Journal of Applied Physiology Ad Instruments
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

J Appl Physiol 101: 833-839, 2006. First published May 25, 2006; doi:10.1152/japplphysiol.01007.2005
8750-7587/06 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
101/3/833    most recent
01007.2005v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sahlin, K.
Right arrow Articles by Tonkonogi, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sahlin, K.
Right arrow Articles by Tonkonogi, M.

Repeated static contractions increase mitochondrial vulnerability toward oxidative stress in human skeletal muscle

Kent Sahlin,1,2,3 Jens Steen Nielsen,1 Martin Mogensen,1 and Michail Tonkonogi2,3,4

1Institute of Sport Sciences and Clinical Biomechanics, University of Southern Denmark, Odense, Denmark; 2Stockholm University College of Physical Education and Sports, Stockholm; 3Department of Physiology and Pharmacology, Karolinska Intitutet, Stockholm; and 4Department of Health and Social Sciences, Dalarna University, Falun, Sweden

Submitted 19 August 2005 ; accepted in final form 9 May 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Repeated static contractions (RSC) induce large fluctuations in tissue oxygen tension and increase the generation of reactive oxygen species (ROS). This study investigated the effect of RSC on muscle contractility, mitochondrial respiratory function, and in vitro sarcoplasmic reticulum (SR) Ca2+ kinetics in human muscle. Ten male subjects performed five bouts of static knee extension with 10-min rest in between. Each bout of RSC (target torque 66% of maximal voluntary contraction torque) was maintained to fatigue. Muscle biopsies were taken preexercise and 0.3 and 24 h postexercise from vastus lateralis. Mitochondria were isolated and respiratory function measured after incubation with H2O2 (HPX) or control medium (Con). Mitochondrial function was not affected by RSC during Con. However, RSC exacerbated mitochondrial dysfunction during HPX, resulting in decreased respiratory control index, decreased mitochondrial efficiency (phosphorylated ADP-to-oxygen consumed ratio), and increased noncoupled respiration (HPX/Con post- vs. preexercise). SR Ca2+ uptake rate was lower 0.3 vs. 24 h postexercise, whereas SR Ca2+ release rate was unchanged. RSC resulted in long-lasting changes in muscle contractility, including reduced maximal torque, low-frequency fatigue, and faster torque relaxation. It is concluded that RSC increases mitochondrial vulnerability toward ROS, reduces SR Ca2+ uptake rate, and causes low-frequency fatigue. Although conclusive evidence is lacking, we suggest that these changes are related to increased formation of ROS during RSC.

calcium homeostasis; exercise; mitochondria; oxidative phosphorylation

REPEATED STATIC CONTRACTIONS (RSC) are characterized by a contraction phase (high-energy turnover and restricted blood flow) followed by a relaxation phase with low-energy turnover, hyperemia, and elevated PO2. RSC performed at low force but over an extended period of time have been associated with muscular disorders (15, 20). The etiology is unclear, but is considered to be multifactorial, involving both physiological and psychological mechanisms (14). RSC performed at higher intensities are frequent in certain sports (e.g., wrestling, sailing, speedway) and are associated with fatigue. The physiological and biochemical responses in muscle to RSC have been investigated only in a few studies.

RSC are associated with oxidative stress, and the generation of reactive oxygen species (ROS) appears to be related to the exerted force (31). Prior treatment of diaphragm muscle fiber bundles with antioxidants (N-acetylcysteine) attenuates fatigue development during RSC (21). Recent studies in humans also demonstrate that administration of N-acetylcysteine during exercise can delay fatigue in endurance-trained subjects (27) and suggest that ROS generation may have an important role in exercise-induced fatigue. Potential sources of ROS during RSC are the electron transport chain and the hypoxanthine-xanthine oxidase system (24, 35). Mitochondria are not only a source but also a target of ROS, and RSC may therefore deteriorate mitochondrial function. RSC have several physiological characteristics in common with ischemia-reperfusion, which is a condition known to induce excessive ROS load and tissue damage (17, 24). Studies in rat heart demonstrate that ischemia-reperfusion reduces mitochondrial ADP-stimulated respiration (state 3) (12, 23) and potentiates ROS-induced reduction in state 3 respiration (22). The oxygen cost of contraction increases successively during RSC (33, 40), and it was suggested (40) that the mechanism was related to reduced efficiency of oxidative phosphorylation (oxphos) [i.e., reduced phosphorylated ADP-to-oxygen consumed (P/O) ratio]. However, the effect of RSC on mitochondrial function and the vulnerability of mitochondria to ROS have not been investigated previously.

RSC result in a long-lasting depression of muscle force, which appears to be independent of muscle acidosis (7, 33, 39, 40). The mechanism of fatigue after RSC is unclear, but it has been hypothesized that RSC result in impaired excitation-contraction (E-C) coupling (40). A characteristic feature of impaired E-C coupling is low-frequency fatigue (LFF) (i.e., more pronounced decrease of force at low than at high frequencies of stimulation). LFF has been observed in humans after RSC (39), and the mechanism may be related to reduced sarcoplasmic reticulum (SR) Ca2+ release (8) or alternatively to a decreased Ca2+ sensitivity of the contractile proteins. Repeated isometric tetani in single mouse muscle fibers resulted in reduced Ca2+ sensitivity, which could be prevented by prior treatment with ROS scavenger (28). It is also known that exposure of single fibers to ROS alters Ca2+ handling at several sites (2). In humans, 30 min of RSC resulted in reduced SR Ca2+ uptake and intrinsic alterations in the Ca2+ pump (39). However, neither SR Ca2+ release nor contractility kinetics (rate of force development and relaxation) have been measured after RSC, and further studies are therefore required to investigate this issue.

