Abstract
Recent studies report that depletion and repletion of muscle taurine (Tau) to endogenous levels affects skeletal muscle contractility in vitro. In this study, muscle Tau content was raised above endogenous levels by supplementing male Sprague-Dawley rats with 2.5% (wt/vol) Tau in drinking water for 2 wk, after which extensor digitorum longus (EDL) muscles were examined for in vitro contractile properties, fatigue resistance, and recovery from fatigue after two different high-frequency stimulation bouts. Tau supplementation increased muscle Tau content by ∼40% and isometric twitch force by 19%, shifted the force-frequency relationship upward and to the left, increased specific force by 4.2%, and increased muscle calsequestrin protein content by 49%. Force at the end of a 10-s (100 Hz) continuous tetanic stimulation was 6% greater than controls, while force at the end of the 3-min intermittent high-frequency stimulation bout was significantly higher than controls, with a 12% greater area under the force curve. For 1 h after the 10-s continuous stimulation, tetanic force in Tau-supplemented muscles remained relatively stable while control muscle force gradually deteriorated. After the 3-min intermittent bout, tetanic force continued to slowly recover over the next 1 h, while control muscle force again began to decline. Tau supplementation attenuated F2-isoprostane production (a sensitive indicator of reactive oxygen species-induced lipid peroxidation) during the 3-min intermittent stimulation bout. Finally, Tau transporter protein expression was not altered by the Tau supplementation. Our results demonstrate that raising Tau content above endogenous levels increases twitch and subtetanic and specific force in rat fast-twitch skeletal muscle. Also, we demonstrate that raising Tau protects muscle function during high-frequency in vitro stimulation and the ensuing recovery period and helps reduce oxidative stress during prolonged stimulation.
Keywords: lipid peroxidation, reactive oxygen species, muscle damage, calpains, low-frequency fatigue
taurine (Tau; 2-aminoethanesulfonic acid), a nontoxic β-amino acid found in most mammalian cells, is reported to function as an osmotic regulator, an antioxidant, a cell membrane stabilizer, a modulator of inflammation, and an intracellular ion regulator (esp. calcium, Ca2+) (for review see Ref. 31). Tau is found in particularly high concentrations (mM) in excitable tissues such as neurons and skeletal muscle (31). Early studies on the function of Tau in skeletal muscle revealed a role in membrane phospholipid stabilization and intracellular Ca2+ regulation, with Tau increasing the rate of sarcoplasmic reticulum (SR) Ca2+ uptake and total storage capacity in SR vesicles (30). More recently, Bakker and Berg (6), using rat fast-twitch extensor digitorum longus (EDL) mechanically skinned fiber preparations, in which most endogenous Tau would be washed out, showed that the readdition of approximately endogenous levels of Tau increased depolarization-induced force responses by ∼20%, despite a small reduction in the sensitivity of the contractile apparatus to Ca2+. In addition, Tau increased SR Ca2+ loading, indicative of increased SR Ca2+ pump activity, and increased the peak, and rate of rise, of caffeine-induced force responses, indicating greater releasable Ca2+ and/or altered activity of SR Ca2+ release channels (6). In a follow-up study, Hamilton et al. (27) reduced endogenous mouse EDL Tau content with the Tau transporter (TauT) uptake inhibitor guanidinoethane sulfonate (GES) and showed dramatic reductions in peak twitch force and a shift to the right of the force-frequency relationship. Studies using a mouse TauT−/− knockout model showed that with almost complete depletion of muscle Tau exercise capacity was reduced by >80% (71). Collectively, these studies provide strong evidence that Tau plays a critical role in modulating skeletal muscle contractile properties. To date, however, no studies have investigated the effect of raising Tau content above endogenous levels on skeletal muscle contractile properties and fatigue resistance in vitro.
There is also mounting evidence that Tau plays a cytoprotective role in various tissues under various conditions (for reviews see Refs. 11, 36, 66). For example, McLaughlin et al. (41) showed that exogenous Tau protected muscle twitch and tetanic force production in the early and late stages of ischemia-reperfusion injury. Moreover, Tau-depleted TauT−/− mice have resting plasma creatine kinase levels (an indicator of muscle plasma membrane damage) double those of wild types (71). This protective effect may be related to Tau's ability to regulate intracellular Ca2+ levels (see above), and thereby attenuate/prevent the activation of Ca2+-activated proteases (e.g., calpains) or lipases, and/or to a direct or indirect antioxidant effect. Although Tau is unlikely to be a direct antioxidant (4, 42), several studies have shown that Tau attenuates nonenzymatic reactive oxygen species (ROS)-induced lipid peroxidation (16, 26, 53, 56, 63, 74). This effect may be mediated by Tau binding to membrane phospholipids, limiting ROS attack (59). Tau's cytoprotective potential has led to the proposition that Tau supplementation may be beneficial in conditions involving increased susceptibility to muscle damage and oxidative stress such as aging and muscular dystrophy (14, 17). To date, no studies have investigated Tau's protective potential during and after severe and potentially damaging muscle contractions in vitro, and whether Tau can attenuate nonenzymatic ROS-induced lipid peroxidation.
Finally, it has been demonstrated in various cell lines and tissues that an increase in Tau levels results in a downregulation of TauT mRNA and protein expression (for reviews see Refs. 28, 64). To date, there have been no studies examining the effect of in vivo Tau supplementation on skeletal muscle TauT protein expression.
Therefore, in this study we examined the effect of raising endogenous Tau content by oral supplementation on rat skeletal muscle TauT protein expression, contractile properties, stability of muscle force during and after severe high-frequency stimulation, and extent of ROS-induced lipid peroxidation. We hypothesize that 2 wk of Tau supplementation would 1) significantly raise muscle Tau content, 2) decrease TauT protein expression, 3) increase twitch force, 4) enhance resistance to fatigue, 5) protect against contraction-induced muscle damage by attenuating calpain activation, and 6) attenuate ROS-induced lipid peroxidation.
MATERIALS AND METHODS
Animals and muscle dissection.
All procedures were submitted to, and approved by, the Victoria University Animal Experimentation Ethics Committee. A total of ninety-four 8-wk-old Sprague-Dawley rats were utilized in this study. Forty-eight rats were fed Tau (2.5% wt/vol) in drinking water ad libitum for 2 wk with an otherwise standard rat chow diet, while 46 rats were given normal drinking water and chow for 2 wk. After supplementation, EDL muscles were dissected under anesthesia (Nembutal; ∼85 mg/kg ip) in accordance with Victoria University Animal Ethics procedures. Although large muscle preparations are generally not recommended for analyzing in vitro contractile function (7), these were deliberately chosen because the use of relatively large whole muscles with repeated or continuous high-frequency electrical stimulation typically leads to significant hypoxia/anoxia (7) and muscle damage (1, 34), thus allowing us to examine any protective effect of Tau on muscle function during, and in recovery from, high-frequency tetanic stimulation. The proximal and distal tendons were isolated, tied with surgical silk, and then cut before disruption of the blood and nerve supply. The experimental muscle was then placed immediately in a Krebs-Henseleit solution containing (in mM) 118 NaCl, 1.18 MgSO4, 4.75 KCl, 1.0 Na2HPO4, 2.5 CaCl2, 24.0 NaHCO3, and 11.0 glucose, pH 7.4 and bubbled with 95% O2-5% CO2 (BOC Gases, Preston, Victoria, Australia) at 25°C. Muscles were mounted horizontally between two field-stimulating platinum plate electrodes by tying the proximal end of the muscle to a force transducer (Research Grade 60-2999, Harvard Apparatus, South Natick, MA) and the distal end to a micromanipulator to allow length adjustment.
Muscle stimulation.
