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MUSCLE CELL BIOLOGY AND CELL MOTILITY

O₂^– production at 37°C plays a critical role in depressing tetanic force of isolated rat and mouse skeletal muscle

¹Department of Zoology, La Trobe University, Bundoora, Victoria, Australia; and ²Institute of Physiology and Biophysics, University of Aarhus, Århus, Denmark

Submitted 26 January 2007 ; accepted in final form 22 April 2007

	ABSTRACT

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To find out whether the decrease in muscle performance of isolated mammalian skeletal muscle associated with the increase in temperature toward physiological levels is related to the increase in muscle superoxide (O₂^–) production, O₂^– released extracellularly by intact isolated rat and mouse extensor digitorum longus (EDL) muscles was measured at 22, 32, and 37°C in Krebs-Ringer solution, and tetanic force was measured in both preparations at 22 and 37°C under the same conditions. The rate of O₂^– production increased marginally when the temperature was increased from 22 to 32°C, but increased fivefold when the temperature was increased from 22 to 37°C in both rat and mouse preparations. This increase was accompanied by a marked decrease in tetanic force after 30 min incubation at 37°C in both rat and mouse EDL muscles. Tetanic force remained largely depressed after return to 22°C for up to 120 min. The specific maximum Ca²⁺-activated force measured in mechanically skinned fibers after the temperature treatment was markedly depressed in mouse fibers but was not significantly depressed in rat muscle fibers. The resting membrane and intracellular action potentials were, however, significantly affected by the temperature treatment in the rat fibers. The effects of the temperature treatment on tetanic force, maximum Ca²⁺-activated force, and membrane potential were largely prevented by 1 mM Tempol (4-hydroxy-2,2,6,6-tetramethylpiperidine-1-oxyl), a membrane-permeable superoxide dismutase mimetic, indicating that the increased O₂^– production at physiological temperatures is largely responsible for the observed depression in tetanic force at 37°C by affecting the contractile apparatus and plasma membrane.

intact mammalian muscle; physiological temperature; superoxide; excitation-contraction coupling; maximum Ca²⁺-activated force; muscle excitability; cytochrome c assay

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There are only a few studies conducted at 37°C with isolated intact mammalian skeletal muscle that examine muscle contractility, as there is a relatively rapid decline in muscle function seen at temperatures above 35°C (9, 16, 27, 31, 40, 41). The longevity of the preparation can be increased by either circulating the Ringer solution bathing the muscle (32) or by having the preparation bathed in a large volume of solution (6), suggesting that deterioration in isolated mammalian skeletal muscle at physiological temperature may be caused by diffusion-limited processes. For example, for many decades, the decline in isolated mammalian skeletal muscle function at physiological temperature was thought to be due to the relatively slow diffusion rate of oxygen into the muscle as a consequence to the removal of vascular perfusion, resulting in an anoxic region in the center of an isolated muscle preparation skeletal muscle (12). Yet recent work by Barclay (4) has shown that diffusive oxygen supply would be adequate to support resting metabolism and energy requirements at low duty cycles in isolated intact rat and mouse extensor digitorum longus (EDL) muscle at 37°C. In addition, the decline in skeletal muscle function at physiological temperature is seen at the level of the single fiber, where oxygen supply to the muscle cell is not limited (16), clearly suggesting that the deterioration in isolated mammalian skeletal muscle function at physiological temperatures is not simply due to the lack of oxygen supply. Diffusion-limited processes can also lead to a buildup of substances in the fibers that could interfere with muscle function, such as ROS, particularly if the rate of production of these substances is increased. Because we could show that O₂^– produced by skeletal rat muscle fiber mitochondria at 37°C can depress the twitch force response in mechanically skinned muscle fibers depleted of cytosolic antioxidants and ROS scavengers (45), it was important to find out whether increasing O₂^– production by raising the muscle temperature is in any way associated with the depression of muscle performance in isolated intact mammalian muscle preparations.

The results clearly show that as the muscle temperature is increased above 32°C, there is a sharp increase in O₂^– produced by the intact rat and mouse muscle, the depression of intact muscle performance at 37°C is markedly attenuated in the presence of a membrane-permeable SOD mimetic and that O₂^– radicals produced in the isolated rat and mouse muscle at physiological temperature interfere with aspects of the excitation-contraction coupling causing depression of the force response to stimulation.

	METHODS

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Animals and preparation of intact muscles. Experiments were performed on the extensor digitorum longus (EDL) muscles of 5- to 15-wk-old mice and 12-wk-old rats that were fed ad libitum and kept in a controlled environment at 21 ± 1°C with a 12:12-h light-dark cycle. Experiments were conducted at La Trobe University, on C57BL/10 mice and Long-Evans hooded rats, which were anesthetized and killed by halothane overdose in accordance with the procedures approved by the Animal Ethics Committee. Experiments were also conducted at Aarhus University on Wistar rats, where the animals were killed by cervical dislocation, followed by decapitation in compliance with Danish animal welfare regulations. In each case, the EDL muscles were rapidly excised and immersed in a Krebs-Ringer solution (KRS) containing (in mM): 122 NaCl, 2.8 KCl, 1.3 CaCl₂, 1.2 MgSO₄, 1.2 KH₂PO₄, 25 NaHCO₃, and 5 D-glucose, bubbled with carbanox (95% oxygen-5% CO₂), pH = 7.4 at 22°C.