The purpose of this study was to test the hypothesis 1) that RSC deteriorate mitochondrial function and increase the vulnerability toward oxidative stress, and 2) that RSC alter muscle contractility and in vitro SR Ca2+ handling.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Subjects.   Ten healthy male subjects were investigated. Their age, weight, and height [mean (range)] were as follows: 26 (21–31) yr, 81 (65–98) kg, and 184 (179–190) cm. Total body fat, measured with impedance technique (Tanita body impedance TBF-300), was 15.5 ± 1.2% (mean ± SE). Subjects were mainly university students and were physically active on a recreational basis, including participation in various sports (in average two times per week) and cycling (average distance ~38 km/wk). Subjects agreed to participate in the experiment after having been informed of the purpose and potential risks involved. The project was approved by the local ethics committee at the Odense University Hospital (VF 20030129).

Experimental protocol.   Before the main experiment (within a week), subjects were familiarized with the protocol of electrical stimulation. Subjects were also trained to perform maximal voluntary isometric contractions (MVC) until they were able to reach maximal torque repeatedly. Subjects were informed to abstain from physical activity 24 h before the experiment and to use the same transportation to the laboratory on both experimental days.

Subjects performed static contraction with the knee extensors of one leg at a target torque corresponding to 66% of MVC until fatigue (Fig. 1). Both the actual torque and the target torque were displayed on a computer screen in front of the subject. Fatigue was defined as the time point when subjects, despite verbal encouragement, were unable to maintain the target torque (>2 s at 10% below target torque). The contraction was repeated five times with 10-min rest in between. Changes in muscle contractility were investigated with a standardized test (see below) before and after the exercise period. The test was (except for immediately postexercise) preceded by warm up (10-min cycling at 80–120 W) and performed on the same leg in each subject (randomly chosen between the dominant and nondominant leg). Capillary blood (prewarmed fingertip) and muscle biopsy were taken after the contractility test. The two postexercise muscle biopsies were taken from the exercised leg, and the preexercise biopsy was taken from the other leg. The first postexercise biopsy was taken ~7 min after the test and thus 18 min after terminating RSC (Fig. 1). The rationale for this time delay was to maximize ROS generation, which is expected to reach a peak during the reperfusion period. Potential acute effects of RSC-induced metabolic perturbations are unlikely to affect the measured mitochondrial function and SR Ca2+ kinetics, since the organelles were studied in standardized solutions during the measurements. The preexercise biopsy was taken from a leg without prior contractility test, and it may therefore be argued that the observed changes of muscle mitochondrial function and/or SR Ca2+ kinetics are due to the contractility test rather than RSC. However, this seems rather unlikely, since the total duration of contraction during the test (including mainly submaximal electrical stimulation) was only ~8 s compared with >220-s contraction during RSC.


Figure 1
View larger version (6K):
[in this window]
[in a new window]
  		Fig. 1. Experimental protocol. Pre, preexercise; WUP, warm-up period (8 min cycling at 80 W and 2 min cycling at 120 W); test, period of muscle contractility test with both voluntary and electrical stimulated contractions (see MATERIALS AND METHODS for details). The arrows denote time of muscle biopsy (NE, nonexercise; E, exercise). The stars denote time of capillary blood. The black bars denote repetitive static contractions (RSC). Each bout of RSC was maintained to fatigue at 66% of maximal voluntary contraction torque (MVC).

 
Muscle biopsy.   After local anesthesia (2–3 ml 2% lidocaine), an incision was made through the skin and fascia (about one-third of the distance between patella and spina ilica anterosuperior) of vastus lateralis muscle. The muscle biopsy was taken with a modified Bergström needle with suction (3). The muscle specimen was dried on a filter paper placed on a glass plate (cooled on ice), and obvious connective tissue and fat were removed. The muscle specimen was divided into three lots, of which one (~10 mg) was immediately frozen in liquid nitrogen and later used for analysis of citrate synthase (CS) activity (see below). Another lot (35–50 mg) was used for determination of SR Ca2+ kinetics and was weighed and homogenized in 1:10 volumes (wt/vol) of ice-cold buffer (300 mM sucrose, 1 mM EDTA, 10 mM NaN3, 40 mM Tris base, and 40 mM histidine at pH 7.8) at 0°C in a 1-ml glass homogenizer with a glass pestle (Kontes Glass Industry, Vineland, NJ). Before homogenization, the muscle sample was rinsed free of contaminating blood by washing in ice-cold buffer. This procedure is important to avoid artifacts due to the influence of blood on the assays of Ca2+ kinetics (30). The homogenate was divided in different portions and frozen in liquid nitrogen for later analyses of Ca2+ kinetics, protein, and myosin heavy chain (MHC) composition (see below). The last muscle portion (84–238 mg) was used for measurements of oxidative function of isolated mitochondria (see below).

Electrical stimulation and measurements of contractility.   Muscle contractility was measured in a custom-made chair (knee angle 90°) with the trunk and hip firmly fixed with belts and the contracting leg fixed (2 cm proximal to the malleoli) in a metal cuff connected to a strain-gauge.

The procedure of electrical stimulation and measuring muscle contractility was similar to that previously described (30). Briefly, two surface electrodes (5 x 9 cm, Pals Axelgaard, Fallbrook, CA) were placed on vastus lateralis muscle. The muscle was stimulated (Digitimer DS7A, Digitimer, Welwyn Garden City, UK) with square-waved pulses of 100 µs with a cutoff voltage of 230 V. The stimulation protocol and data sampling (1 kHz) were controlled from a computer by predesigned macros in Spike 2 version 4.18 (Cambridge Electronic Design, Cambridge, UK) and controlled through a CED Micron 1401 II 16-bit A/D converter (Cambridge Electronic Design).