Square wave pulses (0.2 ms) were produced by a stimulator (Grass S11 stimulator, Quincy, MA) and amplified (40V; CE-1000, Crown Instruments, Elkhart, IN) to ensure sufficient current to produce maximal isometric tetanic contractions. Forces were recorded with a PowerLab 4510 (ADIstruments, Castle Hill, NSW, Australia) running Chart v5.0.2 for Windows. For all muscles, optimal muscle length (Lo) was initially determined by eliciting twitch contractions and adjusting the muscle length until maximum twitch force was produced. Optimal fiber length (Lf) was determined with the previously established Lf-to-Lo ratio of 0.44 for the EDL muscle (9). Next, the force-frequency relationship was determined by stimulating for 500 ms at 10, 30, 50, 80, and 100 Hz. Between each stimulation, muscles were allowed to recover for 3 min. Maximum isometric force (Po) was determined from the greatest force produced during the force-frequency stimulation. Peak isometric twitch force (Pt) was then measured (mean of 3 twitches), after which muscles were subjected to one of two different repetitive stimulation protocols: 1) 10-s continuous stimulation at a frequency of 100 Hz (0.2-ms pulse duration; duty cycle 1.0) followed by a 1-h recovery period; or 2) 3-min intermittent stimulation (1-s stimulation at 100 Hz followed by 4-s recovery; duty cycle 0.2) followed by a 1-h recovery period. The stimulation protocols chosen, although not using physiological frequencies, represented models in which muscles were stimulated continuously and force rapidly declined but recovered relatively quickly (seconds to minutes; Refs. 2, 35) and a more prolonged stimulation protocol of repeated tetani causing a relatively larger depression in force that recovered slowly (minutes to hours; Ref. 10).
For both stimulation protocols, Po (500 ms, 100 Hz, 0.2-ms pulse duration) was monitored during the 1-h recovery at 1, 2, 5, 10, 20, 30, 45, and 60 min to assess recovery/stability of muscle function after stimulation. Muscles were cut from the force transducer either immediately after the stimulation bout (for ROS-induced F2-isoprostane analysis) or after 1 h of recovery (for calpain analysis), blotted dry on Whatman 1 (Maidstone, UK) filter paper, weighed, and freeze clamped with aluminum tongs precooled in liquid N2. Contralateral muscles were also dissected out, blotted dry, weighed, and clamped to serve as controls. All muscles were stored at −80°C until analysis. Muscle cross-sectional area was calculated with muscle mass, Lf, and the reported density for mammalian skeletal muscle (43). Maximal tetanic specific force was calculated as force per cross-sectional area.
Taurine content.
Two milligrams of the powdered muscle was added to 20% (wt/vol) of sulfosalicylic acid placed on ice. Samples were then vortexed for 5 s at a time for a total of 10 min and then centrifuged at 28,000 rpm for 2 min at 0°C. The supernatant was then collected and added to 0.4 M borate buffer and vortexed. After 5 min the samples were vortexed again and recentrifuged before the supernatant was collected and stored at −80°C for further analysis. HPLC was performed on a Phenomenex Hypersil C18 (150 × 4.6 mm, 3 μm) analytical column protected by a Phenomenex Security Guard cartridge (C18, 4 × 3) column guard at 30°C. Gradient composition and detector wavelengths were as described previously by Dunnett and Harris (18), as was derivatization reagent preparation. Samples were derivatized for 7 min before injection with a ratio of 300 μl of sample to 300 μl of reagent. The integrated peak area of a range of standards (r2 = 0.99) was compared with samples to determine the tissue concentration of taurine.
Tau transporter immunoblotting.
TauT protein content was measured in the same control muscles used for Tau content analysis above (n = 8). Frozen muscle was homogenized on ice in 10 volumes of buffer containing 50 mM Tris, 1 mM EDTA, 10% (vol/vol) glycerol, 1% (vol/vol) Triton X-100, 50 mM NaF, 5 mM Na4P2O7, 1 mM DTT, 1 mM PMSF, 10 μg/ml of trypsin inhibitor (Sigma, St. Louis, MO), and 5 μl/ml Protease Inhibitor Cocktail (P8340, Sigma) (pH 7.5). The resulting lysates were left on ice for 20 min and then spun at 10,000 g for 20 min at 4°C. Protein concentration was determined with a bicinchoninic acid (BCA) protein assay (Pierce, Rockford, IL) with BSA as the standard. Lysates were then boiled for 5 min in Laemmli sample buffer. Fifty micrograms of total protein was separated at room temperature on 7.5% acrylamide sodium dodecyl sulfate (SDS)-PAGE gels for 1.5 h at constant voltage (90 V for 30 min, then 60 min at 150 V) and then transferred to polyvinylidene difluoride (PVDF) membranes at 4°C [90 min at a constant 95 V; 37 mmol/l Tris, pH 8.3, 139 mmol/l glycine, and 20% (vol/vol) methanol]. Blots were probed with a rabbit anti-rat TauT polyclonal antibody (1:500; Alpha Diagnostic International, San Antonio, TX). Membranes were then reprobed with mouse anti-β-tubulin monoclonal antibody (1:10,000; Sigma). Binding was detected with IRDye 800-conjugated anti-rabbit IgG (1:5,000; Rockland, Gilbertsville, PA) or IRDye 680-conjugated anti-mouse IgG (1:5,000; Molecular Probes, Invitrogen) secondary antibodies, and protein bands were analyzed by infrared detection (Odyssey imaging system, LI-COR Biosciences, Lincoln, NE). To control for any differences in protein loading, TauT band intensity was expressed relative to tubulin band intensity from the same sample.
F2-isoprostane analysis.
F2-isoprostanes and arachidonic acid were measured as previously described (46, 47). Eight control and eight Tau-supplemented muscles were analyzed immediately after each of the two fatigue protocols described above (32 muscles in total). F2-isoprostanes were analyzed by electron-capture negative ionization GC-MS after solid-phase extraction and HPLC purification using [D4]-8-iso-prostaglandin F2α (Cayman Chemical) as internal standard (46). For arachidonic acid, phospholipids were separated by thin-layer chromatography, and the fatty acid methyl esters then prepared and analyzed by gas-liquid chromatography (GLC) (47). Muscle F2-isoprostane content was expressed relative to muscle arachidonic acid content (13).
Calpain autolysis, calsequestrin, and sarco(endo)plasmic reticulum Ca2+-ATPase immunoblotting.
μ-Calpain and calpain-3 autolysis was determined in eight control and seven Tau-supplemented nonstimulated and stimulated muscles collected after the 1-h recovery from the 3-min intermittent stimulation bout (30 muscles in total). Calsequestrin (CSQ)1 and sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA)1 were measured in the nonstimulated control and Tau-supplemented muscles. Muscle samples were homogenized in 10 volumes of ice-cold extraction buffer [0.4 M Tris-Cl, pH 6.8, and 25 mM EGTA ([Ca2+] < 10 nM)] and SDS added to a final concentration of 4%. Homogenates were incubated at 4°C for 20–40 min, and an aliquot was kept for a protein assay (Quant-iT fluorescence assay, Invitrogen, Sydney, Australia). A further dilution of the homogenate was made with extraction buffer (1:5 vol/vol), which was then added (2:1 vol/vol) to SDS loading buffer (0.125 M Tris·HCl, 10% glycerol, 4% SDS, 4 M urea, 10% mercaptoethanol, and 0.001% bromophenol blue, pH 6.8). Samples were heated to 95°C for 4 min and stored at −20°C until analysis. Samples were analyzed by Western blotting as previously described (50). Protein from the total muscle extracts (∼50 μg) was separated on an 8% SDS-PAGE gel and transferred to nitrocellulose membranes. Membranes were probed with antibodies against μ-calpain (1:1,000, mouse monoclonal, Sigma monoclonal, clone 15C10), calpain-3 (1:200, mouse monoclonal, Novocastra monoclonal 12A2, Newcastle, UK), actin (1:200, rabbit polyclonal, affinity isolated, A2066, Sigma), CSQ1 (1:2,000, mouse monoclonal VIIIDI2 clone, ab2824, Abcam, Cambridge, UK), or SERCA1 isoform (1:200, mouse monoclonal CaF2-5D2 clone, Developmental Studies Hybridoma Bank, University of Iowa), after which goat anti-mouse horseradish peroxidase (HRP) (1:50,000; Bio-Rad, Hercules, CA) was added to the membranes. Bands were visualized with West Pico chemiluminescent substrate (Pierce), and densitometry was performed with Quantity One software (Bio-Rad). After transfer, the gels were stained with BioSafe Coomassie blue (Bio-Rad), and myosin heavy chain (MHC) was used as an indicator of sample loaded. Full-length μ-calpain is visualized as an 80-kDa protein, and its activation is confirmed by its autolysis to 78- and 76-kDa proteins (5). Calpain-3 is observed as a 94-kDa protein that autolyzes to proteins of ∼60, 58, and 55 kDa when activated (65). In rodent muscle a nonspecific band at ∼82 kDa previously described with the 12A2 antibody (49) is likely to correspond to one of the ubiquitous calpains as the antibody detects purified rat m-calpain (Mollica, Murphy, and Lamb, unpublished observations) and possibly detects μ-calpain as previously suggested (3).
