences, Karolinska Institutet at Ersta Hospital, Stockholm, Sweden
Submitted 21 March 2007 ; accepted in final form 2 July 2007
 |
ABSTRACT |
|---|
 TOP ABSTRACT METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
50%, despite increased ROS production. This increase was not associated with a change in the amount of immunoreactive aconitase (Western blot) but was markedly inhibited by cyclosporin A, an inhibitor of the protein phosphatase calcineurin. Immunoprecipitation experiments demonstrated that aconitase was phosphorylated on serine residues. Aconitase in cell-free extracts was inactivated by the addition of the ROS hydrogen peroxide. In conclusion, the results suggest that aconitase activity can be regulated by at least two mechanisms: oxidation/reduction and phosphorylation/dephosphorylation. During contraction, a ROS-mediated inactivation of aconitase can be overcome, possibly by dephosphorylation of the enzyme. The dual-control system may be important in maintaining aerobic ATP production during muscle contraction. glutathione; reactive oxygen species
|
METHODS |
|---|
|
TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Cyclosporin A was from Sigma(C3662). A polyclonal antibody against mitochondrial aconitase was a kind gift from Dr. Luke Szweda (Oklahoma Medical Research Foundation). Glutathiones were analyzed with a commercial kit (Biooxytech GSH/GSSG-412, Oxis Health Products, Portland, OR). All other chemicals were from either Sigma or Boehringer Mannheim.
Adult male NMRI mice were housed at room temperature with a 12:12-h light-dark cycle. Food and water were provided ad libitum. The mice were purchased from B&K Universal (Sollentuna, Sweden). Animals were killed by rapid cervical dislocation, and the extensor digitorum longus (EDL, fast twitch) and soleus (slow twitch) muscles were isolated. All experimental procedures in the animal experiments were approved by the Stockholm North Ethics Committee.
Experimental Design
Human studies. Five healthy volunteers (4 men, 1 woman) participated in the study. Their age, weight, height and maximal workload capacity (Wmax) averaged 26 ± 1 yr, 70.0 ± 5.4 kg, 178 ± 5 cm, and 272 ± 33 W, respectively. Wmax was defined as the maximal workload that the subjects could maintain for 6 min (27). The nature of the study, its purpose, and possible risks were explained to the subjects before they provided written voluntary consent to participate in the study. The experimental protocol in the human study was approved by the Institutional Ethical Committee at the Karolinska Institute. The study conformed to the standards set by the Declaration of Helsinki.
Full details on the experimental protocol are provided elsewhere (27). Briefly, subjects were studied in the morning after an overnight fast. Biopsies were obtained from the lateral aspect of the quadriceps femoris muscle at rest and immediately after 30 min of cycling in the upright position at 75% of Wmax. Thereafter, the subjects rested for 1–2 min and then performed six 60-s bouts of cycling at 125% of Wmax, followed immediately by a third biopsy. The six bouts were separated by 1-min rest periods. Postexercise biopsies were quick frozen in liquid N2 within 10–15 s after termination of exercise. The submaximal and supramaximal cycling protocols were chosen to represent a large range of exercise intensities often performed by humans.
Animal studies.
For contraction studies, stainless steel hooks were tied with nylon thread to the tendons of the muscles. Muscles were then transferred to a stimulation chamber (volume 
10 ml) and mounted between a force transducer and an adjustable holder (World Precision Instruments). The chamber temperature was set at 30°C with a water-jacketed circulation bath. The muscle was bathed in a Tyrode solution with the following composition (in mM): 121 NaCl, 5 KCl, 1.8 CaCl2, 0.4 NaH2PO4, 0.5 MgCl2, 24 NaHCO3, 0.1 EDTA, and 5.5 glucose, with 0.1% fetal calf serum. The pH of the Tyrode solution was set to 7.4 by continuously gassing the solution with 95% O2-5% CO2. Muscles were stimulated with current pulses (0.5-ms duration; 150% of current required for maximum force response) via plate electrodes lying parallel to the fibers. Muscles were set to the length at which tetanic force was maximum (Lo) and then allowed to recover for 30 min. Thereafter, they were stimulated to perform repeated tetanic contractions at 70 Hz (tetanic duration 100 ms, 2 trains/s) for 10 min and frozen in liquid N2 within 10 s after termination of the last contraction. The stimulation protocol resulted in a marked loss of force (see RESULTS) but no irreversible damage as judged by a robust force recovery (28). Nonstimulated muscles from the contralateral leg served as controls and were incubated for 40 min in the same buffer at the same temperature. In some experiments, after Lo was set muscles from the same mouse were bathed in Tyrode solution containing 10 µM cyclosporin A, an inhibitor of the protein phosphatase calcineurin (13), or an equal volume of diluent (ethanol; final concentration 0.1%).