The muscle was stimulated with a current sufficient to elicit either maximal torque (twitch) or 25% of peak twitch torque (20 and 50 Hz). The following protocol was used: 20 Hz (1 s), 2-min pause, 20 Hz (1 s), 0.5-min pause, 50 Hz (1 s), 0.5-min pause, 50 Hz (1 s), 0.5-min pause, 1 Hz. Peak torque at 20 and 50 Hz was determined as the maximal torque produced during 0.4 s. One-half relaxation time (RT1/2) was calculated as the time from the last stimuli to 50% of peak torque at 50 Hz, and one-half contraction time (CT1/2) as the time from the first stimuli until 50% of peak torque. The degree of tetanic fusion during 20-Hz stimulation was calculated from the torque fluctuation (difference between peak and nadir torque) during the last four stimulations. After electrical stimulation, subjects performed three MVC separated by 45 s. Peak torque, i.e., 100% MVC, was defined as the maximal torque maintained during 1 s. The contractile measurements were performed on the same leg, and the coefficients of variation (CVs), calculated previously from duplicate measurements (30), were 6.5% (RT1/2), 6.6% (MVC), 3.3% (20 Hz), and 1.2% (50 Hz).

SR Ca2+ uptake and release rates.   The method used to determine the rates of Ca2+ uptake and release has been described in detail (30). Briefly, muscle homogenate (80 µl ~8 mg of muscle tissue) was mixed with 1 ml buffer [165 mM KCl, 22 mM HEPES, 7.5 mM oxalate, 11 mM NaN3, 5.5 µM N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine, 20 µM CaCl2, 5 mM MgCl2, and 1.5 µM indo 1 (pH 7.0 at 37°C)], and the reaction was initiated by adding ATP to a final concentration of 5 mM. Ca2+ concentration ([Ca2+]) was determined fluorometrically (Ratiomaster RCM, Photon Technology International, Brunswick, NJ) at 37°C using the fluorescent Ca2+ indicator indo 1 (excitation wavelength 355 nm and emission wavelength 400 and 470 nm). The fluorescence was measured at 2 Hz over at least 250 s and converted to free [Ca2+], as previously described (30). After [Ca2+] had reached a plateau, Ca2+ release was initiated by adding 4-chloro-m-cresol (10 mM) (32), and the fluorescence followed during at least 30 s.

All raw data ([Ca2+] and absolute time) were imported into Matlab version 7.0.1 (The MathWorks, Natick, MA) and mathematically analyzed (Curve Fitting Toolbox version 1.1.1, The MathWorks). Initial free [Ca2+] was ≥800 nM in all assays of Ca2+ uptake. Data points between a free [Ca2+] of 800 nM and 20 s before initiating Ca2+ release were fitted to Eq. 1.

Formula 1(1)
where y is the free [Ca2+], t is time, and a, b, and c are constants assigned from Matlab. The rate of Ca2+ uptake at a free [Ca2+] of 800 nM, and the time for the free [Ca2+] to reach 63% of the initial free [Ca2+] ({tau} = 1/b) was calculated from Eq. 1. The data points during the first 30 s of release were mathematically fitted to Eq. 2.

Formula 2(2)
The initial rate of Ca2+ release was determined as the derivate in the first data point in the release process. The data points were well fitted to monoexponential functions, during both the uptake phase (r2 > 0.99) and the release phase (r2 > 0.99).

Protein content in the muscle homogenate was measured in triplicates with a standard kit (Pierce BCA protein reagent no. 23225). Assays of uptake and release rates of Ca2+ were performed in duplicates, and rates were expressed per milligram protein.

The CV as assessed in duplicate assays is larger for Ca2+ release rate (14%) than for Ca2+ uptake rate (5.5%), indicating a larger methodological variation and thus a lower statistical power of the former. The sampling rate (2 Hz) may limit the ability to accurately determine the rapid phase of Ca2+ release.

Fiber-type composition.   MHC composition was analyzed in the same muscle homogenate as used for measurements of Ca2+ kinetics using gel electrophoresis, as previously described (4) and modified for humans (1). Briefly, muscle homogenate (80 µl) was mixed with 200 µl of sample buffer (10% glycerol, 5% 2-mercaptoethanol and 2.3% SDS, 62.5 mM Tris and 0.2% bromophenolblue at pH 6.8), boiled in water bath at 100°C for 3 min, and loaded (10–40 µl) on a SDS-PAGE gel [8% polyacrylamide (100:1 acrylamide-bisacrylamide), 30% glycerol, 67.5 mM Tris base, 0.4% SDS, and 0.1 M glycine]. Gels were run at 80 V for at least 42 h at 4°C, and MHC bands were made visible by staining with Coomassie. The gels were scanned (Linoscan 1400 scanner, Heidelberg, Germany), and MHC bands were quantified densitometrically (Phoretix 1D, nonlinear, Newcastle, UK).

Mitochondrial function.   Isolation of mitochondria was performed as previously described (37). Briefly, muscle specimen was disintegrated with scissors, and mitochondria were isolated by proteinase treatment (Nagarse, 0.2 mg/ml, Sigma P-4789), followed by homogenization and subsequent differential centrifugation. The final mitochondrial pellet was resuspended (0.4 µl/mg initial muscle) in a medium consisting of (in mmol/l) 225 mannitol, 75 sucrose, 10 Tris, 0.1 EDTA, pH 7.40, and stored on ice until analysis.

Oxygen consumption in isolated mitochondria was measured polarographically using a Clark-type electrode (Hansatech DW1) in a water-jacketed glass chamber at a volume of 0.3 ml. The measurements were carried out at 25°C. Respiration in isolated mitochondria was analyzed in a reaction medium containing (in mmol/l) 225 mannitol, 75 sucrose, 10 Tris, 10 KCl, 10 K2HPO4, 0.1 EDTA, 5 pyruvate, 2 malate, 0.08 MgCl2, pH 7.35. Mitochondrial suspension was added to the reaction medium, and state 3 respiration was initiated by the addition of ADP (final concentration 260 µM). When all of the added ADP had been phosphorylated to ATP, the respiratory rate returned to that before the addition of ADP (state 4). To elucidate potential loss of cytochrome c, state 3 was repeated after addition of cytochrome c. Respiratory control index (RCI) was calculated as the ratio of the respiratory rate in state 3 to that in state 4. The P/O ratio was quantified according to the method of Chance and Williams (9).