Statistical analyses.
All data are presented as means ± SE. All two-group comparisons between control and Tau-supplemented groups were analyzed with Student's unpaired t-test. Each individual stimulation frequency in the force-frequency analysis was analyzed with a Student's unpaired t-test. Between-group comparisons during the 1-h recovery time period were analyzed with a repeated-measures two-way ANOVA with Bonferroni's post test. To determine whether there were any differences in basal F2-isoprostane, a one-way ANOVA was used to compare the nonstimulated control and Tau-supplemented muscles serving as contralateral controls for muscles subjected to the 10-s continuous and 3-min intermittent stimulation protocols. Differences in muscle F2-isoprostanes between stimulated and nonstimulated muscle groups for each stimulation protocol (10 s or 3 min) were analyzed with a two-way ANOVA. Significance was set at P < 0.05. Statistical analyses were performed with GraphPad Prism (v 5.0) software.
RESULTS
Effect of Tau supplementation on body mass, muscle mass, Tau content, and TauT protein.
Tau supplementation (n = 48) had no effect on body mass (370 ± 4 vs. 362 ± 5 g) or EDL muscle mass (166 ± 4 vs. 170 ± 2 mg) compared with controls (n = 46), although there was a trend (P = 0.0544) for a higher (2.0%) muscle mass-to-body mass ratio in Tau-supplemented animals compared with controls (0.47 ± 0.010 vs. 0.45 ± 0.008, respectively). There was no difference (P = 0.57) in the dry weight-to-wet weight ratio between the two groups (data not shown). Muscle Tau content was measured in eight representative supplemented nonstimulated muscles and showed a 39.5% increase in muscle Tau content compared with controls (56.9 ± 3.3 vs. 40.8 ± 1.3 mmol/kg wet wt). To determine whether this large increase in muscle Tau content would lead to a downregulation of the TauT, TauT protein was examined by Western blot in the same muscles in which Tau content was measured and expressed relative to a loading control protein, tubulin. Figure 1A shows a representative Western blot of TauT and tubulin from control muscles (lanes 1–4) and Tau-supplemented muscles (lanes 5–8). Despite the large increase in muscle Tau content with supplementation, there was no change in the TauT protein content over the supplementation period (Fig. 1B).
Fig. 1.
Effect of 2-wk taurine (Tau) supplementation on muscle Tau transporter (TauT) content. A: representative Western blot of TauT protein (top) with tubulin (bottom) as a loading control. Lanes 1–4 are control muscles, and lanes 5–8 are Tau-supplemented muscles. B: TauT content expressed relative to a loading control protein, tubulin. There was no significant difference between control (n = 8) and Tau-supplemented (n = 8) muscles (P = 0.41; means ± SE).
Effect of Tau supplementation on twitch and tetanic contractile properties.
Figure 2A shows the mean twitch response for control (n = 46) and Tau-supplemented (n = 48) EDL muscles. Table 1 shows the quantified mean contractile properties. Tau supplementation resulted in 19.0% greater peak twitch force and a 13.5% longer time to peak force, with no difference in the rate of force development between 20% and 80% of peak force. A trend was also found for a longer half-relaxation time (17.5%; P = 0.05) in muscle from Tau-supplemented animals, with no difference in the rate of force decline between 80% and 50% of peak force (Table 1). To examine the effect of elevated Tau on muscle force at higher stimulation frequencies, a force-frequency analysis was performed. As shown in Fig. 2B, muscles supplemented with Tau (n = 48) developed significantly greater relative forces at 10 (7.0%)-, 30 (22.2%)-, 50 (7.8%)-, and 80 (2.1%)-Hz stimulation than controls (n = 46). Fitting a sigmoidal curve to this data showed that Tau supplementation resulted in a leftward shift in the force-frequency curve compared with controls, such that the stimulation frequency required to elicit 50% of maximal force was significantly (P < 0.001; unpaired t-test) lower than in controls (data not shown). Absolute maximum tetanic force (100 Hz) also showed a strong trend to be elevated (3.8%; P = 0.065), while maximal tetanic specific force (force per cross-sectional area) was elevated by 4.2% (P = 0.031) in Tau-supplemented muscles compared with controls (Table 1).
Fig. 2.
Effect of Tau supplementation on the mean twitch response and force-frequency relationship. A: mean twitch response. Error bars have been removed for clarity (n = 46 for controls and n = 48 for Tau supplemented). B: muscles were stimulated for 500 ms (0.2-ms pulse duration) at 10, 30, 50, 80, and 100 Hz. Isometric forces were normalized to the force generated at 100 Hz (n = 46 for controls and n = 48 for Tau supplemented; means ± SE).
Table 1.
Twitch and tetanic contractile properties of EDL muscles from control and Tau-supplemented animals
Peak Force, mN | Time to Peak, ms | Rate of Force Development (20-80%), mN/s | Half-Relaxation Time, ms | Rate of Force Decline (80-50%), mN/s | Tetanic Force, mN | Specific Force, kN/m2 | |
---|---|---|---|---|---|---|---|
Con (n = 46) | 1,027±29 | 16.3±0.4 | 77,340±2,308 | 19.9±0.8 | 30,685±1,004 | 5,797±104 | 401±6 |
Tau (n = 48) | 1,222±22* | 18.5±0.3* | 80,550±1,565 | 23.4±1.5† | 31,472±632 | 6,018±59† | 418±5* |
*P < 0.0001 | *P < 0.0001 | †P = 0.05 | †P = 0.065 | *P = 0.031 |
Values are means ± SE for n rats. EDL, extensor digitorum longus; Tau, taurine; Con, control.
Significantly different from Con.
Effect of Tau supplementation on muscle calsequestrin and SERCA1 protein content.
To investigate whether Tau supplementation may have induced changes in the content of Ca2+ regulatory proteins, we examined the content of the SR Ca2+ binding protein CSQ1 and the SR Ca2+-ATPase SERCA1. As shown in Fig. 3, Tau supplementation resulted in a 49% increase in CSQ1. This difference was significant whether CSQ1 in Tau-supplemented muscles was expressed relative to control muscle values [1.49 ± 0.18 vs. 1.00 ± 0.15 arbitrary units (au); P = 0.036, Fig. 3B] or CSQ1 in both Tau and control samples was expressed relative to the loading control protein actin on each lane (2.99 ± 0.35 vs. 2.00 ± 0.30 au; P = 0.036). Tau supplementation did not, however, result in a change (P = 0.39) in the protein content of SERCA1 compared with controls (Fig. 3C) or normalized to actin.
Fig. 3.
Effect of Tau supplementation on calsequestrin (CSQ)1 and sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA)1 protein content. A: representative Western blots of SERCA1 (top), CSQ1 (middle), and loading control actin (bottom). B: CSQ1 protein expression relative to control samples. *Significantly different from Tau-supplemented muscles (P = 0.036; n = 8 for controls and n = 7 for Tau). C: SERCA1 protein expression relative to control samples (P = 0.39; n = 8 for controls and n = 7 for Tau). For both B and C, results are the same when CSQ1 and SERCA1 are expressed relative to the loading control protein actin (means ± SE).
