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Muscle Ionic Shifts During Exercise: Implications for Fatigue and Exercise Performance

Morten Hostrup, Simeon Peter Cairns, Jens Bangsbo

10.1002/cphy.c190024

Source: Volume 11, Issue 3, July 2021

Published online: June 2021

Full Article on Wiley Online Library



Abstract

Exercise causes major shifts in multiple ions (e.g., K+, Na+, H+, lactate–, Ca2+, and Cl–) during muscle activity that contributes to development of muscle fatigue. Sarcolemmal processes can be impaired by the trans‐sarcolemmal rundown of ion gradients for K+, Na+, and Ca2+ during fatiguing exercise, while changes in gradients for Cl– and Cl– conductance may exert either protective or detrimental effects on fatigue. Myocellular H+ accumulation may also contribute to fatigue development by lowering glycolytic rate and has been shown to act synergistically with inorganic phosphate (Pi) to compromise cross‐bridge function. In addition, sarcoplasmic reticulum Ca2+ release function is severely affected by fatiguing exercise. Skeletal muscle has a multitude of ion transport systems that counter exercise‐related ionic shifts of which the Na+/K+‐ATPase is of major importance. Metabolic perturbations occurring during exercise can exacerbate trans‐sarcolemmal ionic shifts, in particular for K+ and Cl–, respectively via metabolic regulation of the ATP‐sensitive K+ channel (KATP) and the chloride channel isoform 1 (ClC‐1). Ion transport systems are highly adaptable to exercise training resulting in an enhanced ability to counter ionic disturbances to delay fatigue and improve exercise performance. In this article, we discuss (i) the ionic shifts occurring during exercise, (ii) the role of ion transport systems in skeletal muscle for ionic regulation, (iii) how ionic disturbances affect sarcolemmal processes and muscle fatigue, (iv) how metabolic perturbations exacerbate ionic shifts during exercise, and (v) how pharmacological manipulation and exercise training regulate ion transport systems to influence exercise performance in humans. © 2021 American Physiological Society. Compr Physiol 11:1895‐1959, 2021.