O₂^– measurements. The cytochrome c (CytC) assay was used to measure O₂^– in KRS, in which whole intact rat or mouse EDL muscle was incubated at either 22, 32, or 37°C for 30 min with and without various ROS scavengers and reactive nitrogen species (RNS) inhibitors/donors.

CytC is membrane impermeable and when oxidized, CytC(Fe³⁺) can be reduced to CytC(Fe²⁺) through a one-electron transfer reaction (51). CytC(Fe²⁺) has a different absorbance spectrum from that of CytC(Fe³⁺), with a marked increase in peak absorbance at 550 nm (Fig. 1). Absorbances were measured at 22°C with a Hitachi U-1100 (Japan) spectrophotometer. In the presence of O₂^–, CytC(Fe³⁺) is reduced to CytC(Fe²⁺), according to the following reaction:

(1)

Isosbestic points for oxidized and reduced CytC are observed at the wavelengths of 542 and 557 nm (Fig. 1). Using absorbance measurements at 550 nm and at the isosbestic points, the relative amount of reduced CytC of the total CytC present CytC(Fe²⁺)/(CytC_total) can be calculated using Eq. 2:

(2)

where Max_peak is maximum height of the 550 nm absorbance peak measured from the baseline passing through the isosbestic points, when all CytC is reduced in the presence of a cysteine-based solution (see Fig. 1).

View larger version (8K): [in this window] [in a new window]   Fig. 1. Absorbance spectra for fully oxidized () and fully reduced () cytochrome c (CytC; 5 µM) in Krebs-Ringer solution (KRS). CytC was fully oxidized by the addition of 10 mM potassium ferricyanide and fully reduced by the addition of 10 mM sodium dithionite, respectively, to a 10 mM-based cysteine solution, as described by Margoliash and Frohwirt (18).

(3)

where V = volume of the CytC solution in milliliters, and the coefficient 35.7 represents the ratio between total [CytC] (5 µM) and Max_peak (0.14). At the completion of experiments, muscles were weighed on digital scales (Mettler, Switzerland) without blotting.

The rates of reducible molecular species produced by the muscle expressed per unit muscle mass and time can then be calculated. Note that in solution, the reduced CytC remained stable for at least 1 h, which was well within the time when the measurements were made. Furthermore, there were no changes in the absorbance spectra of CytC in solution maintained at 22°C or heated at 37°C. In addition, no changes in the absorbance spectra of CytC were observed when CytC solutions were bubbled with 95% O₂-5% CO₂ compared with CytC solutions that were not bubbled.

Of the ROS and RNS and their derivatives produced by the muscle in the extracellular space, CytC is known to be reduced rapidly by superoxide and oxidized by the hydroxyl radicals, hydrogen peroxide, and nitric oxide (^·NO). On the basis of experiments using denitrifying bacteria, it was originally suggested that ^·NO can also reduce CytC (for a review, see Ref. 22), but more direct experiments with ^·NO and CytC have shown that ^·NO actually oxidizes CytC and reversibly binds to the oxidized CytC without reducing it (42). ^·NO also reacts rapidly with O₂^– forming peroxynitrite that is membrane permeant. To find out whether ^·NO and/or peroxynitrite produced by skeletal muscle at 37°C can cause CytC reduction, an experiment was performed on mouse EDL muscles incubated at 37°C in the presence of oxidized CytC, and a relatively large concentration of nitroprusside (1 mM), a potent donor of ^·NO used with muscle (1). In contrast to results obtained in the absence of the ^·NO donor, when EDL muscles incubated at 37°C produced marked CytC reduction (see RESULTS), there was a 88.3 ± 0.5% (P < 0.0001) drop in CytC reduction in experiments with paired muscles from three mice, when large amounts of ^·NO were produced. This confirms observations made by others showing markedly decreased CytC reduction when both O₂^– and ^·NO are produced by muscle tissue (13). The decrease in CytC reduction under these circumstances is due to the removal of O₂^– in the presence of nitric oxide with the formation of peroxynitrite, which does not cause CytC reduction (10) but can cause CytC oxidation (43). Thus, O₂^– appears to be the only species of ROS or RNS that can actually reduce CytC. Therefore, the use of the CytC assay for measuring O₂^– production is rather conservative because, if anything, it underestimates rather than overestimates the true rate of O₂^– production in the presence of various ROS and RNS. Nevertheless, having said that, great care needs to be taken when making O₂^– measurements using the CytC assay to correct for any changes induced on the CytC assay by factors other than the muscle preparation per se, such as the addition of a chemical or the effect of temperature. Therefore, blank tests without muscle preparations were run in parallel with all proper experiments with the muscle present and were used to correct for any effects of such factors on the CytC assay.