Effect of Tau supplementation on force production during fatiguing protocols.
As shown in Fig. 4A, muscles from Tau-supplemented rats were significantly (P = 0.026) more resistant to fatigue when subjected to 10-s continuous stimulation, with force being reduced by 60.5 ± 1.3% (n = 24) compared with 66.0 ± 2.1% (n = 21) in control muscles. Similarly, as shown in Fig. 4B, when subjected to 3-min intermittent stimulation muscles from Tau-supplemented rats were again more resistant to fatigue (P = 0.009), with force being reduced by 89.9 ± 0.6% (n = 23) compared with 92.4 ± 0.6% (n = 20) in control muscles. In addition, the area under the smoothed fatigue curves was 12.5% (P = 0.0014) greater for muscles from Tau-supplemented rats subjected to 3-min intermittent stimulation compared with controls (data not shown).
Fig. 4.
Effect of Tau supplementation on resistance to fatigue. A: 10-s continuous stimulation (100 Hz, 0.2-ms pulse duration, duty cycle 1.0). *Significantly different from control (P = 0.026; n = 22 for control and n = 20 for Tau supplemented). Error bars have been removed for clarity. B: 3-min intermittent stimulation (100 Hz, 0.2-ms pulse duration for 1 s, then 4-s rest, duty cycle 0.2). *Significantly different from control (P = 0.009; n = 20 for control and n = 23 for Tau supplemented; means ± SE).
Effect of increased muscle Tau content on muscle F2-isoprostane production during fatiguing contractions.
Importantly, there was no difference in basal levels of F2-isoprostanes between any of the four nonstimulated contralateral groups (P = 0.12, 1-way ANOVA; Fig. 5). As shown in Fig. 5A, there was no effect of stimulation on F2-isoprostane levels immediately after 10 s of continuous fatiguing tetanic stimulation in either control (n = 8) or Tau-supplemented (n = 8) muscles. Conversely, after 3 min of intermittent stimulation there was a significant main effect (P = 0.0003, 2-way ANOVA) for F2-isoprostane levels to be increased by stimulation (Fig. 5B), mostly due to the large increase in the control stimulated muscles, and a strong trend for an interaction effect (P = 0.062). The fact that, despite the clear difference in F2-isoprostane production between control and Tau muscles subjected to the 3-min stimulation protocol, the interaction effect just failed to reach significance most likely reflects inadequate statistical power. To highlight this, when the differences in F2-isoprostanes between nonstimulated and stimulated control and Tau muscles were analyzed with unpaired t-tests there was a significant increase (P = 0.0008) of 46.7% in the Tau muscles and no significant change in the control muscles (P = 0.1460). This was further confirmed by using the false discovery rate (FDR) procedure (for review see Ref. 15), which rejected the null hypothesis that there was no difference in the level of F2-isoprostane levels between the control nonstimulated and stimulated muscles, indicating that the ∼47% increase was indeed significant in stimulated control muscles. Conversely, the FDR procedure accepted the null hypothesis that there was no difference between Tau-supplemented nonstimulated and stimulated muscles, indicating that there was no significant change in F2-isoprostane levels in stimulated Tau muscles. Overall, these results indicate that raised muscle Tau content attenuated F2-isoprostane production during prolonged intermittent tetanic stimulation.
Fig. 5.
F2-isoprostane production during fatiguing in vitro contractions. A: 10-s continuous stimulation. B: 3-min intermittent stimulation. #Significant main effect for stimulation (P = 0.0003; 2-way ANOVA); *nonstimulated control different from stimulated control (P = 0.0008; unpaired t-test combined with false discovery rate procedure) (means ± SE). For all groups n = 8.
Effect of Tau supplementation on recovery of muscle tetanic force after severe electrical stimulation.
Severe fatiguing stimulation of relatively large muscles is likely to create a significant hypoxic/anoxic core that will lead to a decline in contractile function due to muscle damage (1). To examine whether an increase in muscle Tau content would confer a degree of protection against this stimulation-induced decline in muscle function, tetanic force was monitored over a 1-h period after each of the two stimulation protocols in muscles from control and Tau-supplemented muscles. As shown in Fig. 6A, characteristic of high-frequency fatigue, after 10 s of continuous tetanic stimulation force produced by control (n = 14) and Tau (n = 17) muscles returned rapidly to ∼80% of initial force. In control muscles, force remained stable over the next 20 min and then began to decline such that the force at 30, 45, and 60 min was significantly lower than at 1 min. At 60 min of recovery control muscle tetanic force had declined to ∼56% of prefatigue force. In muscles from Tau-supplemented animals, however, recovery force remained stable for 45 min after the end of stimulation, with only the 60-min force response slightly lower than at 1 min of recovery. At 60 min of recovery, Tau-supplemented muscle tetanic force was ∼71% of prefatigue force. Compared between the two groups, recovery tetanic force was significantly different for all time points between 20 and 60 min (Fig. 6A).
Fig. 6.
Recovery of tetanic force after fatiguing stimulation. A: tetanic force during 1-h recovery period after 10-s continuous 100-Hz stimulation in muscles from animals supplemented with Tau (n = 17) and controls (n = 14). *P < 0.01, #P < 0.001 significantly different from control value at the same time point. B: tetanic force during 1-h recovery period after 3-min intermittent 100-Hz stimulation in muscles from animals supplemented with Tau (n = 15) and controls (n = 15). ^P < 0.05, *P < 0.01, #P < 0.001 significantly different from control value at the same time point (means ± SE).
After the 3-min intermittent stimulation bout, muscle force showed a minimal increase in recovery within the first 1 min of recovery (Fig. 6B). In control muscles (n = 15), tetanic force did not increase until 20 min into the recovery period, after which force slowly increased to only ∼29% of prefatigue force by 45 min and then began to decline by 60 min (Fig. 6B). In contrast, tetanic force produced by muscles from Tau-supplemented animals (n = 15) began to increase earlier (at 10 min) and continued to increase up to 60 min with no sign of deterioration. Tetanic force in Tau muscles at 60 min of recovery was ∼47% of prefatigue force. Compared between the two groups, recovery tetanic force was significantly different for all time points between 5 and 60 min (Fig. 6B).
Calpain activation in response to severe damaging isometric contractions.
To investigate a possible mechanism that may explain why recovery tetanic force was lower with signs of deterioration in control muscles subjected to the 3-min intermittent stimulation protocol (Fig. 6B), calpain-1 (or μ-calpain) and calpain-3 autolysis (an indicator of calpain activity; Refs. 5, 24, 65) was investigated by Western blot analysis. As shown in Fig. 7, top and middle (lanes 3 and 4 in each), 1 h after the 3-min intermittent stimulation bout, when control muscle force had plateaued and began to decline to only ∼29% of prefatigue force, there was no sign of autolysis of either μ-calpain or calpain-3 in muscles from either Tau-supplemented animals (n = 7, lanes 1 and 2) or control animals (n = 8; e.g., lanes 3 and 4).
Fig. 7.
μ-Calpain and calpain-3 in muscles 1 h after severe fatiguing stimulation. Western blots show μ-calpain (top) and calpain-3 (reprobe, middle) in muscle samples either stimulated (S) or not stimulated (N-S) in control and Tau-supplemented muscles after 3 min of intermittent 100-Hz stimulation. Actin loading control also indicated (bottom). Molecular mass markers are indicated on left. The ∼82-kDa band is likely to correspond to one of the ubiquitous calpains (see materials and methods).
DISCUSSION
This study has produced several novel and important findings. Consistent with our hypotheses, raising muscle Tau content via supplementation resulted in 1) an increase in peak twitch, subtetanic, and maximal specific force, 2) enhanced force production during and 3) in recovery from severe high-frequency stimulation, and 4) reduced nonenzymatic ROS-induced lipid peroxidation during repeated tetanic contractions. Contrary to our hypotheses, however, increased muscle Tau did not result in a downregulation of TauT protein expression. Furthermore, we have shown that 2 wk of Tau supplementation led to an increase in the SR Ca2+ binding protein CSQ1. Finally, we showed for the first time that long-lasting reduced, and deteriorating, muscle force induced by severe in vitro isometric contractions was not associated with the activation (autolysis) of μ-calpain or calpain-3.