Figure 1. Figure 1. Skeletal muscle contraction is controlled via bursts of action potentials initiated in the motor cortex that descend along neurons of the corticospinal tract to stimulate α‐motor‐neurons in the spinal cord (A). The action potentials propagate along the motor‐neuron axon within a peripheral nerve to ultimately instigate contraction of the muscle fibers that it innervates. The motor‐neuron and its attached muscle fibers are referred to as a motor unit. When an action potential reaches the axon terminal at the neuromuscular junction, the events of neuromuscular transmission follow (B). (1) The neurotransmitter acetylcholine (ACh) is released from the axon terminal into the neuromuscular cleft and (2) binds to nicotinic acetylcholine receptors (AChR) located on the abutting muscle fiber membrane. This opens cationic channels, which allows entry of Na+ ions while K+ ions exit. This leads to a small local depolarization known as the endplate potential. The endplate potential is normally of sufficient magnitude so that the adjacent sarcolemma reaches a depolarization threshold and triggers an action potential, which is propagated in both directions via (3) activation of adjacent voltage‐gated Na+‐channels (Nav1.4). Each muscle action potential involves a rapid depolarization phase, that is, upstroke, underpinned by Na+ influx then a repolarization phase, that is, downstroke, involving K+ efflux via Kv channels combined with 4) the Na+ influx being turned off due to inactivation of Nav1.4.
Figure 2. Figure 2. (A) Muscle contractions are activated by a burst of multiple action potentials (I) that propagate inwardly along the tubular (T‐system) membranes and activate voltage‐sensor proteins (DHPR), which via their interaction with the ryanodine receptor channel complex (RyR1), leads to calcium ions (Ca2+) being released from their storage site in the adjacent sarcoplasmic reticulum (II). This Ca2+ diffuses into the myofilament lattice and binds to troponin‐C, leading to force generation by the contractile apparatus as a result of cross‐bridge cycling (III). Upon cessation of excitation, Ca2+ dissociates from troponin‐C and is transported back into the sarcoplasmic reticulum lumen by the Ca2+ ATPase (SERCA) (IV), whereby the fiber relaxes. (B) The myoplasmic Ca2+ concentration ([Ca2+]) largely determines the force production of muscle fibers, which increases with myoplasmic [Ca2+] in a sigmoidal manner (known as the [Ca2+]‐force relationship). The [Ca2+] needed to produce 50% of maximal fiber force reflects the myofibrillar Ca2+ sensitivity (pCa2+50), which is regulated by intrinsic factors, hormones, and exogenous substances.
Figure 3. Figure 3. Ionic shifts in different compartments (e.g., myoplasm, interstitial fluid, plasma) are most severe during exercise requiring recruitment of many motor units and a substantial contribution from anaerobic metabolism. This figure shows the schematic relationships between exercise intensity, and muscle interstitial concentrations of K+ ([K+]) in relation to exercise intensity relative to leg incremental peak power (A) and systemic plasma concentrations of lactate– ([lactate–]) in relation to exercise intensity of maximal oxygen consumption (V̇o2max) (B).
Figure 4. Figure 4. Contracting muscle fibers extrude K+ during the repolarization phase of each action potential so that K+ efflux becomes substantial during high‐intensity exercise. This causes K+ to accumulate in the interstitial fluid surrounding muscle fibers, which can be measured in vivo with the microdialysis technique. Usually, two to four microdialysis probes are inserted through the muscle of interest with the penetration angle being parallel with fiber angle. The probes are perfused (perfusate) with a known concentration of the ion of interest (e.g., K+) at a constant flow‐rate. Within the probe is a semi‐permeable membrane, which allows for the exchange of ions and molecules before the fluid is sampled at the other end of the probe (dialysate). The figure also illustrates interstitial K+ concentrations [K+] of the vastus lateralis muscle at rest, during one‐legged knee extensor exercise to task failure, and in the first 10 min into recovery from exercise.
Figure 5. Figure 5. The rate of interstitial accumulation and release of K+ are intensity‐dependent. This figure shows the net rate of K+ release from the exercising muscle to the blood (right Y‐axis) and interstitial concentrations of K+ ([K+]) in the vastus lateralis muscle (left Y‐axis) during submaximal (A) or intense (B) knee‐extensor exercise.
Figure 6. Figure 6. The Na+/K+‐ATPase is regulated by multiple factors in human skeletal muscle during exercise. Motor‐neuron release of calcitonin gene‐related peptide (cGRP) and increased circulating levels of epinephrine induced by exercise, stimulate Na+/K+‐ATPase activity via activation of downstream effectors. Protein kinase A (PKA) regulates Na+‐affinity of the Na+/K+‐ATPase via the FXYD1 subunit, while myoplasmic Ca2+ transients occurring during repeated muscle contractions activate protein kinase C (PKC), which in turn stimulates maximal catalytic enzyme activity of the Na+/K+‐ATPase via FXYD1. Both PKA and PKC may induce translocation of additional Na+/K+‐ATPase αβ‐complexes to the sarcolemma, whereas the intracellular energy‐sensor protein, 5′adenosine monophosphate‐activated protein kinase (AMPK), may induce endocytosis or attenuate translocation of Na+/K+‐ATPase αβ‐complexes when cellular energetic state is critically low. Contraction‐induced formation of reactive oxygen species (ROS) can cause glutathionylation of the β‐subunits of the Na+/K+‐ATPase, which compromises its function.
Figure 7. Figure 7. Slow‐inactivation of voltage‐sensitive sodium channels (Nav1.4) has been proposed as a mechanism to reduce sarcolemmal excitability and hence cause muscle fatigue. The voltage dependence of slow‐inactivation (S∞) of Na+ currents in rat fast‐twitch EDL fibers (thin solid line) and slow‐twitch soleus fibers (thin dashed line), and human intercostal fast‐twitch fibers (thick solid line) and slow‐twitch fibers (thick dashed line). Boltzmann curves, fitted to sodium currents, are expressed relative to the maximal sodium currents (INa,max). Slow‐twitch fibers = Type 1 fibers, Fast‐twitch fibers = Type 2 fibers. Temperature, 23 °C. Reused, with permission, from Ruff RL, 1996 610.
Figure 8. Figure 8. This figure shows the effect of raised extracellular K+ concentrations ([K+]) on resting membrane potential in rodent muscle fibers in vitro. Data from (A) rat fast‐twitch EDL fibers exposed to 2 to 100 mM extracellular [K+] and (B) mouse fast‐twitch EDL fibers (□) and slow‐twitch soleus fibers (•) fibers exposed to 4 to 14 mM extracellular [K+] ([K+]o). All data were from surface fibers at 25 °C. Solid lines in A and B were obtained by fitting the GHK equation to the data. The dashed line in A shows the calculated K+ equilibrium potential at each [K+]o. (A) Reused, with permission, from DeCoursey TE, et al., 1981 168 and (B) Reused, with permission, from Cairns SP, et al., 1997 95.
Figure 9. Figure 9. This figure illustrates intracellular recordings of sarcolemmal action potentials with altered extracellular ion concentrations. Effect of raised extracellular K+ concentrations ([K+]) on (A) single action potential profile in rat EDL fibers evoked by current injection, (B) sarcolemmal excitability in rat EDL fibers evoked with current injection at various EM blocks. Percentage of excitable fibers (▪), indeterminate fibers are no longer all‐or‐none ( grey), or inexcitable fibers (□), (C) trains of action potentials (3‐9 mM extracellular [K+]) evoked with transverse wire stimulation in amphibian fibers. (D) Effect of lowered extracellular [Na+] (147‐40 mM) on trains of action potentials in mouse soleus fibers. Stimulation at 50 or 125 Hz (for 2 s) with transverse wire electrode. All recordings were obtained in surface fibers at 22 to 25 °C. (A, B) Reused with permission, from, Rich MM and Pinter MJ, 2003 593 © 2003, John Wiley & Sons. (C) Reused with permission, from, Renaud JM and Light P, 1992 591. © 1992, Canadian Science Publishing; (D) Reused, with permission, from Cairns SP, et al., 2003 88, © 2003, The American Physiological Society.
Figure 10. Figure 10. This figure shows the effect of a reduced trans‐sarcolemmal K+ gradient on peak force in mammalian skeletal muscle preparations in vitro. (A) Peak tetanic force‐extracellular K+ concentration ([K+]) relationships for whole slow‐twitch soleus and fast‐twitch EDL muscles from mice. Tetani were evoked at 125 Hz in soleus, 200 Hz in EDL, with supramaximal stimulus pulses (20 V, 0.1 ms) delivered via parallel plate electrodes, 25 °C. (B) Peak twitch force‐intracellular [K+] relationship for mechanically skinned EDL fibers from rat, 25 °C. The peak force is expressed relative to the maximum level in control solutions. (A) Reused with permission, from, Cairns SP, et al., 1997 95 © 1997, The American Physiological Society. (B) Reused, with permission from, Ørtenblad N and Stephenson DG, 2003 541 © 2003, John Wiley & Sons.
Figure 11. Figure 11. This figure shows the peak tetanic force‐resting membrane potential (EM) relationship for mouse EDL muscle in vitro. Peak tetanic force (200 Hz) was obtained in steady‐state conditions at various extracellular K+ concentrations ([K+]) in whole EDL muscles (obtained from Figure 10) is plotted versus resting EM obtained in surface EDL fibers at the same extracellular [K+] (obtained from Figure 8) at 25 °C. Reused, with permission, from Cairns SP, and Lindinger MI. 2008 100 © 2008, John Wiley & Sons.