Force Measurement Experiments

Intact muscle preparations. After dissection, the mouse EDL muscles were tied to stainless-steel pins connected to a force transducer (Grass FT03; Grass Instruments, West Warwick, RI) and micromanipulator (Mitutoy, Tokyo, Japan). Muscles were then placed between stimulating electrodes in a well milled out of Perspex and filled with 3 ml KRS, bubbled with carbanox at room temperature (22 ± 2°C). Force responses were recorded on a chart recorder (Linear, Irvine, CA) or on both the chart recorder and a PC via a PowerLab (AD Instruments, Sydney, Australia). After attachment and immersion in KRS, muscles were allowed to equilibrate for 15 min, during which time the length of the muscle was adjusted to develop optimal tetanic force at 100-Hz stimulation with 1-ms-long supramaximal pulses (train duration 0.5 s; field stimulation 20 V/cm). Muscles were then transferred to a second chamber containing 3 ml KRS constantly bubbled with carbanox and maintained at 37 ± 0.1°C. The mouse EDL muscle produced maximum tetanic force at 100-Hz stimulation at room temperature and at 200 Hz at 37°C (see also Ref. 2). A higher stimulation frequency is required for maximal tetanic force response at 37°C in the mouse than in the rat EDL muscle because of the considerably faster twitches in the mouse EDL than in the rat EDL muscle. The mouse EDL muscles were stimulated (at 100 Hz) every 15 min at 22°C. At 37°C, muscles were stimulated (at 200 Hz) only twice; first, 1 min after the transfer into KRS kept at 37 °C and then once after 30 min, before returning the preparation to KRS at 22 °C. Subsequently, the muscles were stimulated every 15 min in KRS at 22°C for a further 135 min.

Similar experiments were performed on isolated Wistar rat EDL muscles. In this case, the muscles were prepared with 10 mm of the nerve still attached and incubated in standard KRS. Muscles were initially mounted for isometric contractions at 22°C in thermostated chambers containing standard KRS (7 ml) and adjusted to optimal length for force production. The muscles were stimulated to contract tetanically (0.5 s trains, 100 Hz, 0.02-ms pulse duration every 10 min) by applying field stimulation (12 V/cm) across the central region of the muscle through platinum wire electrodes until steady state force was achieved. The temperature was then rapidly increased to 37 ± 0.1°C, with the muscles stimulated every 5 min for 40 min before being returned to 22°C and stimulated every 10 min for a further 50 min. Tempol (4-hydroxy-2,2,6,6-tetramethylpiperidine-1-oxyl) was mixed well in the 22 and 37°C KRS before the transfer of the muscle to that solution.

Skinned fiber preparations. Single mechanically skinned fiber preparations were obtained from Long-Evans hooded rat and mouse EDL muscles that were either kept in 22°C KRS for 30 min or incubated in KRS at 37°C for 30 min before were well blotted on filter paper and transferred to paraffin oil (Ajax, Sydney, Australia) at room temperature (22 ± 2 °C). Single fibers were dissected, mechanically skinned, and viewed at high magnification (x200) on the screen of a TV monitor coupled to a Leitz dissecting microscope. The cross-sectional area was calculated assuming the fiber was circular with a diameter equivalent to the average width of the fiber obtained from at least four measurements along the fiber length (48). The skinned fiber segment was then mounted under oil onto a force transducer (AME 875; Horten, Norway) using fine forceps (jeweler's forceps no. 5) as previously described (48) and then was placed in a relaxing solution containing (in mM): 90 HEPES, 50 EGTA, 1 Mg²⁺, 126 K⁺, 36 Na⁺, 8 ATP, 10 CP; pH = 7.10, pCa (= –log₁₀ [Ca²⁺]) >9 for 2 min. The preparation was then transferred into a series of Ca²⁺-activating solutions containing (in mM): 90 HEPES, 50 Ca-EGTA + EGTA (pCa between 7 and 4.5), 1 Mg²⁺, 126 K⁺, 36 Na⁺, 8 ATP, 10 CP, pH = 7.10 for 5–10 s until force reached a steady value and then returned to the relaxing solution for a further 2 min. Solutions were made as previously described by Stephenson and Williams (42). Force responses were recorded on a chart recorder (Linear), and the specific maximum force was calculated from the maximum force response at pCa 4.5 divided by the cross-sectional area of the fiber.

Electrophysiological Experiments

The resting membrane potential (RMP) and intracellular action potentials (APs) were measured in the outer layer of fibers from Wistar rat EDL muscles by inserting a single intracellular recording microelectrode into an individual fiber as previously described (17). APs were triggered in the muscle by stimulating indirectly via the nerve, using a suction electrode, with single 5-mA constant current pulses. The glass microelectrodes were filled with 3 M KCl and typically had resistances between 10 and 25 M. The microelectrodes were connected to an Axoclamp-2a amplifier, and recordings of membrane potential were displayed on an oscilloscope and recorded by a computer, using Signal 2.09 software (Cambridge Electronic Design, Cambridge, UK), at a minimum of 27 kHz. RMP was measured from the baseline of the AP. AP amplitude was defined as the difference between RMP and the peak potential of the action potential. The maximum rates of depolarization and repolarization of the action potential were determined as the maximum and minimum of the first differential of the action potential signal, respectively. All measurements were made at 22°C before and after muscle exposure to 37°C in KRS.

Chemicals

All Chemicals, unless otherwise noted, were obtained from Sigma (Castle Hill, New South Wales, Australia).

Statistical Analyses

Results are expressed as means ± SE. Curve fitting and statistical analyses were performed using GraphPad Prism software (San Diego, CA). Statistical significance was tested at P < 0.05 using one-way ANOVA and Student's t-tests, as appropriate.