Effect of Tau supplementation on muscle Tau content.
The absolute muscle Tau concentration in this study for nonsupplemented rats (41 mmol/kg wet wt) was higher than those published previously for rat EDL muscle. Interestingly, however, the values previously published for EDL muscle Tau vary tremendously, for example, ∼1.0 (39), ∼3.0 (57), 14 (16), and 17 (32) mmol/kg wet wt. The differences among all these studies, including ours, are likely due to differences in animal strain, age, diet, and the methods used for Tau extraction and analysis (HPLC vs. automated amino acid analyzer). Regardless of the absolute Tau content, the important aspect of this study is that when all samples were analyzed at the same time and under the same conditions we showed that Tau supplementation increased muscle Tau by ∼40% above endogenous levels.
Effect of Tau supplementation on Tau transporter protein expression.
Our study is the first to examine the effect of Tau supplementation on skeletal muscle TauT protein expression with no evidence of a decrease in TauT protein expression over the 2-wk period, despite ∼40% increase in Tau content. Previous studies have shown that exposure of various cell lines to Tau results in a downregulation of TauT mRNA and protein levels (64). In rats in vivo a high-Tau diet (3% wt/vol) has been shown to decrease kidney TauT mRNA and reduce Tau transport Vmax, indicative of reduced protein expression (38). Further research is required to examine the effect of more prolonged Tau supplementation on TauT regulation in skeletal muscle.
Effect of Tau supplementation on twitch and tetanic contractile properties.
Bakker and Berg (6) previously showed in rat mechanically skinned EDL fibers bathed in aqueous solutions that the reintroduction of approximately endogenous levels of Tau resulted in ∼20% greater depolarization-induced isometric force. In the present study, we show that raising Tau content above endogenous levels in whole EDL muscles also resulted in ∼20% increase in peak twitch force. This increase in twitch force could be due to an increased sensitivity of the contractile apparatus to Ca2+ with increased endogenous Tau leading to a decrease in intracellular ionic strength due to increased water influx similar to that observed with elevated creatine in skinned fibers (48). Interestingly, however, we did not detect any difference between control and Tau-supplemented muscles in wet weight-to-dry weight ratio, a crude indicator of tissue water content. Another possibility is that increased Tau may lead to increased SR Ca2+ accumulation (6, 30, 58) and/or increased SR Ca2+ release by increasing the activity of the SR ryanodine receptor (RyR)/Ca2+ release channel (6), leading to an increase in twitch force. Our novel finding that Tau supplementation increased the content of the SR Ca2+ binding protein CSQ1, and therefore the total capacity to store Ca2+ (51), is consistent with the first of these possibilities. Although there are currently no data on the effect of increasing CSQ1 on whole skeletal muscle Ca2+ kinetics and contractile properties, Shin et al. (62) reported that overexpression of CSQ1 in C2C12 myotubes resulted in increased caffeine- and voltage-induced SR Ca+ release due to greater SR Ca2+ load, while CSQ1-null mice display reduced SR Ca2+ release and reduced Ca2+ transient amplitude (55). In regard to function of the RyR/Ca2+ release channel, Baker and Berg (6) showed that acute Tau application to mechanically skinned fibers increased caffeine-induced force responses and proposed that Tau potentiated SR Ca2+ release, presumably by increasing RyR/Ca2+ release channel activity, while changes in SR CSQ1 content also have the potential to alter RyR/Ca2+ release channel function (8). Thus Tau-induced alterations in CSQ1 content and increase in Ca2+ release may help explain our observed increase in twitch force with Tau supplementation. Further work is required to help clarify these mechanisms.
In the present study, we found the time to peak twitch force was longer, with no change in the rate of force development and a trend (P = 0.05) for a slight increase in twitch half-relaxation time in muscles from Tau-supplemented animals. The longer twitch half-relaxation time in our whole muscles is likely due to the higher peak force reached therefore taking longer for force to decline to 50%. This result suggests that raising Tau above endogenous levels does not further increase the maximal rate of SR Ca2+ accumulation during twitch contractions. This is consistent with our finding of no change in the abundance of SERCA1 protein with Tau supplementation.
As well as increasing twitch force, Tau supplementation induced an upward shift in the force-frequency relationship, with higher forces produced between 10- and 80-Hz stimulation, likely due to a combination of the higher peak forces and the slight slowing of the half-relaxation time. Hamilton et al. (27) showed the opposite effect in mouse EDL muscle depleted of endogenous Tau with the competitive inhibitor of Tau uptake, GES, confirming an important role for Tau in regulating skeletal muscle contractile properties. In addition, our results also show that Tau supplementation resulted in a leftward shift in force-frequency curve, consistent with the notion of increased Ca2+ sensitivity of the contractile apparatus in the presence of increased muscle Tau.
Effect of Tau supplementation on force production during continuous and intermittent high-frequency stimulation.
To determine whether elevated Tau would protect muscle function, we deliberately subjected relatively large muscles to severe contraction protocols that would produce significant hypoxia/anoxia (7) and lead to muscle damage (34). In the present study we showed that raised muscle Tau led to ∼6% more force production at the end of the 10-s stimulation bout, an effect that may be due to the greater sensitivity of the contractile apparatus to Ca2+ but could also be related to Tau's demonstrated ability to inhibit KATP channels (68), delay K+ leak through Ca2+-activated K+ (KCa) channels due to either a direct action of Tau (69) or better intracellular Ca2+ handling (6, 30), and thus delay membrane potential rundown (19, 25). It does not seem that increased ROS was a factor in this brief stimulation protocol because we did not detect any increase in F2-isoprostane levels (an indicator of nonenzymatic ROS-induced lipid peroxidation and a sensitive measure of oxidative stress; Ref. 44) after fatigue in either control or Tau-supplemented muscles.
Repeated tetani for 3 min resulted in a large reduction in force (∼90%) and a very slow recovery over the next 1 h. Although by the end of the 3-min stimulation force was only ∼2.5% higher in Tau-supplemented muscle compared with controls, the area under the force curve was 12% greater in Tau-supplemented muscles, largely because of the greater force production during the middle portion of the curve (see Fig. 3B). In single-fiber experiments, this region of the force curve is much flatter but can be made steeper by the addition of cyanide and the abolition of oxidative metabolism (72, 75). The effect of Tau on this portion of the curve suggests that Tau may have protected mitochondrial function and thus helped maintain ATP-dependent processes within the cell (60). Although we did not measure mitochondrial function or ATP, several studies have demonstrated that Tau plays an important role in mitochondrial function (12, 20, 29, 54, 56, 61, 69, 74). In this context, it is also interesting to note that Tau levels are much higher in mitochondrially rich slow-twitch muscles (e.g., soleus) than glycolytic fast-twitch muscles (e.g., EDL; Ref. 32).
Our results show for the first time in an isolated in vitro system, free from other potential sources of contamination and using the highly sensitive and specific gas chromatography/electron capture negative ionization mass spectrometry method (45, 46), that repeated tetanic contractions lead to significant skeletal muscle oxidative stress with a ∼47% increase in F2-isoprostane content. Importantly, raised muscle Tau attenuated this contraction-induced lipid peroxidation, in agreement with other studies using less sensitive measures in whole exercising animals (16), which suggests a role for Tau in protecting membranes from direct ROS attack and/or reducing overall ROS production. Further work is required to elucidate the exact mechanism behind Tau's protective effect.
Effect of Tau supplementation on stability/recovery of muscle force after severe tetanic contractions.