Figure 12. Figure 12. Effect of prior‐exposure to altered extracellular ion concentrations on the fatigue profile during repeated intermittent tetanic stimulation in mouse soleus muscle. (A) 2K, 4K, 7K, represent 2, 4, 7 mM extracellular K+ concentrations ([K+]), respectively. (B) 147Na, 80Na represents 147, 80 mM extracellular Na+ concentrations ([Na+]), respectively. Na+ was replaced with choline. (C) 10Ca, 1.3Ca, 0Ca, represents 10, 1.3, 0 mM extracellular Ca2+ concentrations ([Ca2+]), respectively. (D) control (128 mM extracellular Cl– concentration ([Cl−])), low Cl− (10 mM extracellular [Cl−]). Fatigue protocol: 125 Hz for 500 ms, every 1 s, for 100 s. Supramaximal pulses (20 V, 0.1 ms) delivered via parallel plate electrodes, 25 °C. Equilibration in each solution was for at least 40 min. Reproduced from (A) Reused, with permission, from Cairns SP, et al., 1985 99 © 2015, The American Physiological Society. (B) Reused, with permission, from Cairns SP, et al., 2003, 88 © 2003, The American Physiological Society. (C) Reused, with permission, from Cairns SP, et al., 1985 96 © 1998, The American Physiological Society, and (D) Reused, with permission, from Cairns SP, et al., 2004 101 © 2004, The American Physiological Society.
Figure 13. Figure 13. Modulation of sarcolemmal K+ transporters on muscle fatigue profile. (A) Effect of blockade with ouabain (10−5 M) on fatigue during repeated tetani in isolated rat EDL muscle. Fatigue protocol: 60 Hz for 1 s every 4 s, for 3.5 min. Supramaximal pulses (10‐12 V, 0.2 ms) delivered via transverse wire electrodes, 30 °C. Reused, with permission, from Clausen T., 2011 132 © 2011, John Wiley & Sons. (B) Effect of pinacidil, which opens KATP channels; (C) Effect of bumetanide, which inhibits NKCC1; (D) NS‐1619, which opens BKCa channels. (B‐D) Fatigue protocol: Repeated tetanic stimulation of isolated rat soleus muscle at 33 Hz for 1 s every 3 s for 6 min, 26 °C. Reused, with permission, from Kristensen M., et al., 2006 392 © 2006, The American Physiological Society.
Figure 14. Figure 14. Interactive effects of lowered extracellular Na+ concentrations ([Na+]) and raised extracellular K+ concentrations ([K+]) on contraction of rodent muscle in vitro. (A) Effect of lowered extracellular [Na+] at 4 mM extracellular [K+] (○) and 8 mM extracellular [K+] (•) in rat soleus, 30 °C. Peak tetanic force (30 Hz) is expressed relative to the maximum level in the control solution (B). Interactive effects of lowered extracellular [Na+], raised extracellular [K+], and altered extracellur Ca2+ concentrations ([Ca2+]o) on peak tetanic force (125 Hz) in mouse soleus muscle, 25 °C. 8K, 8 mM extracellular [K+]; 100 Na, 100 mM extracellular [Na+], 10Ca, 10 mM extracellular [Ca2+]; 0Ca, Ca2+ free conditions. #Indicates predicted response if effects were additive (8K + 100Na). (A) Reused, with permission, from Overgaard K, et al., 1997 544 © 1997, Springer Nature. (B) Reused, with permission, from Cairns SP and Lindinger MI, 2008 100 © 2008, John Wiley & Sons.
Figure 15. Figure 15. Effect of modulation of Ca2+ entry through sarcolemmal store‐operated Ca2+ channels (TRPC channels) on fatigue kinetics during repeated tetani in mouse soleus muscles. (A) Pharmacological inhibition of TRPC channels with gadolinium (Gd3+), GsMTx‐4, or SKF‐96365, and (B) Genetic modification of TRPC channels. TRPC1−/− knock‐out, TRPC1+/+ control. Fatigue protocol: 50 Hz for 500 ms, every 1 s, for 2 min. (A) Reused, with permission, from Ducret T, et al., 2006 179 © 2006, John Wiley & Sons. (B) Reused, with permission, from Zanou N, et al., 2010 747 © 2010, The American Physiological Society.
Figure 16. Figure 16. Interactive effects of lowered extracellular Cl− concentrations ([Cl−]) with raised extracellular K+ concentrations ([K+]) on tetanic contractions in isolated rodent muscle. (A) Depressive effects of raised extracellular [K+] (4‐9 mM) after pre‐exposure to lowered extracellular [Cl−] (from 127 to 10 mM) for 60 min. (•) 10 mM Cl−, (▪) 9 mM K+ with 127 mM Cl−, (▴) 9 mM K+ with 10 mM Cl−. Tetani were evoked at 125 Hz at 25 °C, in mouse soleus. (B) Restorative effects of lowered extracellular [Cl−] (80 or 0 mM), or 9‐anthracene‐carboxylate (9AC), or 24% CO2, after first raising extracellular [K+] to 11 mM for 60 min on peak tetanic force (solid bars) or M‐wave area (open bars). Tetani were evoked at 30 Hz at 30 °C, in rat soleus. (A) Reused, with permission, from Cairns SP, et al., 2004 101 © 2004, The American Physiological Society. (B) Reused, with permission, from Pedersen TH, et al., 2005 552 © 2005, Rockefeller University Press.
Figure 17. Figure 17. Effects of lactic acid/lactate– on peak force in K+‐depressed rat soleus muscles in vitro. (A) 20 mM lactic acid (open circles) was added after 90 min exposure to 11 mM extracellular [K+]. Tetani (30 Hz) were evoked by direct stimulation with supramaximal 1.0 ms pulses, transverse wire electrodes, 30 °C. (B) 20 mM l‐lactate was added after 90 min exposure to 11 mM extracellular [K+]. Tetani (60 Hz) were evoked by nerve stimulation with supramaximal 0.2 ms pulses, 30 °C. (A) Reused, with permission, from Nielsen OB, et al., 2001 525 © 2001, John Wiley & Sons. (B) Reused, with permission, from de Paoli FV, et al., 2010 163 © 2010, John Wiley & Sons.
Figure 18. Figure 18. Effect of raised levels of lactate– and H+ induced by preceding arm exercise on the rate of leg lactate– (A) and K+ release and muscle interstitial concentrations of K+ [K+] (C) during subsequent knee‐extensor exercise to task failure, as well as on the muscle lactate– concentration and pH of the vastus lateralis (B). “Arm+Leg” denotes knee‐extensor exercise that was preceded by arm exercise (red color), whereas “Leg” denotes a control situation without preceding arm exercise (blue color). Extracted from Bangsbo J, et al., 1996 38 and Nordsborg N, et al., 2003 531.
Figure 19. Figure 19. Mixed venous plasma K+ concentration [K+] during (A) and in recovery (B) from progressive bike ergometer exercise increasing by 50 W every 3 min in trained cyclists (blue) and untrained individuals (red). Adapted from Marcos E, and Ribas J. 1995 458.
Figure 20. Figure 20. Muscle interstitial‐to‐femoral venous K+ concentration [K+] gradient during knee‐extensor exercise to task failure before (red) and after (blue) a period of training in otherwise untrained individuals. Adapted from Nielsen JJ, et al., 2004 522.
Figure 21. Figure 21. Mixed venous blood concentrations of lactate– ([lactate–]) in relation to workload before (red) and after (blue) a period of training in the otherwise untrained individual.
Figure 22. Figure 22. A period of training enhances the ability to counter ionic shifts during exercise for a given intensity. Training reduces and delays exercise‐related accumulation of extracellular K+ and lactate– as well as the influx of Na+ and Cl–. Furthermore, training reduces and delays the rate of decline of myoplasmic pH. These training‐induced changes are attributed to adaptations of the ion transport and H+‐buffer capacity of skeletal muscle. In addition, increments in the capacity of aerobic energy systems with training postpone the reliance on anaerobic metabolism and possibly contribute to the reduced exercise‐related ionic shifts relative to workload. Solid and dashed arrows denote after and before a period of training, respectively.
Figure 23. Figure 23. Training induces several adaptations involved in ion transport of skeletal muscle as shown in this figure. (a) Training upregulates the number of Na+/K+‐ATPase complexes to counter K+ and Na+ disturbances during exercise. (b) Training increases the content of monocarboxylate transporter isoforms 1 and 4 (MCT1/4) and Na+/H+ exchanger isoform 1 (NHE1) to augment the capacity for lactate– and H+ extrusion. A training‐induced increase in the content of H+ acceptors enhances the myoplasmic H+ buffer capacity and attenuates acidosis during intense exercise—in effect delaying or lowering the amount of ATP‐sensitive K+ channels (KATP) being in their open configuration to reduce K+ efflux via these channels. (c) Training enhances the ability of skeletal muscle to match metabolic demands during exercise (i.e., increased metabolic stability) and to resist metabolic perturbations (e.g., H+ accumulation and loss of adenosine nucleotides, such as ATP). Such adaptations may postpone the opening of chloride channel isoform 1 (ClC‐1) and KATP as well as counter reductions of glycolytic rate; the latter of which is important to sustain the activity of the Na+/K+‐ATPase and SERCA. (d) Proteins regulating sarcoplasmic reticulum Ca2+ handling adapt according to the training performed, in which endurance‐based training, such as aerobic training and speed endurance training, may reduce the content of SERCA isoforms and sarcolipin, whereas repeated sprint training may increase ryanodine receptor isoform 1 (RyR1) and SERCA content via an expansion of sarcoplasmic reticulum volume. Furthermore, training makes muscle fibers less susceptible to exercise‐mediated fragmentation of RyR1. Solid green and red arrows indicate an enhancement/upregulation or reduction/downregulation, respectively; dashed green and red arrows indicate a stimulation/activation or inhibition, respectively; dashed black arrows indicate a modulation.
Figure 24. Figure 24. Muscle interstitial (blue circles) and intracellular (red squares) concentrations of K+ ([K+]) during repeated intense exercise. Adapted from Mohr M, et al., 2004 495 and Lindinger MI, et al., 1995 435. Note that performance declines as bouts are repeated.