Mitochondria were incubated at 0°C for 30 min with freshly prepared H2O2 [final concentration 0.095 mM (HPX) or with suspension medium (Con)]. The order between HPX and Con was randomized between subjects but maintained constant in biopsies taken from the same subject.

Mitochondrial Ca2+ resistance.   Mitochondrial Ca2+ resistance (MCaR) for opening of mitochondrial permeability transition pore was determined on isolated mitochondria, as previously described (16). An aliquot of mitochondrial suspension equivalent to 5 x 10–2 units CS, corresponding to 19.1 µg protein, was resuspended in 100 µl of buffer (pH 7.4) containing 150 mM KCl, 5 mM Tris, 5 mM K2HPO4, 5 mM malate, and 5 mM pyruvate. Equivalent pulses of CaCl2 (2 µmol of Ca2+ per unit CS) were added to the mitochondria every 3 min, and mitochondrial swelling was monitored continuously as the change in absorbance at 540 nm until absorbance decreased rapidly. A rapid decrease in absorbance (limit >0.0175 absorbance units/min) indicates progressive swelling of mitochondria due to opening of mitochondrial permeability transition pore.

Plasma CK and thiobarbituric acid-reactive substances and muscle CS activity.   Plasma creatine kinase activity was analysed with a standardized kit (Vitros, Ortho-clinical Diagnostic, Indigo Creek, NY). Total content of thiobarbituric acid reactive substances (TBARS) was analyzed in plasma using a fluorometrical method (6). Briefly, plasma (10 µl) was precipitated with phosphothungstic acid (10%), resuspended in a thiobarbituric acid reagent (0.67%), heated in water bath at 100°C for 60 min, and extracted with butanol. The fluorescence of the butanol phase was measured (excitation 515 nm, emission 545 nm), and TBARS was estimated with a standard curve constructed from tetraethoxypropane. All plasma samples from each subject were analyzed simultaneously and in duplicates (CV 10%).

CS activity was measured spectrophotometrically (25°C) in isolated mitochondria and in freeze-dried muscle and dissected free from nonmuscle constituents, using a technique described previously (37).

Mitochondrial lipid peroxidation (4-hydroxynonenal).   Mitochondrial 4-hydroxynonenal (HNE) protein adducts were determined with Western blotting of samples taken pre- and 0.3 h postexercise. Equal amounts of mitochondria (as judged from CS activity) were loaded on SDS-polyacrylamide gels. After electrophoresis, proteins were transferred to a polyvinylidene difluoride membrane, and HNE protein adducts were detected with polyclonal antibodies (HNE12-S, Alpha Diagnosics, San Antonio, TX). After incubation with secondary antibodies conjugated with horseradish peroxidase, the membrane was incubated with detection reagent (ECL, Amersham Biosciences). An X-ray film was exposed to the membrane, and the optical density of the whole lane was quantified by using Molecular Analyst 1.5 (Bio-Rad).

Statistical methods.   Values are presented as means ± SE. Statistical significance of differences was tested with a nonparametric analysis of variance (Friedman's test) followed by post hoc analysis with Wilcoxon's signed-rank test. Correlations between parameters were performed with Spearman rank correlation. Statistical significance was accepted at 5% level. All statistical analysis was conducted in Statview version 5.0 (SAS Institute, Cary, NC).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Muscle fiber-type distribution is reflected by the relative composition of MHC isozymes. MHC composition was not different between legs. When calculated from the average of the three biopsies from each subject, MHC composition was [mean ± SE (range)]: MHC I, 42.5 ± 2.2% (30.8–51.7%); MHC IIA, 56.8 ± 2.2% (47.9–68.0%) and MHC IIX, 0.9 ± 0.3% (0–1.9%).

The endurance during RSC decreased successively, being 52 ± 3, 47 ± 3, 42 ± 2, 40 ± 3, and 38 ± 3 s for the five sustained bouts of isometric contractions at 66% of MVC. Endurance was 26% lower during the last vs. the initial bout (P < 0.05). The sum of endurance during the five bouts (220 ± 13 s) was not correlated to the actual values or the change in values (post – pre) of mitochondrial function or Ca2+ kinetics. Maximal torque declined after RSC, and the depression was more pronounced during low-frequency stimulations (30% decline in twitch torque, P < 0.05) than during MVC (–14%; P < 0.05; Table 1). This was also reflected by a decline in 20- to 50-Hz force ratio (–15%, P < 0.05), which remained depressed 24 h postexercise (–11%; P < 0.05). The decline in 20- to 50-Hz torque ratio was related to the decline in twitch torque (r = 0.83, P < 0.01). RT1/2, measured before RSC, was inversely related to the degree of torque fluctuation at 20 Hz (r = –0.84, P < 0.01; Fig. 2) and suggests that both parameters are related to torque relaxation. Following RSC, RT1/2 decreased and torque fluctuation increased (Table 1). Both of these changes demonstrate that torque relaxation become faster after RSC.


View this table:
[in this window]
[in a new window]
  		Table 1. Effect of RSC on muscle contractility and in vitro Ca2+ kinetics

 

Figure 2
View larger version (5K):
[in this window]
[in a new window]
  		Fig. 2. Correlation between one-half relaxation time (RT1/2) and fluctuation of torque at 20 Hz. RT1/2 is the time between last stimuli (50 Hz) and decline of torque by 50%. Torque fluctuation denotes the relative degree of unfused torque, i.e., the average difference between the peak and nadir of torque (normalized to peak torque) during the last 4 stimuli. The figure shows values obtained preexercise (r = –0.84; P < 0.01, n = 10). RT1/2 was also inversely correlated to torque fluctuation when measured at 3 h (r = 0.72; P < 0.05) and 24 h (r = 0.82; P < 0.01) postexercise but not at 0.3 h postexercise.