Many years ago it was shown that isometric contractions of whole skeletal muscles can result in irreversible force loss, and intracellular enzyme release (34) that is exacerbated by hypoxia/anoxia (21, 40). This effect could be mimicked by the use of mitochondrial inhibitors and alleviated by the removal of extracellular Ca2+ (33). These findings suggest that contractions under hypoxic/anoxic conditions lead to disruption of the sarcolemmal membrane resulting in an influx of extracellular Ca2+ and Ca2+-induced muscle damage and that this process may involve ATP depletion via hypoxia/anoxia-mediated inhibition of oxidative phosphorylation. Our results clearly show that raised muscle Tau had a protective effect on skeletal muscle function after fatiguing/damaging contractions. These results are supported by a recent results from Yatabe et al. (73) showing reduced urinary markers of muscle damage after exhaustive running following 2 wk of Tau supplementation. Moreover, TauT−/− mice, with severely depleted muscle Tau content, display resting plasma creatine kinase levels (an indicator of muscle plasma membrane damage) double those of wild types (71). Studies have also shown that raising endogenous Tau levels confers protection against skeletal muscle ischemia-reperfusion injury (37, 41, 70). Given that ischemia-reperfusion injury is thought to be largely caused by increased intracellular Ca2+ and increased mitochondrial ROS production (52), this suggests that Tau's protective effect is related to its ability to regulate intracellular Ca2+ and/or to its indirect antioxidant properties, and is consistent with Tau's effect of reducing lipid peroxidation (see above) by either reducing ROS production or helping to maintain lipid membrane integrity.
Calpain activation after severe repeated tetanic contractions.
It has also been speculated that prolonged force depression after fatiguing repeated tetani could be due to activation of Ca2+-dependent cysteine proteases (calpains) and disruption of critical SR, T tubule, contractile, and/or cytoskeletal proteins (2, 23). Studies using calpain inhibitors (e.g., leupeptin, which inhibits μ-calpain and m-calpain but not calpain-3) have failed to convincingly demonstrate a role for either μ-calpain or m-calpain in this phenomenon (2, 10). After 1-h recovery following the 3-min intermittent stimulation protocol, we found no autolysis of either μ-calpain or calpain-3 despite the low and deteriorating loss of tetanic force in control muscles, and therefore could not attribute Tau's protective effect to a direct effect on calpain activation. These findings suggest that any increase in intracellular [Ca2+] during the prolonged in vitro stimulation was neither high enough nor for a long enough period of time to activate either μ-calpain or calpain-3, similar to the observation of Murphy et al. (49) with high-intensity cycling or endurance running. They also reinforce other studies using protease inhibitors showing that μ-calpain activation during or immediately after exercise is unlikely to play a role in the prolonged reduction of force production after repeated isometric tetani and suggest that other proteolytic and/or lipolytic processes are involved in the deterioration of muscle force after severe tetanic contractions under hypoxic/anoxic conditions.
Our results, combined with recent studies that reintroduced endogenous levels of Tau (6) or depleted muscle Tau (27, 71), confirm that Tau plays an important role in modulating skeletal muscle function. Our findings of increased force production and enhanced fatigue resistance with tetanic stimulation warrant further investigation into possible ergogenic effects of high-dose Tau supplementation during high-intensity exercise in humans. In this context, it remains to be determined whether muscle Tau can be elevated to the same extent in humans as in rodent muscle, and what dosage would be required to achieve this. Although considered nontoxic in healthy humans, very high doses could result in gastrointestinal distress. In a recent human study by Galloway et al. (22), muscle Tau failed to increase after 7 days of oral supplementation (∼5 g/day), leading the authors to speculate that this may have been due to a downregulation of TauT protein. In light of our results, notwithstanding any species differences, the reason for the lack of increase in muscle Tau in that study may be the Tau dosage being too low and, in particular, the relatively short duration, and not changes in TauT protein. Dose-response studies are, however, required to examine this issue further.
Our study also provides further evidence that elevated Tau plays a protective role in skeletal muscle (14, 36), possibly by protecting lipid membrane structures from direct oxidant attack and/or by reducing overall ROS production per se during repeated contractions. Our finding of increased CSQ1 with Tau supplementation also reenforces the previous studies demonstrating a role for Tau in modulation of intracellular Ca2+ handling (14). These effects may aid recovery from damaging muscular activity. It has also been proposed that Tau supplementation may be useful in conditions that involve reduced force production and increased susceptibility to muscle damage and oxidative stress such as aging and muscular dystrophy (14, 17).
Conclusions.
In conclusion, 2 wk of Tau supplementation increased rat fast-twitch muscle Tau content without changing TauT protein content. This increase in muscle Tau resulted in increased in vitro isometric force production and enhanced fatigue resistance and protected muscle function during recovery. Our results point to a potential ergogenic effect of raising muscle Tau content with oral supplementation. Further work is required to investigate the mechanism of Tau's action and whether similar results can be obtained in healthy and diseased human populations.
Acknowledgments
The SERCA1 monoclonal antibody developed by D. M. Fambrough was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the National Institute of Child Health and Human Development and maintained by the Department of Biological Sciences, University of Iowa (Iowa City, IA).
REFERENCES
- 1.Allen DG, Whitehead NP, Yeung EW. Mechanisms of stretch-induced muscle damage in normal and dystrophic muscle: role of ionic changes. J Physiol 567: 723–735, 2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2.Allen DG, Lamb GD, Westerblad H. Skeletal muscle fatigue: cellular mechanisms. Physiol Rev 88: 287–332, 2008. [DOI] [PubMed] [Google Scholar]
- 3.Anderson LV, Davison K, Moss JA, Richard I, Fardeau M, Tomé FM, Hübner C, Lasa A, Colomer J, Beckmann JS. Characterization of monoclonal antibodies to calpain 3 and protein expression in muscle from patients with limb-girdle muscular dystrophy type 2A. Am J Pathol 153:1169–1179, 1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4.Aruoma OI, Halliwell B, Hoey BM, Butler J. The antioxidant action of taurine, hypotaurine and their metabolic precursors. Biochem J 256: 251–255, 1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 5.Baki A, Tompa P, Alexa A, Molnar O, Friedrich P. Autolysis parallels activation of mu-calpain. Biochem J 318: 897–901, 1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6.Bakker AJ, Berg HM. Effect of taurine on sarcoplasmic reticulum function and force in skinned fast-twitch skeletal muscle fibres of the rat. J Physiol 538: 185–194, 2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7.Barclay CJ Modeling diffusive O2 supply to isolated preparations of mammalian skeletal and cardiac muscle. J Muscle Res Cell Motil 26: 225–235, 2005. [DOI] [PubMed] [Google Scholar]