Figure 1. Skeletal muscle contraction is controlled via bursts of action potentials initiated in the motor cortex that descend along neurons of the corticospinal tract to stimulate α‐motor‐neurons in the spinal cord (A). The action potentials propagate along the motor‐neuron axon within a peripheral nerve to ultimately instigate contraction of the muscle fibers that it innervates. The motor‐neuron and its attached muscle fibers are referred to as a motor unit. When an action potential reaches the axon terminal at the neuromuscular junction, the events of neuromuscular transmission follow (B). (1) The neurotransmitter acetylcholine (ACh) is released from the axon terminal into the neuromuscular cleft and (2) binds to nicotinic acetylcholine receptors (AChR) located on the abutting muscle fiber membrane. This opens cationic channels, which allows entry of Na+ ions while K+ ions exit. This leads to a small local depolarization known as the endplate potential. The endplate potential is normally of sufficient magnitude so that the adjacent sarcolemma reaches a depolarization threshold and triggers an action potential, which is propagated in both directions via (3) activation of adjacent voltage‐gated Na+‐channels (Nav1.4). Each muscle action potential involves a rapid depolarization phase, that is, upstroke, underpinned by Na+ influx then a repolarization phase, that is, downstroke, involving K+ efflux via Kv channels combined with 4) the Na+ influx being turned off due to inactivation of Nav1.4.