Analysis
Human muscles were divided into aliquots at –20°C. One aliquot was freeze dried, dissected free of nonmuscle constituents, powdered, thoroughly mixed, and used for analysis of glutathiones (see below). Mouse muscles were also freeze dried and treated likewise for analysis of glutathiones. Enzyme activities were analyzed on wet-frozen aliquots of human muscles and wet-frozen whole soleus and EDL muscles.
For enzyme activities, muscles were homogenized (50 µl/mg wet wt) with ground glass homogenizers in ice-cold buffer consisting of (in mM) 50 Tris, 5 citrate, 0.6 MnCl2, and 1 cysteine, with 0.05% (vol/vol) Triton X-100, pH 7.4 (14). The homogenate was centrifuged at 1,400 g (4°C) for 1 min. Aliquots of the supernatant were frozen for subsequent analyses of citrate synthase (CS) and 
-hydroxyacyl-CoA dehydrogenase (HAD) with standard spectrophotometric techniques (1, 3).
For analysis of aconitase, we used a spectrophotometric assay following the conversion of citrate to isocitrate coupled with isocitrate dehydrogenase at 340 nm (14, 16). Preliminary experiments revealed that activity decreased by 20% when supernatants were stored on ice for 2 h. Freezing the supernatants at –20°C for 1 wk resulted in a complete loss of activity, and freeze-drying of frozen muscles resulted in a loss of 80% of activity. Therefore, freshly prepared supernatant (see above) was assayed within 10 min after homogenization. Activities were typically measured between 10 and 50 min, since occasionally a burst of activity was observed during the first 10 min of the reaction. This phenomenon was previously observed and attributed to a rapid conversion of citrate to isocitrate and subsequent formation of NADPH (14).
All enzyme activities were assayed at room temperature (22°C) under conditions that yielded linearity with respect to extract volume and time (data not shown). Protein content was measured in the supernatant with the Bio-Rad assay (Bio-Rad). For glutathiones, oxidized glutathione (GSSG) and the sum of reduced glutathione (GSH) and GSSG were assayed spectrophotometrically after the reduction of 5,5'-dithiobis(2-nitrobenzoic acid) at 412 nm as described elsewhere (29). In both assays GSSG was used as a standard, and the data are given in GSSG equivalents and should be multiplied by 2 to express as GSH equivalents.
Western blots were performed on the enzyme extracts (see above) for aconitase. Briefly, 15 µg of supernatant protein was separated by SDS-PAGE (4–12% Bis-Tris gels, Invitrogen) and transferred onto polyvinylidene difluoride (PVDF) membranes. Membranes were blocked in 5% (wt/vol) nonfat milk in Tris-buffered saline containing 0.05% Tween 20, followed by incubation with primary antibody made up in 5% (wt/vol) bovine serum albumin (1:2,000 dilution) overnight at 4°C. Membranes were then washed and incubated for 1 h at room temperature with secondary antibody (donkey anti-rabbit at 1:2,000 dilution). Immunoreactive bands were visualized with enhanced chemiluminescence (Super Signal, Pierce). Band densities were analyzed with Image J (National Institutes of Health; http//rsb.info.nih.gov/j/).