- 8.Beard NA, Laver DR, Dulhunty AF. Calsequestrin and the calcium release channel of skeletal and cardiac muscle. Prog Biophys Mol Biol 85: 33–69, 2004. [DOI] [PubMed] [Google Scholar]
- 9.Brooks SV, Faulkner JA. Contractile properties of skeletal muscles from young, adult and aged mice. J Physiol 404: 71–82, 1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 10.Bruton JD, Lannergren J, Westerblad H. Mechanisms underlying the slow recovery of force after fatigue: importance of intracellular calcium. Acta Physiol Scand 162: 285–293, 1998. [DOI] [PubMed] [Google Scholar]
- 11.Canas PE The role of taurine and its derivatives on cellular hypoxia: a physiological view. Acta Physiol Pharmacol Ther Latinoam 42: 133–137, 1992. [PubMed] [Google Scholar]
- 12.Chang L, Xu J, Yu F, Zhao J, Tang X, Tang C. Taurine protected myocardial mitochondria injury induced by hyperhomocysteinemia in rats. Amino Acids 27: 37–48, 2004. [DOI] [PubMed] [Google Scholar]
- 13.Choy K, Beck K, Png FY, Wu BJ, Leichtweiss SB, Thomas SR, Hou JY, Croft KD, Mori TA, Stocker R. Processes involved in the site-specific effect of probucol on atherosclerosis in apolipoprotein E gene knockout mice. Arterioscler Thromb Vasc Biol 25: 1684–1690, 2005. [DOI] [PubMed] [Google Scholar]
- 14.Conte Camerino D, Tricarico D, Pierno S, Desaphy JF, Liantonio A, Pusch M, Burdi R, Camerino C, Fraysse B, De Luca A. Taurine and skeletal muscle disorders. Neurochem Res 29: 135–142, 2004. [DOI] [PubMed] [Google Scholar]
- 15.Curran-Everett D Multiple comparisons: philosophies and illustrations. Am J Physiol Regul Integr Comp Physiol 279: R1–R8, 2000. [DOI] [PubMed] [Google Scholar]
- 16.Dawson R, Biasetti M, Messina S, Dominy J. The cytoprotective role of taurine in exercise-induced muscle injury. Amino Acids 22: 309–324, 2002. [DOI] [PubMed] [Google Scholar]
- 17.De Luca A, Pierno S, Liantonio A, Conte Camerino D. Pre-clinical trials in Duchenne dystrophy: what animal models can tell us about potential drug effectiveness. Neuromuscul Disord 12, Suppl 1: S142–S146, 2002. [DOI] [PubMed] [Google Scholar]
- 18.Dunnett M, Harris RC. High-performance liquid chromatographic determination of imidazole dipeptides, histidine, 1-methylhistidine and 3-methylhistidine in equine and camel muscle and individual muscle fibres. J Chromatogr B Biomed Sci Appl 688: 47–55, 1997. [DOI] [PubMed] [Google Scholar]
- 19.Duty S, Allen DG. The effects of glibenclamide on tetanic force and intracellular calcium in normal and fatigued mouse skeletal muscle. Exp Physiol 80: 529–541, 1995. [DOI] [PubMed] [Google Scholar]
- 20.El Idrissi A Taurine increases mitochondrial buffering of calcium: role in neuroprotection. Amino Acids 34: 321–328, 2008. [DOI] [PubMed] [Google Scholar]
- 21.Fredsted A, Mikkelsen UR, Gissel H, Clausen T. Anoxia induces Ca2+ influx and loss of cell membrane integrity in rat extensor digitorum longus muscle. Exp Physiol 90: 703–714, 2005. [DOI] [PubMed] [Google Scholar]
- 22.Galloway SDR, Talanian JL, Shoveller AK, Heigenhauser GJF, Spriet LL. Seven days of oral taurine supplementation does not increase muscle taurine content or alter substrate metabolism during prolonged exercise in humans. J Appl Physiol 105: 643–651, 2008. [DOI] [PubMed] [Google Scholar]
- 23.Gissel H The role of Ca2+ in muscle cell damage. Ann NY Acad Sci 1066: 166–180, 2005. [DOI] [PubMed] [Google Scholar]
- 24.Goll DE, Thompson VF, Li H, Wei W, Cong J. The calpain system. Physiol Rev 83: 731–801. [DOI] [PubMed]
- 25.Gong B, Legault D, Miki T, Seino S, Renaud JM. KATP channels depress force by reducing action potential amplitude in mouse EDL and soleus muscle. Am J Physiol Cell Physiol 285: C1467–C1474, 2003. [DOI] [PubMed] [Google Scholar]
- 26.Haber CA, Lam TKT, Yu Z, Gupta N, Goh T, Bogdanovic E, Giacca A, Fantus IG. N-acetylcysteine and taurine prevent hyperglycemia-induced insulin resistance in vivo: possible role of oxidative stress. Am J Physiol Endocrinol Metab 285: E744–E753, 2003. [DOI] [PubMed] [Google Scholar]
- 27.Hamilton EJ, Berg HM, Easton CJ, Bakker AJ. The effect of taurine depletion on the contractile properties and fatigue in fast-twitch skeletal muscle of the mouse. Amino Acids 31: 273–278, 2006. [DOI] [PubMed] [Google Scholar]
- 28.Han X, Patters AB, Jones DP, Zelikovic I, Chesney RW. The taurine transporter: mechanisms of regulation. Acta Physiol (Oxf) 187: 61–76, 2006. [DOI] [PubMed] [Google Scholar]
- 29.Hansen SH, Andersen ML, Birkedal H, Cornett C, Wibrand F. The important role of taurine in oxidative metabolism. Adv Exp Med Biol 583: 129–35, 2006. [DOI] [PubMed] [Google Scholar]
- 30.Huxtable R, Bressler R. Effect of taurine on a muscle intracellular membrane. Biochim Biophys Acta 323: 573–583, 1973. [DOI] [PubMed] [Google Scholar]
- 31.Huxtable JR Physiological actions of taurine. Physiol Rev 72: 101–163, 1992. [DOI] [PubMed] [Google Scholar]
- 32.Iwata H, Obara T, Kim BK, Baba A. Regulation of taurine transport in rat skeletal muscle. J Neurochem 47: 158–163, 1986. [DOI] [PubMed] [Google Scholar]
- 33.Jackson MJ, Jones DA, Edwards RH. Experimental skeletal muscle damage: the nature of the calcium-activated degenerative processes. Eur J Clin Invest 14: 369–374, 1984. [DOI] [PubMed] [Google Scholar]
- 34.Jones DA, Jackson MJ, Edwards RH. Release of intracellular enzymes from an isolated mammalian skeletal muscle preparation. Clin Sci (Lond) 65: 196–201, 1983. [DOI] [PubMed] [Google Scholar]
- 35.Jones DA High- and low-frequency fatigue revisited. Acta Physiol Scand 156: 265–270, 1996. [DOI] [PubMed] [Google Scholar]
- 36.Kingston R, Kelly CJ, Murray P. The therapeutic role of taurine in ischaemia-reperfusion injury. Curr Pharm Des 10: 2401–2410, 2004. [DOI] [PubMed] [Google Scholar]
- 37.Kingston R, Kearns S, Kelly C, Murray P. Effects of systemic and regional taurine on skeletal muscle function following ischaemia-reperfusion injury. J Orthop Res 23: 310–314, 2005. [DOI] [PubMed] [Google Scholar]
- 38.Matsell DG, Bennett T, Han X, Budreau AM, Chesney RW. Regulation of the taurine transporter gene in the S3 segment of the proximal tubule. Kidney Int 52: 748–754, 1997. [DOI] [PubMed] [Google Scholar]
- 39.Matsuzaki Y, Miyazaki T, Miyakawa S, Bouscarel B, Ikegami T, Tanaka N. Decreased taurine concentration in skeletal muscles after exercise for various durations. Med Sci Sports Exerc 34: 793–797, 2002. [DOI] [PubMed] [Google Scholar]
- 40.McCall KE, Duncan CJ. Independent pathways causing cellular damage in mouse soleus muscle under hypoxia. Comp Biochem Physiol A 94: 799–804, 1989. [DOI] [PubMed] [Google Scholar]
- 41.McLaughlin R, Bowler D, Kelly CJ, Kay E, Bouchier-Hayes D. Taurine protects against early and late skeletal muscle dysfunction secondary to ischaemia reperfusion injury. Eur J Surg 166: 375–379, 2000. [DOI] [PubMed] [Google Scholar]
- 42.Mehta TR, Dawson R. Taurine is a weak scavenger of peroxynitrite and does not attenuate sodium nitroprusside toxicity to cells in culture. Amino Acids 20: 419–433, 2001. [DOI] [PubMed] [Google Scholar]
- 43.Mendez J, Keys A. Density and composition of mammalian skeletal muscle. Metab Clin Exp 9: 184–199, 1960. [Google Scholar]