Figure 2. (A) Muscle contractions are activated by a burst of multiple action potentials (I) that propagate inwardly along the tubular (T‐system) membranes and activate voltage‐sensor proteins (DHPR), which via their interaction with the ryanodine receptor channel complex (RyR1), leads to calcium ions (Ca2+) being released from their storage site in the adjacent sarcoplasmic reticulum (II). This Ca2+ diffuses into the myofilament lattice and binds to troponin‐C, leading to force generation by the contractile apparatus as a result of cross‐bridge cycling (III). Upon cessation of excitation, Ca2+ dissociates from troponin‐C and is transported back into the sarcoplasmic reticulum lumen by the Ca2+ ATPase (SERCA) (IV), whereby the fiber relaxes. (B) The myoplasmic Ca2+ concentration ([Ca2+]) largely determines the force production of muscle fibers, which increases with myoplasmic [Ca2+] in a sigmoidal manner (known as the [Ca2+]‐force relationship). The [Ca2+] needed to produce 50% of maximal fiber force reflects the myofibrillar Ca2+ sensitivity (pCa2+50), which is regulated by intrinsic factors, hormones, and exogenous substances.


Figure 3. Ionic shifts in different compartments (e.g., myoplasm, interstitial fluid, plasma) are most severe during exercise requiring recruitment of many motor units and a substantial contribution from anaerobic metabolism. This figure shows the schematic relationships between exercise intensity, and muscle interstitial concentrations of K+ ([K+]) in relation to exercise intensity relative to leg incremental peak power (A) and systemic plasma concentrations of lactate– ([lactate–]) in relation to exercise intensity of maximal oxygen consumption (V̇o2max) (B).