 
Ca2+ kinetics was measured in muscle homogenates. The average rate of SR Ca2+ uptake shortly after RSC was 11% lower than that after 24-h recovery (P < 0.05, Table 1) and that preexercise, albeit the latter did not reach statistical significance. SR Ca2+ release rate was not significantly changed (P > 0.05) by RSC.

RSC had no effect on mitochondrial function [state 3 and state 4 respiration, P/O ratio, resistance to Ca2+-induced pore opening (MCaR) during Con; Table 2]. In vitro exposure of mitochondria to ROS resulted in lower state 3 respiration (–47%; P < 0.001), lower RCI (–43%; P < 0.001), and lower P/O ratio (–4%; P < 0.01). The ROS-induced impairment of mitochondrial function was exacerbated by RSC. When related to Con, there was a number of changes in mitochondrial function after RSC during HPX (Fig. 3), including reduced RCI (P < 0.05, pre- vs. 0.3 and 24 h postexercise), reduced P/O ratio (P < 0.05, pre- vs. 0.3 h postexercise), and increased noncoupled respiration (state 4) (P < 0.05, pre- vs. 24 h postexercise). Addition of cytochrome c could not reverse the ROS-induced deterioration of mitochondrial respiration (data not shown).


View this table:
[in this window]
[in a new window]
  		Table 2. Effect of RSC on mitochondrial function during control

 

Figure 3
View larger version (22K):
[in this window]
[in a new window]
  		Fig. 3. Vulnerability of mitochondrial parameters to oxidative stress. Mitochondria were isolated from muscle biopsies taken preexercise and 0.3 and 24 h postexercise and incubated in 0.1 mM H2O2 (HPX) or mitochondrial suspension medium (Con) for 30 min. State 3 and state 4 denote mitochondrial respiration with pyruvate-malate in the presence and absence of ADP, respectively. RCI, respiratory control index (state 3/state 4); P/O, ratio between phosphorylated ADP to oxygen consumed. Values (means ± SE) are shown as percentage of corresponding values during Con. *P < 0.05 vs. preexercise. {dagger}P < 0.05 vs. 0.3 h postexercise.

 
Plasma CK activity increased moderately after RSC and was 57% higher 24 h postexercise than that preexercise (333 ± 54 vs. 212 ± 70 U/l plasma; P < 0.001). There was no evidence of increased lipid peroxidation when measured as the concentration of TBARS in plasma [6.8 ± 0.7 (pre), 7.2 ± 0.3 (0 h), 6.9 ± 0.4 (3 h), and 7.5 ± 0.3 (24 h) µmol/l plasma, P > 0.05] or in urine (data not shown). The sensitivity of this method to detect local oxidative stress in muscle tissue may, however, be low (see DISCUSSION). Mitochondrial lipid peroxidation was measured as HNE protein adducts with Western blotting. There was no significant difference in HNE protein adducts in mitochondria isolated from muscle samples taken pre- and postexercise (93 ± 10% of preexercise value).


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Mitochondrial function.   In vitro ROS exposure of mitochondria from preexercise biopsies resulted in a pronounced decline in state 3 respiration, RCI, and P/O ratio (Fig. 3). Mitochondria isolated from muscle biopsies taken after RSC was more vulnerable to ROS exposure, as demonstrated by further depressions of RCI and oxphos, as well as increased noncoupled respiration (Fig. 3). These alterations remained when cytochrome c was added to the mitochondria, and thus the defects could not be attributed to a loss of cytochrome c due to damaged outer mitochondrial membrane. The augmented mitochondrial vulnerability to ROS after RSC is similar to that observed in rat cardiac tissue after ischemia-reperfusion (23), which was attributed to depletion of the mitochondrial antioxidant reserve (i.e., glutathione, GSH). Following ischemia-reperfusion of rat cardiac tissue, GSH decreased by 10 and 30% in the myocardium and mitochondria, respectively (23). Strenuous exercise in humans has previously been shown to reduce GSH in plasma (18) and in muscle (10, 27). Intravenous infusion with an antioxidant (N-acetylcysteine) prevented the exercise-induced fall in total glutathione and reduced glutathione in muscle of endurance-trained subjects (27). Furthermore, patients with circulatory shock exhibit both reduced GSH levels and mitochondrial dysfunction in skeletal muscle (11). Due to limited material, we were unable to measure mitochondrial glutathione status. However, because RSC is an ischemia-reperfusion type of exercise, it is possible that the increased mitochondrial vulnerability to ROS after RSC was caused by a reduction of the mitochondrial antioxidant reserve (e.g., GSH).

There are some studies that indicate that the vulnerability to oxidative stress is increased by endurance training. First, the ergogenic effect of intravenous infusion of antioxidants (N-acetylcysteine) was dependent on the training status of the subjects. Performance increased by 26% in endurance-trained subjects (27) but remained unchanged in recreationally active men (peak O2 uptake < 48 ml O2·min–1·kg body wt–1) (26). Furthermore, mitochondrial vulnerability to oxidative stress is increased in rats after rigorous swim training (23). The subjects included in the present study were active on a recreational basis, but were not classified as athletes. Although mitochondrial function was unaffected by RSC during Con, it can be speculated that mitochondria from endurance-trained athletes would be more susceptible to RSC than that from the present group of subjects.

Based on experiments in horses, it was concluded that maximal mitochondrial respiration was severely reduced after exhaustive exercise (19). However, later studies in humans, in whom more advanced techniques have been used, have not been able to confirm these data. Mitochondrial respiratory function, including oxphos, was maintained following fatiguing exercise of long duration (36), at high intensity (38), or of eccentric type (41). The results from the present study extend this list to RSC, which had no acute influence on mitochondrial function, including oxphos.