	RESULTS

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Effect of Temperature on O₂^– Detected Extracellularly

O₂^– produced by intact rat and mouse EDL muscles incubated at different temperatures was detected in the KRS using the CytC assay (Fig. 1). When the temperature was increased from 22 to 32°C, the average rate of CytC reduction measured in nanomoles reducing equivalents per minute gram wet weight over a 30-min interval for intact rat EDL muscle increased marginally from 2.3 ± 0.3 at 22°C to 3.4 ± 0.2 at 32°C in paired experiments (n = 4, P < 0.05, one-sided Student's t-test) and remained effectively constant for intact mouse EDL muscle (2.3 ± 0.1 at 22°C vs. 2.7 ± 0.2 at 32°C in paired experiments, n = 6, P > 0.05, one-sided Student's t-test). However, when the temperature was increased from 22 to 37°C, there was a fivefold increase in the rate of CytC reduction (measured in nanomoles reducing equivalents per minute per gram wet weight) for both rat (2.2 ± 0.3 at 22°C vs. 10.5 ± 0.9 at 37°C in paired experiments using intact rat EDL muscle, n = 6, P < 0.001) and mouse (2.3 ± 0.3 at 22°C vs. 11.1 ± 1.0 at 37°C in paired experiments using intact mouse EDL muscle, n = 6, P < 0.001) (see Fig. 2).

View larger version (10K): [in this window] [in a new window]   Fig. 2. Rates of superoxide entering KRS at 22, 32, and 37°C. Extracellular O2– (in nanomoles per minute per gram wet weight) was measured in KRS using the CytC assay for both isolated rat and mouse EDL muscle preparations treated at either 22, 32, or 37°C. When the temperature of intact rat EDL muscle was increased from 22 to 32°C (A), the measured CytC reduction in KRS after 30 min marginally increased [paired muscles from same animal, n (rats) = 4, P < 0.05, one-sided Student's t-test] yet effectively remained constant for intact mouse (C) EDL muscle under the same conditions [paired muscles from same animal, n (mice) = 6, P > 0.05, one-sided Student's t-test]. A fivefold increase in the rate of CytC reduction was observed when the muscle temperature was increased from 22 to 37°C for both rat (B; n = 6, P < 0.001) and mouse (D; n = 6, P < 0.001) EDL muscle. Statistical significance: *P < 0.05; ***P < 0.001 for responses at 32°C/37°C compared with responses at 22°C.

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To find out whether hydrogen peroxide produced in the muscle significantly affects the reduction/oxidation of CytC in the KRS at 37°C, experiments were performed on three pairs of EDL muscles from three mice in the absence and in the presence of 30 µM ebselen, a membrane-permeant glutathione peroxidase mimetic (38). Because the results (expressed as nanomoles reducing equivalents per minute per gram wet weight) in the absence of ebselen were close and not statistically significantly different from the results in the presence of ebselen (9.0 ± 0.2 vs. 8.3 ± 0.2, n = 3, P > 0.05, paired Student's t-test), one can conclude that hydrogen peroxide does not significantly alter the reduction/oxidation of CytC in the KRS under our conditions.

Note that all extracellular measurements of superoxide presented so far were from experiments in which the muscles were left free in solution and that results (expressed in nanomoles reducing equivalents per minute per gram wet weight) from experiments with mouse EDL muscles kept at optimal length for tetanic force (37°C in KRS for 30 min) were not significantly different from those with free muscles in solution (8.7 ± 0.3, n = 3 vs. 9.3 ± 0.9, n = 17, P > 0.8, unpaired t-test).

Effect of Temperature on Tetanic Force Production in Intact Mouse and Rat Muscle

To assess the effect of temperature on force production in intact mouse and rat EDL muscles for conditions similar to those in which O₂^– was measured, muscles were briefly (0.5 s) tetanically stimulated every 15 min (mouse) or every 5–10 min (rat) for up to 165 min (mouse) and 90 min (rat) at 22°C (control responses) or were stimulated with the same pattern as control preparations before transfer from KRS at 22°C to KRS at 37°C for 40 min (rat) and 30 min (mouse) and after transfer from KRS at 37°C to KRS at 22°C (test responses). Representative force responses obtained with the rat EDL muscle for control and test procedures are shown in Fig. 3. Control experiments with mouse muscle showed that maximal tetanic forces at 22°C and 37°C were achieved when muscles were stimulated with supramaximal stimuli at 100 Hz and 200 Hz, respectively (see METHODS). Therefore, in all experiments, the mouse muscles were stimulated at 100 Hz at 22°C and at 200 Hz at 37°C. For rat muscle, maximum tetanic force was achieved at 100 Hz stimulation, both at 22 and 37°C. Tetanic force responses in intact rat (n = 4) and mouse (n = 3) EDL muscle kept at 22°C remained relatively stable (Fig. 4). In the rat muscle, there was a small drop in force of 0.17%/min, while in the mouse, there was effectively no decrease in the tetanic force after 160 min stimulation at 22°C (101 ± 2% initial tetanic force, n = 3). The greater decline in tetanic force seen in the rat than in the mouse EDL muscle at room temperature may be related to diffusional problems associated with the larger size of the rat EDL compared with that of the mouse EDL muscle. In contrast, tetanic force production after 30 min in KRS at 37°C decreased markedly to 49.0 ± 4.0% initial tetanic force in rat EDL muscles (n = 4, one-way ANOVA, P < 0.01) and to 31.0 ± 12.0% initial tetanic force in mouse EDL muscles (n = 6, one-way ANOVA, P < 0.01).

View larger version (9K): [in this window] [in a new window]   Fig. 3. Representative traces of tetanic force production in two rat EDL muscles. Muscle kept in KRS at 22°C for 90 min (A). Muscle kept initially at 22°C in KRS (first trace), then exposed for 40 min to 37°C in KRS (second trace), and thereafter returned to 22°C in KRS for 50 min (third trace, 90 min) (B).