For immunoprecipitation of aconitase, EDL muscles were isolated and stimulated as above (one basal and the contralateral muscle stimulated). To obtain sufficient protein, two basal (or two stimulated) muscles from two mice were pooled. Muscles were homogenized in ice-cold buffer consisting of (in mM) 25 KH2PO4, 5 EDTA, and 50 KF, with 0.05% (vol/vol) Triton X-100, pH 7.25. Homogenates were centrifuged at 1,400 g (4°C) for 1 min. Supernatants were assayed for protein and used for immunoprecipitation. Equal amounts of protein (800 µg) were incubated with 10 µl of rabbit anti-aconitase antibody overnight at 4°C, with rotation, followed by addition of 50 µl of protein G agarose suspension (Santa Cruz Biotechnology) and an additional 4-h incubation at 4°C with rotation. Samples were washed three times with a buffer consisting of (in mM) 20 HEPES, 150 NaCl, 5 EDTA, 1 Na3VO4, and 25 KF, with 20% (vol/vol) glycerol and protease inhibitor cocktail, pH 7.6, eluted with 40 µl of LDS sample buffer, and heated for 10 min at 70°C. Samples were then loaded onto NuPAGE Novex 4–12% Bis-Tris gel (Invitrogen), separated by electrophoresis, and transferred onto a PVDF membrane (Bio-Rad). Membranes were blocked in 5% (wt/vol) nonfat milk in Tris-buffered saline containing 0.05% Tween 20, followed by incubation with primary antibody (anti-phosphoserine, 1.5 µl/ml, Zymed Laboratories). Membranes were then incubated with horseradish peroxidase (HRP)-conjugated antibody (HRP-conjugated anti-rabbit Ig, 1:20,000 dilution, Amersham), and immunoreactive bands were visualized with enhanced chemiluminescence (Super Signal).
Force was sampled online and stored on a desktop computer for subsequent analysis. Tetanic force was measured as the peak force during the 100 ms of stimulation.
Statistics
Significant differences between means were determined with a one-way repeated-measures ANOVA followed by the Holm-Sidak method (human study). Student's t-test for paired samples was used for the isolated mouse muscle studies. P < 0.05 was regarded as significant. Values are presented as means ± SE.
|
RESULTS |
|---|
|
TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Subjects performed submaximal and supramaximal exercise that resulted in increased ROS production, as judged by increases in GSSG (Table 1). Despite the increased ROS production, aconitase activity did not decrease. The activities of two other mitochondrial enzymes were also measured, one representative for the TCA cycle (CS) and one representative for mitochondrial -oxidation (HAD). There were no significant changes in the activities of these enzymes after exercise. Human quadriceps femoris muscle consists of various fiber types, and the heterogeneity could mask changes that occur in a given fiber type. Consequently, additional experiments were performed on isolated mouse muscles of defined fiber composition.
|
Muscles were stimulated with a protocol that leads to oxidative stress as judged by altered glutathione status and increased fluorescence of the intracellular fluorescent indicator 5(and 6)-chloromethyl-2',7'-dichlorodihydrofluorescein (29, 30). Under the stimulation conditions of the present study, GSSG content in the EDL increases twofold (from 1.1 ± 0.1 to 2.2 ± 0.1 µmol/g dry wt) (30). Surprisingly, aconitase activity increased almost 50% in the fast-twitch EDL muscle after repeated contractions, whereas it remained unchanged in the slow-twitch soleus muscle (Fig. 1A). This protocol results in marked fatigue in both muscles, albeit the soleus is more fatigue resistant than the EDL (Fig. 1B). Neither of the other measured mitochondrial enzyme activities was altered with stimulation (Table 2).
|
|
The finding that aconitase activity increased in EDL muscle after repeated contraction in the presence of oxidative stress raised the possibility that the enzyme was also regulated by a non-ROS-dependent mechanism. Indeed, it has been demonstrated that aconitase undergoes phosphorylation/dephosphorylation and this results in altered activity (9, 12, 25, 34). Therefore, an additional series of experiments was performed with cyclosporin A, an inhibitor of the protein phosphatase calcineurin. Cyclosporin A had no significant effect on the activity of aconitase in resting EDL muscles but significantly decreased the activity after repeated contractions (Fig. 2A). Cyclosporin A did not affect force production (initial force = 303 ± 29 mN for control and 301 ± 19 mN for cyclosporin A) or fatigue development during repeated contractions (Fig. 2B).