Muscle contractility and Ca2+ kinetics.   Endurance at 66% MVC decreased successively during RSC. The relatively slow recovery of endurance (10-min rest between bouts) is consistent with previous findings (34) and is likely related to remaining high muscle lactate. MVC decreased by 14% after RSC (~10 min postexercise) and remained depressed 3 h postexercise. Previous studies have shown that muscle metabolic state is recovered 1 h after RSC, but that MVC remains reduced (39). The long-lasting depression of MVC observed in the present and previous studies (39) is, therefore, most likely caused by nonmetabolic factors, e.g., reduced SR Ca2+ release and/or reduced myofibrillar Ca2+ sensitivity. The observed long-lasting LFF (reduced 20- to 50-Hz torque ratio) after RSC is consistent with previous findings (39) and supports the hypothesis of disturbances in Ca2+ handling.

LFF is most likely a consequence of reduced cytosolic Ca2+ levels and/or reduced myofibrillar Ca2+ sensitivity. The rate of SR Ca2+ release, as measured in vitro, was not reduced after RSC (Table 1). However, this method only measures a late step in E-C coupling, and we cannot exclude that Ca2+ release in vivo is reduced due to disturbances in t-tubular function, defects in signaling between the t-tubular and the SR system, or reduced SR content of Ca2+. Studies in isolated mouse muscle fibers showed that myofibrillar Ca2+ sensitivity decreased after RSC performed at body temperature (37°C) and that this could be prevented by ROS scavengers (28). Furthermore, exposure of muscle fibers to ROS (without RSC) reduced myofibrillar Ca2+ sensitivity (2). Thus it seems reasonable to suggest that the reduction in torque at low frequencies, which was observed after RSC, is caused by ROS-induced reduction in myofibrillar Ca2+ sensitivity and/or reduced SR Ca2+ release. Further studies are required to investigate this hypothesis.

The rate of SR Ca2+ uptake was lower in muscle biopsies taken shortly after RSC than those after 24-h recovery. The reduced rate of SR Ca2+ uptake is consistent with a previous study in humans, in whom 30 min of RSC resulted in a pronounced reduction in SR Ca2+ uptake rate and SR Ca2+ ATPase activity (39). Exposure of single fibers to ROS reduces the rate of Ca2+ removal (2), and the depression of SR Ca2+ uptake after RSC may, therefore, be an effect of increased ROS load. Conclusive evidence for a cause-effect relation is, however, lacking.

Torque relaxation is determined by the rate of Ca2+ removal from the cytosol and the rate of cross-bridge turnover. Everything else being the same, one would expect that a reduced maximal rate of Ca2+ uptake would result in a slower torque relaxation. However, previous studies have demonstrated that, despite a depressed Ca2+ pump function, relaxation of force was unchanged after prolonged exercise (5) or even more rapid after eccentric exercise (30). Similarly, in the present study, we observed a faster torque relaxation after RSC, despite a lower rate of Ca2+ uptake (P < 0.05; 0.3 h vs. 24 h postexercise). The anomaly between faster torque relaxation and slower maximal Ca2+ uptake may relate to changes in other parts of Ca2+ handling (i.e., reduced myofibrillar Ca2+ sensitivity or reduced in vivo Ca2+ release).

Unchanged mitochondrial resistance to Ca2+ overload.   In a previous study, endurance exercise increased MCaR with almost 50% (16), and we suggested that this could serve as a protection of mitochondria to exercise-induced Ca2+ exposure. Preconditioning by exposure of muscle to brief periods of ischemia or contraction is known to protect cardiac tissue from subsequent periods of ischemia-reperfusion in (13), and skeletal muscle from, contraction-induced damage (25). There is evidence that the mechanism is related to activation of mitochondrial ATP-sensitive potassium channels, which attenuates mitochondrial Ca2+ overload (29). Being an ischemia-reperfusion type of exercise, RSC could be regarded as a form of preconditioning and thus expected to attenuate mitochondrial Ca2+ overload during subsequent periods of contractile activity. MCaR was, however, not significantly changed after RSC. However, MCaR is a measure of how much Ca2+ can be taken up by mitochondria before pore opening and does not include factors such as rate of Ca2+ accumulation or degree of Ca2+ overload. The unchanged MCaR does not exclude that RSC prevents mitochondria from Ca2+ overload.

Methodological considerations.   Mitochondrial function can be studied with a number of techniques, including isolated mitochondria and permeabilized fibers. The reason for using isolated mitochondria in this study was that this model, in contrast to permeabilized fibers, can be used to assess intrinsic mitochondrial functional parameters, such as efficiency (P/O ratio), RCI, and PTP opening. An inherent limitation with this model is that only a fraction of the available mitochondria is harvested. The yield of harvested mitochondria was rather high in this study (~28%), and there were no systematic differences in the yield between pre- and postexercise biopsies. However, we cannot exclude that the results of this study primarily reflect that of a subpopulation of mitochondria (e.g., subsarcolemmal mitochondria).

The rationale for the experimental protocol (i.e., RSC) was to maximize ROS generation. The model will induce large fluctuations of muscle oxygen tension with anoxia during the main part of the contraction and rapid, elevated PO2 during the hyperemic recovery phase. This ischemic-reperfusion model is known to be associated with increased ROS generation. Furthermore, exercise to fatigue is known to induce adenine nucleotide catabolism (34), and the associated increase in substrate (hypoxanthine) and conversion of xanthine dehydrogenase into xanthine oxidase would provide a further source of ROS. Lipid peroxidation products (TBARS) were measured in plasma and urine but could not demonstrate an increased ROS load after RSC. However, it could be argued that the working muscle was too small (one-leg knee-extension) to influence markers of ROS (lipid peroxidation products) in plasma and urine. Furthermore, we measured the degree of lipid peroxidation in mitochondria by analyzing HNE protein adducts. Again, we were unsuccessful in demonstrating exercise-induced lipid peroxidation, i.e., markers of tissue damage.

We observed unchanged mitochondrial function after RSC but increased vulnerability to ROS exposure. It is possible that the antioxidative defense (e.g., glutathione) was sufficient to protect mitochondria against RSC-induced ROS load, but that this caused a reduction of the mitochondrial antioxidant reserve, rendering the mitochondria more vulnerable to subsequent ROS exposure. The absence of increased mitochondrial lipid peroxidation is consistent with this scenario.