View larger version (18K): [in this window] [in a new window]   Fig. 4. Changes in tetanic force production in rat (A) and mouse (B) EDL when exposed to 22 and 37°C. Tetanic force production in muscles kept in KRS at 22°C (open bars; n = 4 for rat, n = 3 for mouse). Tetanic force production significantly declined (P < 0.01) in both rat and mouse EDL muscle preparations after being exposed to 37°C for 30 min (stacked dark gray bars; n = 4 for rat, n = 6 for mouse) and recovered marginally after 50 min (rat) and 135 min (mouse) in KRS at 22°C (simple dark gray bars; n = 4 for rat, n = 6 for mouse). 1 mM Tempol significantly prevented the decline in tetanic force responses at 37 °C in rat and mouse muscle preparations (stacked light gray bars; n = 8 for rat, n = 5 for mouse) and further increased tetanic force recovery in KRS at 22 °C (simple light gray bars; n = 8 for rat, n = 5 for mouse). There was no statistically significant difference at 165 min between tetanic forces from muscles exposed to 1 mM Tempol and muscles kept throughout at 22°C in mouse (P > 0.05, one-tailed Student's t-test); however, muscles did differ at 90 min in rat (P < 0.05, one-tailed Student's t-test). Statistical significance: *P < 0.05; **P < 0.01 for respective responses compared with responses at 22°C; P < 0.05 for respective responses after recovery at 22°C compared with last responses at 37°C (40 min for rat, 30 min for mouse).

Changes in Tetanic Force and O₂^– Production in the Presence of Tempol

Tempol is a stable, membrane-permeable nitroxide that acts as a SOD mimetic, and its action can therefore be compared with that of an enzyme (15). Tempol is first oxidized by the protonated form of superoxide (·OOH) to yield H₂O₂ and an oxo-ammonium cation, which is then further reduced by another superoxide molecule to yield O₂ and regenerate the Tempol molecule, with the net removal of two superoxide molecules (5, 15). Tempol is therefore ideally suited to find out whether the increased O₂^– production at 37°C is responsible for the decrease in tetanic force production since, unlike classical superoxide scavengers, it is not used as substrate while "scavenging" O₂^–. In experiments in which isolated mouse (n = 5) and rat (n = 8) EDL muscles were equilibrated in KRS at 22°C with 1 mM Tempol and then transferred to 37°C with 1 mM Tempol and incubated for 30 min, tetanic force decreased significantly less than when no Tempol was present in solutions (one-way ANOVA, P < 0.05 for both mouse and rat EDL muscles) (Fig. 4). Furthermore, upon return to 22°C, tetanic force recovered to values that were rather close to the corresponding control values for muscles incubated at 22°C only (Fig. 4). The protective effect of Tempol on the tetanic response at 37°C was smaller at Tempol concentrations below 1 mM and almost completely disappeared when Tempol concentration was increased from 1 to 5 mM (data not shown). Importantly, the rate of O₂^– production detected extracellularly at 37°C in mouse EDL muscles was more than halved in the presence of 1 mM Tempol (4.4 ± 0.8, n = 4 vs. 9.3 ± 0.9 nanomoles reducing equivalents per minute per gram wet weight, n = 17, P < 0.05).

When 1 mM Tempol was present in the KRS at 22°C, there was no change in tetanic force in mouse EDL muscles (100.8 ± 1.0%, n = 3), but increasing Tempol concentration to 5 mM caused a 20% decrease in tetanic force after 20 min (80.0 ± 8.97%, n = 3). Also, 1 mM Tempol did not significantly (P > 0.05) affect the kinetics of the tetanic force response (e.g., 20 to 80% rise time at 22°C for rat EDL muscle: 0.064 ± 0.004 s, n = 8 in the absence of Tempol vs. 0.065 ± 0.005 s, n = 8 in the presence of Tempol and 80 to 20% relaxation time at 22°C for rat EDL muscle: 0.027 ± 0.002 s, n = 8 in the absence of Tempol vs. 0.032 ± 0.002 s, n = 8 in the presence of Tempol).

The inhibitory effect of 5 mM Tempol on the force response is most likely related to the increase in the concentration of the oxo-ammonium cation, the oxidized form of Tempol, which at higher concentrations, will oxidize instead of protect target molecules from oxidation. This is because under conditions of high concentrations of synthetic SOD mimetics like Tempol and native Cu,Zn SODs, the [O₂^–] drops to very low levels and therefore, to regenerate the reduced state of the SOD or SOD mimetic molecule, the second-order reaction requires the marked buildup in the concentration of the oxidized state of the SOD (Cu²⁺, Zn-SOD) or SOD mimetic molecule (the oxo-ammonium cation) to compensate for the decrease in [O₂^–] (25).

Because the oxidation of Tempol by O₂^– yields H₂O₂ (see above), one can also rule out the possibility that the drop in tetanic response after 30 min exposure to 37°C is related to H₂O₂ production. This is because although more H₂O₂ is produced in the presence of 1 mM Tempol than in its absence, the decline in the force response is markedly prevented instead of being enhanced.

Thus, the results obtained with Tempol strongly suggest that a large part of the decrease in the force production in intact rat and mouse muscle fibers at 37°C is related to the increased production of superoxide, which can act at different sites in the process of excitation-contraction coupling.