|
|
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
In addition to control by oxidation/reduction, recent studies indicate that aconitase can also be controlled by phosphorylation/dephosphorylation (9, 12, 25). The present findings demonstrate that aconitase in skeletal muscle is also phosphorylated, specifically on serine residues. However, it is not known whether phosphorylation of these sites alters the activity of the mitochondrial isoform of aconitase, nor is it known whether other phosphorylation sites (e.g., threonine or tyrosine) exist. It is well documented that protein kinase C can phosphorylate and inactivate cytosolic aconitase (12, 25). Moreover, aconitase from isolated potato tuber mitochondria can be phosphorylated as well, although the kinase responsible has not been identified (9). To our knowledge, there is no published information on the identity of the enzymes that regulate the phosphorylation state of mitochondrial aconitase in the basal state.
Aconitase can be activated by the action of the Ca2+-dependent protein phosphatase calcineurin (34). The present cyclosporin A experiments support the idea that calcineurin-mediated activation of aconitase is of physiological significance in fast-twitch muscle during repeated contractions. For this mechanism to exist, it is requisite that Ca2+-dependent calcineurin is expressed in mitochondria and that Ca2+ increases in mitochondria during contraction. Indeed, calcineurin is expressed in mitochondria (31), and Ca2+ increases in skeletal muscle mitochondria after repeated contractions (5). Cyclosporin A is also known to inhibit the mitochondrial permeability transition pore (11) and can thereby result in decreased calcineurin activity by altering mitochondrial Ca2+ handling.
In human skeletal muscle, the maximal activity of 2-oxoglutarate dehydrogenase (converts 2-oxoglutarate to succinyl-CoA) is considered to be rate limiting for flux through the TCA cycle (4). It follows that this enzyme would limit oxygen consumption and aerobic ATP consumption. The maximal activity of aconitase in human muscle (measured by conversion of citrate to isocitrate and not cis-aconitate to isocitrate) is similar to, or even, lower than that of 2-oxoglutarate dehydrogenase (cf. Refs. 4, 15; see also Table 1). Thus ROS-mediated inactivation of aconitase could also compromise aerobic ATP production during exercise (2). In line with this idea, it was recently demonstrated that after ischemia and reperfusion of rat hearts, which results in oxidative stress, both aconitase and 2-oxoglutarate dehydrogenase were inactivated and state 3 respiration (ADP-stimulated oxygen consumption) of cardiac mitochondria was decreased to 30% of control (26). Similarly, addition of H2O2 to cardiomyocytes in culture resulted in marked inactivation of aconitase (but not 2-oxoglutarate dehydrogenase) and impaired acetate oxidation (18). In contrast, in exercising humans oxygen consumption is not decreased during sustained exercise, even at the point of fatigue (19, 33). This latter observation is consistent with the findings of the present study, in which aconitase activity was maintained or even increased during exercise conditions that result in increased ROS production. We suggest that the dephosphorylation of aconitase serves as a mechanism that contributes to activation of the enzyme in the face of ROS production during exercise/contractions. This suggestion is based on the results from the isolated mouse muscle preparations. However, extrapolation of findings from isolated mouse muscle preparations to intact humans should be performed with caution, since variables in the latter (e.g., presence of blood-borne hormones, cytokines) that are not present in the isolated preparation could influence the results.
Recently, another mode of mitochondrial aconitase protection in the face of oxidative stress was described. It was demonstrated that the mitochondrial iron-binding protein frataxin prevents oxidant-induced inactivation of aconitase (7, 8). Whether frataxin also protects aconitase from inactivation by exercise-induced ROS remains to be investigated.
In conclusion, our data suggest that aconitase can be regulated by at least two mechanisms: oxidation/reduction and phosphorylation/dephosphorylation. Thus during contraction a ROS-mediated inactivation of aconitase can be overcome, possibly by dephosphorylation of the enzyme. This dual-control system is probably of physiological significance, since ROS-mediated inactivation of aconitase could result in compromised aerobic ATP production during muscle contraction.
|
GRANTS |
|---|
|
TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
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 |
|---|
|
TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
2. Andersson U, Leighton B, Young ME, Blomstrand E, Newsholme EA. Inactivation of aconitase and oxoglutarate dehydrogenase in skeletal muscle in vitro by superoxide anions and/or nitric oxide. Biochem Biophys Res Commun 249: 512–516, 1998.[CrossRef][ISI][Medline]
3. Bass A, Brdiczka D, Eyer P, Hofer S, Pette D. Metabolic differentiation of distinct muscle types at the level of enzymatic organization. Eur J Biochem 10: 198–206, 1969.[ISI][Medline]
4. Blomstrand E, Radegran G, Saltin B. Maximum rate of oxygen uptake by human skeletal muscle in relation to maximal activities of enzymes in the Krebs cycle. J Physiol 501: 455–460, 1997.