In summary, RSC resulted in 1) increased vulnerability of mitochondrial function to ROS exposure, 2) reduced rate of SR Ca2+ uptake (0 h vs. 24 h postexercise), 3) long-lasting muscle fatigue, especially at low frequencies, and 4) a faster torque relaxation. Previous studies have demonstrated that most of these changes can be provoked by exposure of the muscle or mitochondria to ROS or are associated with ROS-generating conditions. Measurements of lipid peroxidation markers in plasma (TBARS) or mitochondria (HNE protein adducts) could not verify the hypothesis of an increased ROS load after RSC. Although conclusive evidence is lacking, we suggest that the observed changes are related to oxidative stress induced by RSC.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
ACKNOWLEDGMENTS
 REFERENCES
 
The study received economical support from The Danish Medical Research Council (22–02-0346) and The Swedish Research Council (project 13020).


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
REFERENCES
 
We thank the subjects for cooperation during the experiments.


   FOOTNOTES
 

Address for reprint requests and other correspondence: K. Sahlin, GIH, Stockholm Univ. College of Physical Education and Sports, Box 5626, SE 11486 Stockholm, Sweden

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


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 

  1. Andersen JL and Aagaard P. Myosin heavy chain IIX overshoot in human skeletal muscle. Muscle Nerve 23: 1095–1104, 2000.[CrossRef][Web of Science][Medline]
  2. Andrade FH, Reid MB, Allen DG, and Westerblad H. Effect of hydrogen peroxide and dithiothreitol on contractile function of single skeletal muscle fibres from the mouse. J Physiol 509: 565–575, 1998.[Abstract/Free Full Text]
  3. Bergstrom J. Percutaneous needle biopsy of skeletal muscle in physiological and clinical research. Scand J Clin Lab Invest 35: 609–616, 1975.[Medline]
  4. Betto DD, Zerbato E, and Betto R. Type 1, 2A, and 2B myosin heavy chain electrophoretic analysis of rat muscle fibers. Biochem Biophys Res Commun 138: 981–987, 1986.[CrossRef][Web of Science][Medline]
  5. Booth J, McKenna MJ, Ruell PA, Gwinn TH, Davis GM, Thompson MW, Harmer AR, Hunter SK, and Sutton JR. Impaired calcium pump function does not slow relaxation in human skeletal muscle after prolonged exercise. J Appl Physiol 83: 511–521, 1997.[Abstract/Free Full Text]
  6. Brown RK and Kelly FJ. Peroxides and other products. In: Free Radicals, A Practical Approach, edited by Punchard NA and Kelly FJ. Oxford, UK: Oxford University Press, 1996, p. 119–145.
  7. Bruton JD, Lannergren J, and Westerblad H. Effects of CO2-induced acidification on the fatigue resistance of single mouse muscle fibers at 28°C. J Appl Physiol 85: 478–483, 1998.[Abstract/Free Full Text]
  8. Bruton JD, Lannergren J, and Westerblad H. Mechanisms underlying the slow recovery of force after fatigue: importance of intracellular calcium. Acta Physiol Scand 162: 285–293, 1998.[CrossRef][Web of Science][Medline]
  9. Chance B and Williams GR. Respiratory enzymes in oxidative phosphorylation. I. Kinetics of oygen utilization. J Biol Chem 217: 383–393, 1955.[Free Full Text]
  10. Cooper MB, Jones DA, Edwards RH, Corbucci GC, Montanari G, and Trevisani C. The effect of marathon running on carnitine metabolism and on some aspects of muscle mitochondrial activities and antioxidant mechanisms. J Sports Sci 4: 79–87, 1986.[Medline]
  11. Corbucci GG, Gasparetto A, Candiani A, Crimi G, Antonelli M, Bufi M, De Blasi RA, Cooper MB, and Gohil K. Shock-induced damage to mitochondrial function and some cellular antioxidant mechanisms in humans. Circ Shock 15: 15–26, 1985.[Web of Science][Medline]
  12. Crestanello JA, Doliba NM, Babsky AM, Niibori K, Osbakken MD, and Whitman GJ. Mitochondrial function during ischemic preconditioning. Surgery 131: 172–178, 2002.[CrossRef][Web of Science][Medline]
  13. Crestanello JA, Doliba NM, Babsky AM, Niibori K, Whitman GJ, and Osbakken MD. Ischemic preconditioning improves mitochondrial tolerance to experimental calcium overload. J Surg Res 103: 243–251, 2002.[CrossRef][Web of Science][Medline]
  14. Edwards RH. Hypotheses of peripheral and central mechanisms underlying occupational muscle pain and injury. Eur J Appl Physiol 57: 275–281, 1988.
  15. Edwards RH, Hill DK, Jones DA, and Merton PA. Fatigue of long duration in human skeletal muscle after exercise. J Physiol 272: 769–778, 1977.[Abstract/Free Full Text]
  16. Fernstrom M, Tonkonogi M, and Sahlin K. Effects of acute and chronic endurance exercise on mitochondrial uncoupling in human skeletal muscle. J Physiol 554: 755–763, 2004.[Abstract/Free Full Text]
  17. Ferrari R, Guardigli G, Mele D, Percoco GF, Ceconi C, and Curello S. Oxidative stress during myocardial ischaemia and heart failure. Curr Pharm Des 10: 1699–1711, 2004.[CrossRef][Web of Science][Medline]
  18. Gohil K, Viguie C, Stanley WC, Brooks GA, and Packer L. Blood glutathione oxidation during human exercise. J Appl Physiol 64: 115–119, 1988.[Abstract/Free Full Text]
  19. Gollnick PD, Bertocci LA, Kelso TB, Witt EH, and Hodgson DR. The effect of high-intensity exercise on the respiratory capacity of skeletal muscle. Pflügers Arch 415: 407–413, 1990.[CrossRef][Web of Science][Medline]