Maximum Ca²⁺-Activated Force and Sensitivity to Ca²⁺

The decline in tetanic force after muscle exposure to 37°C in the absence of Tempol could be due to effects on the contractile apparatus. Thus, force would be reduced if the ability of the contractile apparatus to generate maximum Ca²⁺-activated force and/or the sensitivity to Ca²⁺ were reduced.

Indeed, the capacity of the contractile apparatus in rat muscle fibers to develop force was shown to decline in response to increased temperature (48). We measured, therefore, the maximum Ca²⁺ -activated specific force in mechanically skinned fibers isolated from intact rat and mouse muscles treated at 37°C for 30 min and from contralateral muscles maintained throughout at 22°C.

As shown in Fig. 5A, the maximum Ca²⁺-activated specific force of mechanically skinned fibers from rat EDL muscle treated at 37°C for 30 min was not significantly lower than that from fibers kept at 22°C (176.3 ± 19.5 kN/m², n = 6 vs. 196.7 ± 21.5 kN/m², n = 6; P > 0.1, one-tailed t-test). In contrast, however, the skinned fibers from the 37°C treated mouse EDL muscles produced 55% less specific force than those from the 22°C control muscles (77 + 12 kN/m², n = 16 vs. 163.6 ± 17.4 kN/m², n = 10; P < 0.001; see Fig. 5 caption). Importantly, this decline in specific maximum Ca²⁺-activated force was not seen in the presence of 1 mM Tempol (174.1 ± 17.4 kN/m², n = 4), as the maximum Ca²⁺-activated force was not significantly different from that in fibers kept at 22°C (P > 0.5).

View larger version (15K): [in this window] [in a new window]   Fig. 5. Maximum Ca2+-activated force production was measured at 22°C in mechanically skinned muscle fibers from both rat (A) and mouse (B) muscles. Measurements were made at 22°C from mechanically skinned muscle fibers obtained from EDL muscle kept either at 22°C (open bars; n = 6 for rat, n = 10 for mouse), after exposure to 37°C for 30 min (dark gray bars; n = 6 for rat, n = 16 for mouse). B: light gray bar shows maximum Ca2+-activated force production in mechanically skinned muscle fibers from mouse muscles exposed to 37°C for 30 min in the presence of 1 mM Tempol (n = 4). Results shown by the dark gray bar in B are significantly different from results shown by the clear and the light gray bars (P < 0.001, one-way ANOVA).

View larger version (6K): [in this window] [in a new window]   Fig. 6. Isometric force-pCa curves obtained at 22°C with mechanically skinned muscle fibers from mouse EDL muscles kept in KRS at 22°C () and after muscles were exposed to 37°C in KRS for 30 min (). Number of fibers = 6 for both treatments. Measurements were made at 22°C from mechanically skinned muscle fibers obtained from EDL muscle kept either at 22°C () or after exposure to 37°C for 30 min ().

Because the decrease in force production in rat muscle could not be explained by the ability of the contractile apparatus to produce force, experiments were performed to determine whether there were any changes in rat muscle fiber excitability.

For this, the RMP and intracellular APs were measured in single fibers from the outer layer of fibers from rat EDL muscles to identify whether changes in the fiber excitability could be responsible for the deterioration in muscle performance at 37°C. As seen in Fig. 7A, the RMP becomes significantly depolarized (P < 0.001) by about 10 mV (from approximately –80 to –70 mV) and the amplitude of the AP is reduced when the muscle is incubated at 37°C for 30 min (Fig. 7B). The addition of 1 mM Tempol to the KRS before and during incubation at 37°C did prevent the depolarization of the cell membrane at 37°C, and the amplitude of the AP was not different from that of control muscles that were not exposed to 37°C.

View larger version (12K): [in this window] [in a new window]   Fig. 7. Resting membrane potential (A), action potential amplitude (B), action potential maximum rates of depolarization (C) and action potential maximum rates of repolarization (D) in rat EDL fibers. Open bars: measurements made from single fibers (n = 21) from muscles kept at 22°C in KRS. Measurements made from single fibers from muscles that were exposed to 37°C in KRS for 40 min (solid bars; n = 25), or muscles that were exposed to 37°C in KRS for 40 min in the presence of 1 mM Tempol (gray bars; n =25). These measurements were made at 22°C within 5 min (45 min results) or 50–60 min after the muscle was returned to KRS at 22°C (100 min results). Statistical significance: *P < 0.05, **P < 0.01, ***P < 0.001 for respective results compared with responses at 22°C; P < 0.01, P < 0.001 for respective responses after recovery at 22°C compared with last responses at 37°C.

	DISCUSSION

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Increased O₂^– Production and Decreased Muscle Function

The results presented here clearly show that when isolated intact rat and mouse EDL muscles are incubated in a relatively small volume of KRS (3–7 ml) at 22°C, there is little depression in the tetanic force response for several hours with our low-duty cycle pattern of stimulation. At 22°C, there is also little O₂^– detected in the KRS, in which the muscles are incubated. However, when muscles are exposed to 37°C in KRS, there is a much greater amount of O₂^– detected than at 22°C, and this is accompanied by a marked and largely irreversible decline in tetanic force. Importantly, the irreversible decrease in tetanic force seen after exposure to KRS at 37°C was largely prevented in both rat and mouse in the presence of 1 mM Tempol, which similarly to SOD, is not used like a substrate during O₂^– removal but can be continuously regenerated (15). Tempol is not electrically charged and is membrane permeable, allowing it to penetrate all intracellular compartments of the muscle fibers and remove O₂^– where it is produced. In our previous study (45), we have shown that in the resting rat EDL muscle at 37°C, mitochondria are the major source of O₂^– measured either in the myoplasm or in the extracellular compartment. Because Tempol is membrane permeable, it will enter the mitochondrial matrix where most O₂^– is produced.