Figure 4. Contracting muscle fibers extrude K+ during the repolarization phase of each action potential so that K+ efflux becomes substantial during high‐intensity exercise. This causes K+ to accumulate in the interstitial fluid surrounding muscle fibers, which can be measured in vivo with the microdialysis technique. Usually, two to four microdialysis probes are inserted through the muscle of interest with the penetration angle being parallel with fiber angle. The probes are perfused (perfusate) with a known concentration of the ion of interest (e.g., K+) at a constant flow‐rate. Within the probe is a semi‐permeable membrane, which allows for the exchange of ions and molecules before the fluid is sampled at the other end of the probe (dialysate). The figure also illustrates interstitial K+ concentrations [K+] of the vastus lateralis muscle at rest, during one‐legged knee extensor exercise to task failure, and in the first 10 min into recovery from exercise.


Figure 5. The rate of interstitial accumulation and release of K+ are intensity‐dependent. This figure shows the net rate of K+ release from the exercising muscle to the blood (right Y‐axis) and interstitial concentrations of K+ ([K+]) in the vastus lateralis muscle (left Y‐axis) during submaximal (A) or intense (B) knee‐extensor exercise.


Figure 6. The Na+/K+‐ATPase is regulated by multiple factors in human skeletal muscle during exercise. Motor‐neuron release of calcitonin gene‐related peptide (cGRP) and increased circulating levels of epinephrine induced by exercise, stimulate Na+/K+‐ATPase activity via activation of downstream effectors. Protein kinase A (PKA) regulates Na+‐affinity of the Na+/K+‐ATPase via the FXYD1 subunit, while myoplasmic Ca2+ transients occurring during repeated muscle contractions activate protein kinase C (PKC), which in turn stimulates maximal catalytic enzyme activity of the Na+/K+‐ATPase via FXYD1. Both PKA and PKC may induce translocation of additional Na+/K+‐ATPase αβ‐complexes to the sarcolemma, whereas the intracellular energy‐sensor protein, 5′adenosine monophosphate‐activated protein kinase (AMPK), may induce endocytosis or attenuate translocation of Na+/K+‐ATPase αβ‐complexes when cellular energetic state is critically low. Contraction‐induced formation of reactive oxygen species (ROS) can cause glutathionylation of the β‐subunits of the Na+/K+‐ATPase, which compromises its function.


Figure 7. Slow‐inactivation of voltage‐sensitive sodium channels (Nav1.4) has been proposed as a mechanism to reduce sarcolemmal excitability and hence cause muscle fatigue. The voltage dependence of slow‐inactivation (S∞) of Na+ currents in rat fast‐twitch EDL fibers (thin solid line) and slow‐twitch soleus fibers (thin dashed line), and human intercostal fast‐twitch fibers (thick solid line) and slow‐twitch fibers (thick dashed line). Boltzmann curves, fitted to sodium currents, are expressed relative to the maximal sodium currents (INa,max). Slow‐twitch fibers = Type 1 fibers, Fast‐twitch fibers = Type 2 fibers. Temperature, 23 °C. Reused, with permission, from Ruff RL, 1996 610.


Figure 8. This figure shows the effect of raised extracellular K+ concentrations ([K+]) on resting membrane potential in rodent muscle fibers in vitro. Data from (A) rat fast‐twitch EDL fibers exposed to 2 to 100 mM extracellular [K+] and (B) mouse fast‐twitch EDL fibers (□) and slow‐twitch soleus fibers (•) fibers exposed to 4 to 14 mM extracellular [K+] ([K+]o). All data were from surface fibers at 25 °C. Solid lines in A and B were obtained by fitting the GHK equation to the data. The dashed line in A shows the calculated K+ equilibrium potential at each [K+]o. (A) Reused, with permission, from DeCoursey TE, et al., 1981 168 and (B) Reused, with permission, from Cairns SP, et al., 1997 95.