  20. Hagberg M and Wegman DH. Prevalence rates and odds ratios of shoulder-neck diseases in different occupational groups. Br J Ind Med 44: 602–610, 1987.[Web of Science][Medline]
  21. Khawli FA and Reid MB. N-acetylcysteine depresses contractile function and inhibits fatigue of diaphragm in vitro. J Appl Physiol 77: 317–324, 1994.[Abstract/Free Full Text]
  22. Leichtweis SB, Leeuwenburgh C, Chandwaney R, Parmelee DJ, and Ji LL. Ischaemia-reperfusion induced alterations of mitochondrial function in hypertrophied rat heart. Acta Physiol Scand 156: 51–60, 1996.[CrossRef][Web of Science][Medline]
  23. Leichtweis SB, Leeuwenburgh C, Parmelee DJ, Fiebig R, and Ji LL. Rigorous swim training impairs mitochondrial function in post-ischaemic rat heart. Acta Physiol Scand 160: 139–148, 1997.[CrossRef][Web of Science][Medline]
  24. Li C and Jackson RM. Reactive species mechanisms of cellular hypoxia-reoxygenation injury. Am J Physiol Cell Physiol 282: C227–C241, 2002.[Abstract/Free Full Text]
  25. McArdle F, Spiers S, Aldemir H, Vasilaki A, Beaver A, Iwanejko L, McArdle A, and Jackson MJ. Preconditioning of skeletal muscle against contraction-induced damage: the role of adaptations to oxidants in mice. J Physiol 561: 233–244, 2004.[Abstract/Free Full Text]
  26. Medved I, Brown MJ, Bjorksten AR, and McKenna MJ. Effects of intravenous N-acetylcysteine infusion on time to fatigue and potassium regulation during prolonged cycling exercise. J Appl Physiol 96: 211–217, 2004.[Abstract/Free Full Text]
  27. Medved I, Brown MJ, Bjorksten AR, Murphy KT, Petersen AC, Sostaric S, Gong X, and McKenna MJ. N-acetylcysteine enhances muscle cysteine and glutathione availability and attenuates fatigue during prolonged exercise in endurance-trained individuals. J Appl Physiol 97: 1477–1485, 2004.[Abstract/Free Full Text]
  28. Moopanar TR and Allen DG. Reactive oxygen species reduce myofibrillar Ca2+ sensitivity in fatiguing mouse skeletal muscle at 37°C. J Physiol 564: 189–199, 2005.[Abstract/Free Full Text]
  29. Murata M, Akao M, O'Rourke B, and Marban E. Mitochondrial ATP-sensitive potassium channels attenuate matrix Ca(2+) overload during simulated ischemia and reperfusion: possible mechanism of cardioprotection. Circ Res 89: 891–898, 2001.[Abstract/Free Full Text]
  30. Nielsen JS, Madsen K, Jorgensen LV, and Sahlin K. Effects of lengthening contraction on calcium kinetics and skeletal muscle contractility in humans. Acta Physiol Scand 184: 203–214, 2005.[CrossRef][Web of Science][Medline]
  31. O'Neill CA, Stebbins CL, Bonigut S, Halliwell B, and Longhurst JC. Production of hydroxyl radicals in contracting skeletal muscle of cats. J Appl Physiol 81: 1197–1206, 1996.[Abstract/Free Full Text]
  32. Ortenblad N, Sjogaard G, and Madsen K. Impaired sarcoplasmic reticulum Ca2+ release rate after fatiguing stimulation in rat skeletal muscle. J Appl Physiol 89: 210–217, 2000.[Abstract/Free Full Text]
  33. Sahlin K, Cizinsky S, Warholm M, and Hoberg J. Repetitive static muscle contractions in humans–a trigger of metabolic and oxidative stress? Eur J Appl Physiol 64: 228–236, 1992.[CrossRef][Web of Science]
  34. Sahlin K and Ren JM. Relationship of contraction capacity to metabolic changes during recovery from a fatiguing contraction. J Appl Physiol 67: 648–654, 1989.[Abstract/Free Full Text]
  35. Sjodin B, Hellsten Westing Y, and Apple FS. Biochemical mechanisms for oxygen free radical formation during exercise. Sports Med 10: 236–254, 1990.[Web of Science][Medline]
  36. Tonkonogi M, Harris B, and Sahlin K. Mitochondrial oxidative function in human saponin-skinned muscle fibres: effects of prolonged exercise. J Physiol 510: 279–286, 1998.[Abstract/Free Full Text]
  37. Tonkonogi M and Sahlin K. Rate of oxidative phosphorylation in isolated mitochondria from human skeletal muscle: effect of training status. Acta Physiol Scand 161: 345–353, 1997.[CrossRef][Web of Science][Medline]
  38. Tonkonogi M, Walsh B, Tiivel T, Saks V, and Sahlin K. Mitochondrial function in human skeletal muscle is not impaired by high intensity exercise. Pflügers Arch 437: 562–568, 1999.[CrossRef][Web of Science][Medline]
  39. Tupling R, Green H, Grant S, Burnett M, and Ranney D. Postcontractile force depression in humans is associated with an impairment in SR Ca2+ pump function. Am J Physiol Regul Integr Comp Physiol 278: R87–R94, 2000.[Abstract/Free Full Text]
  40. Vollestad NK, Sejersted OM, Bahr R, Woods JJ, and Bigland-Ritchie B. Motor drive and metabolic responses during repeated submaximal contractions in humans. J Appl Physiol 64: 1421–1427, 1988.[Abstract/Free Full Text]
  41. Walsh B, Tonkonogi M, Malm C, Ekblom B, and Sahlin K. Effect of eccentric exercise on muscle oxidative metabolism in humans. Med Sci Sports Exerc 33: 436–441, 2001.[CrossRef][Web of Science][Medline]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
101/3/833    most recent
01007.2005v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sahlin, K.
Right arrow Articles by Tonkonogi, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sahlin, K.
Right arrow Articles by Tonkonogi, M.

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