The fact that 1 mM Tempol had a marked protective effect on tetanic responses of isolated intact rat and mouse EDL muscle at 37°C further strengthens our previous observation on the protective effect of 1 mM Tempol on twitch responses at 37°C in mechanically skinned fibers of the rat (45) and demonstrates that the presence of diffusible myoplasmic ROS scavengers, which are lost from the mechanically skinned fiber, are not sufficient to prevent the gradual loss of muscle function in isolated intact preparations at 37°C. Therefore, Tempol (1 mM) may be used in isolated skeletal muscle preparations exposed to elevated temperatures to reduce the deterioration of force production and muscle function. Note, however, that like SOD and other nitroxide SOD mimetics, Tempol loses its protective action at higher concentrations (5 mM), probably because of its increased pro-oxidative action due to the marked increase in the concentration of its oxidized state (oxo-ammonium ion) as a consequence of the decrease of [O₂^–] to very low levels (25). Taken together, the results provide evidence that the tetanic force depression occurring at 37°C in isolated intact rat and mouse EDL muscles is closely related to the increased O₂^– production. The protective effect of 1 mM Tempol on muscle function also rules out the involvement of hydrogen peroxide in the gradual loss of muscle performance reported here at 37°C because, as mentioned above, hydrogen peroxide is produced in the first step of reaction of Tempol with O₂^–.

Isolated mammalian skeletal muscle lacks endogenous extracellular SOD (EC-SOD), a SOD isoform present in plasma and in the extracellular fluid, which binds reversibly, with a relatively high affinity to multiple sites on collagen and the basement membrane of muscle fibers through a heparin-binding carboxy terminal on its tail (23, 26). Using EC-SOD knockout mice, Park et al. (26) recently showed that EC-SOD protects skeletal muscle in vivo from O₂^–-induced injury. The presence of EC-SOD bound to multiple sites on collagen and the basement membrane of muscle fibers in vivo and in situ would act as a sink, preventing the accumulation of O₂^– in the vicinity of the plasma membranes in the extracellular space, thus creating conditions for the efficient loss of O₂^– from the intracellular environment. By removing the O₂^– sink (represented by EC-SOD), O₂^– will accumulate in isolated muscle to higher levels than would occur in vivo.

It is relevant to point out here that O₂^– can cross the membranes not only in its protonated form but also in exchange for bicarbonate (HCO₃^–) via the HCO₃^–-chloride anion exchanger (24) that is expressed in skeletal muscle (49).

An interesting observation from the present study is that muscle O₂^– production measured in KRS increased only marginally, as the temperature increased from 22 to 32°C but increased many fold, as the temperature increased to 37°C. This may explain why most attempts to experiment with isolated mammalian skeletal muscle at physiologically relevant temperatures stop at temperatures in the vicinity of 32°C.

O₂^– also increases markedly during intense muscle contraction (14, 35, 36) and has been implicated in contributing to muscle fatigue as O₂^– scavengers such as Tiron and Tempol have been shown to help prevent fatigue in isolated single intact mouse fibers stimulated at high frequency, while incubated at 37°C (21).

Sites of O₂^– Action in Muscle

The main sites of O₂^– action in skeletal muscle responsible for the decrease in force appear to be different in the rat and mouse EDL muscles treated at 37°C, even though the apparent rates of O₂^– production measured in the KRS were very similar for the two muscles.

Thus, a large proportion of the depression in tetanic force in mouse EDL muscles exposed to 37°C for 30 min compared with controls (Fig. 4B) can be explained by the reduced ability of the contractile apparatus to produce maximal force due to O₂^– production as indicated by the 55% decrease in the maximally Ca²⁺-activated specific force in mechanically skinned fibers from muscles exposed for 30 min to 37°C (Fig. 5B). Importantly, when the muscles were incubated at 37°C in the presence of 1 mM Tempol, there was no statistically significant drop in the capacity of the contractile apparatus to produce force. This shows that the contractile apparatus in isolated mouse EDL muscle is actually quite susceptible to O₂^– causing oxidation of various chemical groups on the contractile apparatus, most likely on the myosin heads (7, 48, 50). In contrast, in the rat EDL muscle, the tetanic force dropped by about 60% after 40 min in KRS at 37°C but without significant drop in specific maximum Ca²⁺-activated force (Fig. 5A).

There was no change in the sensitivity of the contractile apparatus to Ca²⁺ when mouse EDL muscles were exposed to 37°C for 30 min, and we have also previously shown that there was no change in the sensitivity of the contractile apparatus to Ca²⁺ in resting isolated rat muscle exposed to temperatures in the range 40–43°C (48). This was also shown to be the case by Moopanaar and Allen (20), where a decrease of myofibrillar Ca²⁺ sensitivity in mouse fibers only occurred when the muscle was fatigued.