Figure 9. This figure illustrates intracellular recordings of sarcolemmal action potentials with altered extracellular ion concentrations. Effect of raised extracellular K+ concentrations ([K+]) on (A) single action potential profile in rat EDL fibers evoked by current injection, (B) sarcolemmal excitability in rat EDL fibers evoked with current injection at various EM blocks. Percentage of excitable fibers (▪), indeterminate fibers are no longer all‐or‐none ( grey), or inexcitable fibers (□), (C) trains of action potentials (3‐9 mM extracellular [K+]) evoked with transverse wire stimulation in amphibian fibers. (D) Effect of lowered extracellular [Na+] (147‐40 mM) on trains of action potentials in mouse soleus fibers. Stimulation at 50 or 125 Hz (for 2 s) with transverse wire electrode. All recordings were obtained in surface fibers at 22 to 25 °C. (A, B) Reused with permission, from, Rich MM and Pinter MJ, 2003 593 © 2003, John Wiley & Sons. (C) Reused with permission, from, Renaud JM and Light P, 1992 591. © 1992, Canadian Science Publishing; (D) Reused, with permission, from Cairns SP, et al., 2003 88, © 2003, The American Physiological Society.


Figure 10. This figure shows the effect of a reduced trans‐sarcolemmal K+ gradient on peak force in mammalian skeletal muscle preparations in vitro. (A) Peak tetanic force‐extracellular K+ concentration ([K+]) relationships for whole slow‐twitch soleus and fast‐twitch EDL muscles from mice. Tetani were evoked at 125 Hz in soleus, 200 Hz in EDL, with supramaximal stimulus pulses (20 V, 0.1 ms) delivered via parallel plate electrodes, 25 °C. (B) Peak twitch force‐intracellular [K+] relationship for mechanically skinned EDL fibers from rat, 25 °C. The peak force is expressed relative to the maximum level in control solutions. (A) Reused with permission, from, Cairns SP, et al., 1997 95 © 1997, The American Physiological Society. (B) Reused, with permission from, Ørtenblad N and Stephenson DG, 2003 541 © 2003, John Wiley & Sons.


Figure 11. This figure shows the peak tetanic force‐resting membrane potential (EM) relationship for mouse EDL muscle in vitro. Peak tetanic force (200 Hz) was obtained in steady‐state conditions at various extracellular K+ concentrations ([K+]) in whole EDL muscles (obtained from Figure 10) is plotted versus resting EM obtained in surface EDL fibers at the same extracellular [K+] (obtained from Figure 8) at 25 °C. Reused, with permission, from Cairns SP, and Lindinger MI. 2008 100 © 2008, John Wiley & Sons.


Figure 12. Effect of prior‐exposure to altered extracellular ion concentrations on the fatigue profile during repeated intermittent tetanic stimulation in mouse soleus muscle. (A) 2K, 4K, 7K, represent 2, 4, 7 mM extracellular K+ concentrations ([K+]), respectively. (B) 147Na, 80Na represents 147, 80 mM extracellular Na+ concentrations ([Na+]), respectively. Na+ was replaced with choline. (C) 10Ca, 1.3Ca, 0Ca, represents 10, 1.3, 0 mM extracellular Ca2+ concentrations ([Ca2+]), respectively. (D) control (128 mM extracellular Cl– concentration ([Cl−])), low Cl− (10 mM extracellular [Cl−]). Fatigue protocol: 125 Hz for 500 ms, every 1 s, for 100 s. Supramaximal pulses (20 V, 0.1 ms) delivered via parallel plate electrodes, 25 °C. Equilibration in each solution was for at least 40 min. Reproduced from (A) Reused, with permission, from Cairns SP, et al., 1985 99 © 2015, The American Physiological Society. (B) Reused, with permission, from Cairns SP, et al., 2003, 88 © 2003, The American Physiological Society. (C) Reused, with permission, from Cairns SP, et al., 1985 96 © 1998, The American Physiological Society, and (D) Reused, with permission, from Cairns SP, et al., 2004 101 © 2004, The American Physiological Society.


Figure 13. Modulation of sarcolemmal K+ transporters on muscle fatigue profile. (A) Effect of blockade with ouabain (10−5 M) on fatigue during repeated tetani in isolated rat EDL muscle. Fatigue protocol: 60 Hz for 1 s every 4 s, for 3.5 min. Supramaximal pulses (10‐12 V, 0.2 ms) delivered via transverse wire electrodes, 30 °C. Reused, with permission, from Clausen T., 2011 132 © 2011, John Wiley & Sons. (B) Effect of pinacidil, which opens KATP channels; (C) Effect of bumetanide, which inhibits NKCC1; (D) NS‐1619, which opens BKCa channels. (B‐D) Fatigue protocol: Repeated tetanic stimulation of isolated rat soleus muscle at 33 Hz for 1 s every 3 s for 6 min, 26 °C. Reused, with permission, from Kristensen M., et al., 2006 392 © 2006, The American Physiological Society.