5. Bruton J, Tavi P, Aydin J, Westerblad H, Lännergren J. Mitochondrial and myoplasmic [Ca2+] in single fibres from mouse limb muscles during repeated tetanic contractions. J Physiol 551: 179–190, 2003.
6. Bulteau AL, Ikeda-Saito M, Szweda LI. Redox-dependent modulation of aconitase activity in intact mitochondria. Biochemistry 42: 14846–14855, 2003.[CrossRef][Medline]
7. Bulteau AL, Lundberg KC, Ikeda-Saito M, Isaya G, Szweda LI. Reversible redox-dependent modulation of mitochondrial aconitase and proteolytic activity during in vivo cardiac ischemia/reperfusion. Proc Natl Acad Sci USA 102: 5987–5991, 2005.
8. Bulteau AL, O'Neill HA, Kennedy MC, Ikeda-Saito M, Isaya G, Szweda LI. Frataxin acts as an iron chaperone protein to modulate mitochondrial aconitase activity. Science 305: 242–245, 2004.
9. Bykova NV, Egsgaard H, Moller IM. Identification of 14 new phosphoproteins involved in important plant mitochondrial processes. FEBS Lett 540: 141–146, 2003.[CrossRef][ISI][Medline]
10. Castro L, Rodriguez M, Radi R. Aconitase is readily inactivated by peroxynitrite, but not by its precursor, nitric oxide. J Biol Chem 269: 29409–29415, 1994.
11. Duchen MR. Roles of mitochondria in health and disease. Diabetes 53, Suppl 1: S96–S102, 2004.
12. Eisenstein RS, Tuazon PT, Schalinske KL, Anderson SA, Traugh JA. Iron-responsive element-binding protein. Phosphorylation by protein kinase C. J Biol Chem 268: 27363–27370, 1993.
13. Garcia-Roves PM, Jones TE, Otani K, Han DH, Holloszy JO. Calcineurin does not mediate exercise-induced increase in muscle GLUT4. Diabetes 54: 624–628, 2005.
14. Gardner PR. Aconitase: sensitive target and measure of superoxide. Methods Enzymol 349: 9–23, 2002.[ISI][Medline]
15. Haller RG, Henriksson KG, Jorfeldt L, Hultman E, Wibom R, Sahlin K, Areskog NH, Gunder M, Ayyad K, Blomqvist CG. Deficiency of skeletal muscle succinate dehydrogenase and aconitase. Pathophysiology of exercise in a novel human muscle oxidative defect. J Clin Invest 88: 1197–1206, 1991.[ISI][Medline]
16. Hausladen A, Fridovich I. Measuring nitric oxide and superoxide: rate constants for aconitase reactivity. Methods Enzymol 269: 37–41, 1996.[ISI][Medline]
17. Hollander J, Fiebig R, Gore M, Bejma J, Ookawara T, Ohno H, Ji LL. Superoxide dismutase gene expression in skeletal muscle: fiber-specific adaptation to endurance training. Am J Physiol Regul Integr Comp Physiol 277: R856–R862, 1999.
18. Janero DR, Hreniuk D. Suppression of TCA cycle activity in the cardiac muscle cell by hydroperoxide-induced oxidant stress. Am J Physiol Cell Physiol 270: C1735–C1742, 1996.
19. Katz A, Broberg S, Sahlin K, Wahren J. Leg glucose uptake during maximal dynamic exercise in humans. Am J Physiol Endocrinol Metab 251: E65–E70, 1986.