Because the tetanic force dropped by about 60% after 40 min in KRS at 37°C in the rat EDL fibers but without a decrease in myofibrillar Ca²⁺ sensitivity and without a significant drop in maximum Ca²⁺-activated force (Fig. 5B), the major site of O₂^– action in the rat fibers must be at another step in the excitation-contraction coupling. As seen in Fig. 7A, there was significant depolarization of the rat fibers by about 10 mV after exposure to 37°C. This depolarization must cause some Na⁺ channels to slow-inactivate, resulting in observed decreased AP amplitude (Fig. 7B) and rates of depolarization (Fig. 7C) and repolarization (Fig. 7D), which would translate in decreased activation of the voltage sensors, decreased Ca²⁺ release and decreased force output. Interestingly, 1 mM Tempol prevented the depolarization of the fibers at rest, the reduction in the amplitude of the AP, and the decrease in the maximal rate of repolarization, but it did not prevent the decrease in the maximal rate of depolarization during an AP (Fig. 7) and only partially prevented the decline of the tetanic force (Fig. 4B). One explanation for the decrease in the resting membrane potential at 37°C in the absence of Tempol could be that the elevated O₂^– increases Na⁺ permeability (see discussion in Ref. 45) and decreases the activity of the Na⁺-K⁺-pump. A recent study by McKenna et al. (19) showed that infusion of the nonspecific ROS scavenger, N-acetylcysteine, conveyed a protective effect on the reduction in maximal Na⁺-K⁺ pump activity induced by fatiguing exercise. A reduced Na⁺-K⁺ pump activity would result in membrane depolarization, leading to slow inactivation of Na⁺ channels (37) and reduction in the Na⁺ current responsible for the depolarizing phase of the action potential. This may be further exacerbated by an elevated intracellular [Na⁺], reducing the driving force for the Na⁺ current. These effects can be seen from the reduction in maximum rate of depolarization and the marked reduction in AP amplitude after exposure to 37°C for 40 min (Fig. 7). The reduced AP amplitude would also result in a slower rate of repolarization as the driving force for the K⁺ current is now smaller. The ability of Tempol to prevent the depolarization but not the reduction in maximum rate of depolarization of the AP following 40 min at 37°C, suggests that there may also be an effect directly on the Na⁺ channel. Regardless, of the mechanism, it is clear that elevation of muscle temperature from 22 to 37°C causes a significant reduction in muscle excitability.

The small drop in the tetanic force in mouse EDL muscles kept at 37°C for 30 min in the presence of 1 mM Tempol is likely due to some events in the excitation-contraction coupling such as Ca²⁺ release from the SR that are more directly affected by prolonged exposure to physiological temperature. Indeed, the ability of the SR to accumulate Ca²⁺ is known to decrease at higher temperatures (11, 39, 46, 47), suggesting that the 30–40% decrease in tetanic force observed after 30–40 min at 37°C in the presence of 1 mM Tempol is due to SR Ca²⁺ depletion. The gradual recovery in tetanic force to levels similar to those in the control muscles kept at 22°C can be explained by SR Ca²⁺ depletion at 37°C followed by recovery of SR Ca²⁺ content upon return of the muscle to 22°C. Note that some recovery in tetanic force was seen in all muscles after 2 h at 22°C, even in the absence of Tempol, further supporting the view that temperature in the range 22–37°C has effects on excitation-contraction coupling that are not necessarily mediated by O₂^–.

Finally, we wish to point out that the differences in the effect of O₂^– on excitation-contraction coupling between rat and mouse skeletal muscles are not qualitative differences. For example, the maximum Ca²⁺-activated force production is affected in the same way in rat EDL muscle as it is affected in mouse EDL muscle at 37°C, but at the slightly higher temperature of 40°C (48). In the same vein, one would expect that temperature-induced O₂^– production would affect membrane excitability in a similar manner in the mouse fibers as described here for the rat fibers, but the temperature range over which these effects are revealed may differ by several degrees Celsius. Therefore, we believe that the temperature-induced O₂^– effects observed in this study are relevant for all eutherian mammals, including humans, but the precise temperature at which these effects manifest themselves is likely to be slightly different between species.

General Remarks

In conclusion, this study has shown that there is a marked and relatively sudden increase in O₂^– production in mammalian skeletal muscle as the temperature is raised between 32 and 37°C and that this increase in O₂^– production rather than limitations in O₂ delivery to the isolated rat and mouse EDL muscle largely contributes to the irreversible force depression encountered with isolated mammalian skeletal muscle by affecting different steps in the excitation-contraction coupling. The study also showed that Tempol at concentrations around 1 mM can be used as an effective tool to prevent O₂^–- induced effects on isolated mammalian muscles at physiological temperatures. The presence of EC-SOD bound to collagen and basement membrane of muscle cells in vivo, together with the efficient perfusion of the muscle with blood may facilitate the efficient removal of O₂^–, preventing the accumulation of O₂^– and ROS in the muscle above levels that cause a depression in the force response.

	GRANTS

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We thank Australian Research Council and National Health and Medical Research Council (Australia) (D. G. Stephenson), the Lundbeck Foundation (W. A. Macdonald) and the La Trobe University Institute for Advanced Study (C. van der Poel) for financial support.

	FOOTNOTES

 

Address for reprint requests and other correspondence: D. G. Stephenson, Dept. of Zoology, La Trobe Univ., Melbourne, Victoria 3086, Australia (e-mail: george.stephenson{at}latrobe.edu.au)

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.

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