- 44.Milne GL, Musiek ES, Morrow JD. F2-isoprostanes as markers of oxidative stress in vivo: an overview. Biomarkers 10: 10–23, 2005. [DOI] [PubMed] [Google Scholar]
- 45.Milne GL, Sanchez SC, Musiek ES, Morrow JD. Quantification of F2-isoprostanes as a biomarker of oxidative stress. Nat Protoc 2: 221–226, 2007. [DOI] [PubMed] [Google Scholar]
- 46.Mori TA, Croft KD, Puddey IB, Beilin LJ. An improved method for the measurement of urinary and plasma F2-isoprostanes using gas chromatography-mass spectrometry. Anal Biochem 268: 117–127, 1999. [DOI] [PubMed] [Google Scholar]
- 47.Mori TA, Burke V, Puddey IB, Watts GF, O'Neal DN, Best JD, Beilin LJ. Purified eicosapentaenoic and docosahexaenoic acids have differential effects on serum lipids and lipoproteins, LDL particle size, glucose, and insulin in mildly hyperlipidemic men. Am J Clin Nutr 71: 1085–1094, 2000. [DOI] [PubMed] [Google Scholar]
- 48.Murphy RM, Stephenson DG, Lamb GD. Effect of creatine on contractile force and sensitivity in mechanically skinned single fibers from rat skeletal muscle. Am J Physiol Cell Physiol 287: C1589–C1595, 2004. [DOI] [PubMed] [Google Scholar]
- 49.Murphy RM, Snow RJ, Lamb GD. μ-Calpain and calpain-3 are not autolyzed with exhaustive exercise in humans. Am J Physiol Cell Physiol 290: C116–C122, 2006. [DOI] [PubMed] [Google Scholar]
- 50.Murphy RM, Goodman CA, McKenna MJ, Bennie J, Leikis M, Lamb GD. Calpain-3 is autolyzed and hence activated in human skeletal muscle 24 h following a single bout of eccentric exercise. J Appl Physiol 103: 926–931, 2007. [DOI] [PubMed] [Google Scholar]
- 51.Murphy RM, Larkins NT, Mollica JP, Beard NA, Lamb GD. Calsequestrin content and SERCA determine normal and maximal Ca2+ storage levels in sarcoplasmic reticulum of fast- and slow-twitch fibres of rat. J Physiol 587: 443–460, 2009. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 52.Murphy E, Steenbergen C. Mechanisms underlying acute protection from cardiac ischemia-reperfusion injury. Physiol Rev 88: 581–609, 2008. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 53.Nandhini TA, Anuradha CV. Inhibition of lipid peroxidation, protein glycation and elevation of membrane ion pump activity by taurine in RBC exposed to high glucose. Clin Chim Acta 336: 129–135, 2003. [DOI] [PubMed] [Google Scholar]
- 54.Palmi M, Youmbi GT, Fusi F, Sgaragli GP, Dixon HBF, Frosini M, Tipton KF. Potentiation of mitochondrial Ca2+ sequestration by taurine. Biochem Pharmacol 58: 1123–1131, 1999. [DOI] [PubMed] [Google Scholar]
- 55.Paolini C, Quarta M, Nori A, Boncompagni S, Canato M, Volpe P, Allen PD, Reggiani C, Protasi F. Reorganized stores and impaired calcium handling in skeletal muscle of mice lacking calsequestrin-1. J Physiol 583: 767–784, 2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 56.Parvez S, Tabassum H, Banerjee BD, Raisuddin S. Taurine prevents tamoxifen-induced mitochondrial oxidative damage in mice. Basic Clin Pharmacol Toxicol 102: 382–387, 2008. [DOI] [PubMed] [Google Scholar]
- 57.Pierno S, De Luca A, Camerino C, Huxtable RJ, Camerino DC. Chronic administration of taurine to aged rats improves the electrical and contractile properties of skeletal muscle fibers. J Pharmacol Exp Ther 286: 1183–1190, 1998. [PubMed] [Google Scholar]
- 58.Punna S, Ballard C, Hamaguchi T, Azuma J, Schaffer S. Effect of taurine and methionine on sarcoplasmic reticular Ca2+ transport and phospholipid methyltransferase activity. J Cardiovasc Pharmacol 24: 286–292, 1994. [PubMed] [Google Scholar]
- 59.Schaffer SW, Azuma J, Madura JD. Mechanisms underlying taurine-mediated alterations in membrane function. Amino Acids 8: 231–246, 1995. [DOI] [PubMed] [Google Scholar]
- 60.Schaffer SW, Azuma J, Mozaffari M. The role of antioxidant activity of taurine in diabetes. Can J Physiol Pharmacol 87: 91–99, 2009. [DOI] [PubMed] [Google Scholar]
- 61.Scholte HR, Yu Y, Ross JD, Oosterkamp II, Boonman AMC, Busch HFM. Rapid isolation of muscle and heart mitochondria, the lability of oxidative phosphorylation and attempts to stabilize the process in vitro by taurine, carnitine and other compounds. Mol Cell Biochem 174: 61–66, 1997. [PubMed] [Google Scholar]
- 62.Shin DW, Pan Z, Kim EK, Lee JM, Bhat MB, Parness J, Kim DH, Ma J. A retrograde signal from calsequestrin for the regulation of store-operated Ca2+ entry in skeletal muscle. J Biol Chem 278: 3286–92, 2003. [DOI] [PubMed] [Google Scholar]
- 63.Shiny KS, Kumar SH, Farvin KH, Anandan R, Devadasan K. Protective effect of taurine on myocardial antioxidant status in isoprenaline-induced myocardial infarction in rats. J Pharm Pharmacol 57: 1313–1317, 2005. [DOI] [PubMed] [Google Scholar]
- 64.Tappaz ML Taurine biosynthetic enzymes and taurine transporter: molecular identification and regulations. Neurochem Res 29: 83–96, 2004. [DOI] [PubMed] [Google Scholar]
- 65.Taveau M, Bourg N, Sillon G, Roudaut C, Bartoli M, Richard I. Calpain 3 is activated through autolysis within the active site and lyses sarcomeric and sarcolemmal components. Mol Cell Biol 23: 9127–9135, 2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 66.Timbrell JA, Seabra V, Waterfield CJ. The in vivo and in vitro protective properties of taurine. Gen Pharmacol 26: 453–462, 1995. [DOI] [PubMed] [Google Scholar]
- 67.Tricarico D, Barbieri M, Conte Camerino D. Taurine blocks ATP-sensitive potassium channels of rat skeletal muscle fibres interfering with the sulphonylurea receptor. Br J Pharmacol 130: 827–834, 2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 68.Tricarico D, Barbieri M, Conte Camerino D. Voltage-dependent antagonist/agonist actions of taurine on Ca2+-activated potassium channels of rat skeletal muscle fibers. J Pharmacol Exp Ther 298: 1167–1171, 2001. [PubMed] [Google Scholar]
- 69.Wan FS, Li GH, Zhang J, Yu LH, Zhao XM. Protective effects of taurine on myocardial mitochondria and their enzyme activities in rate with severe burn. Zhonghua Shao Shang Za Zhi 24: 171–174, 2008. [PubMed] [Google Scholar]
- 70.Wang J, Yan LI, Zhang L, Zhao J, Pang Y, Tang C, Zhang J. Taurine inhibits ischemia/reperfusion-induced compartment syndrome in rabbits. Acta Pharmacol Sin 26: 821–827, 2005. [DOI] [PubMed] [Google Scholar]
- 71.Warskulat U, Flogel U, Jacoby C, Hartwig HG, Thewissen M, Merx MW, Molojavyi A, Heller-Stilb B, Schrader J, Haussinger D. Taurine transporter knockout depletes muscle taurine levels and results in severe skeletal muscle impairment but leaves cardiac function uncompromised. FASEB J 18: 577–579, 2004. [DOI] [PubMed] [Google Scholar]
- 72.Westerblad H, Allen DG. Changes of myoplasmic calcium concentration during fatigue in single mouse muscle fibers. J Gen Physiol 98: 615–635, 1991. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 73.Yatabe Y, Miyakawa S, Ohmori H, Mishima H, Adachi T. Effects of taurine administration on exercise. Adv Exp Med Biol 643: 245–252, 2009. [DOI] [PubMed] [Google Scholar]
- 74.Yildirim Z, Kilic N, Ozer C, Babul A, Take G, Erdogan D. Effects of taurine in cellular responses to oxidative stress in young and middle-aged rat liver. Ann NY Acad Sci 1100: 553–561, 2007. [DOI] [PubMed] [Google Scholar]
- 75.Zhang SJ, Bruton JD, Katz A, Westerblad H. Limited oxygen diffusion accelerates fatigue development in mouse skeletal muscle. J Physiol 572: 551–559, 2006. [DOI] [PMC free article] [PubMed] [Google Scholar]