20. Lawler JM, Powers SK, Visser T, Van Dijk H, Kordus MJ, Ji LL. Acute exercise and skeletal muscle antioxidant and metabolic enzymes: effects of fiber type and age. Am J Physiol Regul Integr Comp Physiol 265: R1344–R1350, 1993.
21. Leeuwenburgh C, Fiebig R, Chandwaney R, Ji LL. Aging and exercise training in skeletal muscle: responses of glutathione and antioxidant enzyme systems. Am J Physiol Regul Integr Comp Physiol 267: R439–R445, 1994.
22. 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.
23. Murrant CL, Reid MB. Detection of reactive oxygen and reactive nitrogen species in skeletal muscle. Microsc Res Tech 55: 236–248, 2001.[CrossRef][ISI][Medline]
24. Nulton-Persson AC, Szweda LI. Modulation of mitochondrial function by hydrogen peroxide. J Biol Chem 276: 23357–23361, 2001.
25. Pitula JS, Deck KM, Clarke SL, Anderson SA, Vasanthakumar A, Eisenstein RS. Selective inhibition of the citrate-to-isocitrate reaction of cytosolic aconitase by phosphomimetic mutation of serine-711. Proc Natl Acad Sci USA 101: 10907–10912, 2004.
26. Sadek HA, Humphries KM, Szweda PA, Szweda LI. Selective inactivation of redox-sensitive mitochondrial enzymes during cardiac reperfusion. Arch Biochem Biophys 406: 222–228, 2002.[CrossRef][ISI][Medline]
27. Sakamoto K, Arnolds DE, Ekberg I, Thorell A, Goodyear LJ. Exercise regulates Akt and glycogen synthase kinase-3 activities in human skeletal muscle. Biochem Biophys Res Commun 319: 419–425, 2004.[CrossRef][ISI][Medline]
28. Sandström ME, Abbate F, Andersson DC, Zhang SJ, Westerblad H, Katz A. Insulin-independent glycogen supercompensation in isolated mouse skeletal muscle: role of phosphorylase inactivation. Pflügers Arch 448: 533–538, 2004.[ISI][Medline]
29. Sandström ME, Zhang SJ, Silva JP, Reid MB, Westerblad H, Katz A. Role of reactive oxygen species in contraction-mediated glucose transport in skeletal muscle. J Physiol 575: 251–262, 2006.
30. Sandström ME, Zhang SJ, Westerblad H, Katz A. Mechanical load plays little role in contraction-mediated glucose transport in mouse skeletal muscle. J Physiol 579: 527–534, 2007.
31. Sayen MR, Gustafsson AB, Sussman MA, Molkentin JD, Gottlieb RA. Calcineurin transgenic mice have mitochondrial dysfunction and elevated superoxide production. Am J Physiol Cell Physiol 284: C562–C570, 2003.
32. Schriner SE, Linford NJ, Martin GM, Treuting P, Ogburn CE, Emond M, Coskun PE, Ladiges W, Wolf N, Van Remmen H, Wallace DC, Rabinovitch PS. Extension of murine life span by overexpression of catalase targeted to mitochondria. Science 308: 1909–1911, 2005.
33. Spencer MK, Yan Z, Katz A. Carbohydrate supplementation attenuates IMP accumulation in human muscle during prolonged exercise. Am J Physiol Cell Physiol 261: C71–C76, 1991.
34. Tokheim AM, Martin BL. Association of calcineurin with mitochondrial proteins. Proteins 64: 28–33, 2006.[CrossRef][ISI][Medline]
This article has been cited by other articles:
|
|
 |
|
  J. P. McClung, N. E. Andersen, T. N. Tarr, C. H. Stahl, and A. J. Young Physical Activity Prevents Augmented Body Fat Accretion in Moderately Iron-Deficient Rats J. Nutr., July 1, 2008; 138(7): 1293 - 1297. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
) and EDL (
) muscles. Values are means ± SE for 7 muscles. C: aconitase protein expression in EDL extracts in the basal state and after repeated contractions. Top: Western blots for a heart sample and paired EDL muscles in the basal state and after repeated contractions (Stim). Bottom: mean ± SE values for aconitase protein expression given as % of the basal value (=100%) for each pair (n = 4; P > 0.05).