Figure 14. Interactive effects of lowered extracellular Na+ concentrations ([Na+]) and raised extracellular K+ concentrations ([K+]) on contraction of rodent muscle in vitro. (A) Effect of lowered extracellular [Na+] at 4 mM extracellular [K+] (○) and 8 mM extracellular [K+] (•) in rat soleus, 30 °C. Peak tetanic force (30 Hz) is expressed relative to the maximum level in the control solution (B). Interactive effects of lowered extracellular [Na+], raised extracellular [K+], and altered extracellur Ca2+ concentrations ([Ca2+]o) on peak tetanic force (125 Hz) in mouse soleus muscle, 25 °C. 8K, 8 mM extracellular [K+]; 100 Na, 100 mM extracellular [Na+], 10Ca, 10 mM extracellular [Ca2+]; 0Ca, Ca2+ free conditions. #Indicates predicted response if effects were additive (8K + 100Na). (A) Reused, with permission, from Overgaard K, et al., 1997 544 © 1997, Springer Nature. (B) Reused, with permission, from Cairns SP and Lindinger MI, 2008 100 © 2008, John Wiley & Sons.


Figure 15. Effect of modulation of Ca2+ entry through sarcolemmal store‐operated Ca2+ channels (TRPC channels) on fatigue kinetics during repeated tetani in mouse soleus muscles. (A) Pharmacological inhibition of TRPC channels with gadolinium (Gd3+), GsMTx‐4, or SKF‐96365, and (B) Genetic modification of TRPC channels. TRPC1−/− knock‐out, TRPC1+/+ control. Fatigue protocol: 50 Hz for 500 ms, every 1 s, for 2 min. (A) Reused, with permission, from Ducret T, et al., 2006 179 © 2006, John Wiley & Sons. (B) Reused, with permission, from Zanou N, et al., 2010 747 © 2010, The American Physiological Society.


Figure 16. Interactive effects of lowered extracellular Cl− concentrations ([Cl−]) with raised extracellular K+ concentrations ([K+]) on tetanic contractions in isolated rodent muscle. (A) Depressive effects of raised extracellular [K+] (4‐9 mM) after pre‐exposure to lowered extracellular [Cl−] (from 127 to 10 mM) for 60 min. (•) 10 mM Cl−, (▪) 9 mM K+ with 127 mM Cl−, (▴) 9 mM K+ with 10 mM Cl−. Tetani were evoked at 125 Hz at 25 °C, in mouse soleus. (B) Restorative effects of lowered extracellular [Cl−] (80 or 0 mM), or 9‐anthracene‐carboxylate (9AC), or 24% CO2, after first raising extracellular [K+] to 11 mM for 60 min on peak tetanic force (solid bars) or M‐wave area (open bars). Tetani were evoked at 30 Hz at 30 °C, in rat soleus. (A) Reused, with permission, from Cairns SP, et al., 2004 101 © 2004, The American Physiological Society. (B) Reused, with permission, from Pedersen TH, et al., 2005 552 © 2005, Rockefeller University Press.


Figure 17. Effects of lactic acid/lactate– on peak force in K+‐depressed rat soleus muscles in vitro. (A) 20 mM lactic acid (open circles) was added after 90 min exposure to 11 mM extracellular [K+]. Tetani (30 Hz) were evoked by direct stimulation with supramaximal 1.0 ms pulses, transverse wire electrodes, 30 °C. (B) 20 mM l‐lactate was added after 90 min exposure to 11 mM extracellular [K+]. Tetani (60 Hz) were evoked by nerve stimulation with supramaximal 0.2 ms pulses, 30 °C. (A) Reused, with permission, from Nielsen OB, et al., 2001 525 © 2001, John Wiley & Sons. (B) Reused, with permission, from de Paoli FV, et al., 2010 163 © 2010, John Wiley & Sons.


Figure 18. Effect of raised levels of lactate– and H+ induced by preceding arm exercise on the rate of leg lactate– (A) and K+ release and muscle interstitial concentrations of K+ [K+] (C) during subsequent knee‐extensor exercise to task failure, as well as on the muscle lactate– concentration and pH of the vastus lateralis (B). “Arm+Leg” denotes knee‐extensor exercise that was preceded by arm exercise (red color), whereas “Leg” denotes a control situation without preceding arm exercise (blue color). Extracted from Bangsbo J, et al., 1996 38 and Nordsborg N, et al., 2003 531.


Figure 19. Mixed venous plasma K+ concentration [K+] during (A) and in recovery (B) from progressive bike ergometer exercise increasing by 50 W every 3 min in trained cyclists (blue) and untrained individuals (red). Adapted from Marcos E, and Ribas J. 1995 458.


Figure 20. Muscle interstitial‐to‐femoral venous K+ concentration [K+] gradient during knee‐extensor exercise to task failure before (red) and after (blue) a period of training in otherwise untrained individuals. Adapted from Nielsen JJ, et al., 2004 522.


Figure 21. Mixed venous blood concentrations of lactate– ([lactate–]) in relation to workload before (red) and after (blue) a period of training in the otherwise untrained individual.


Figure 22. A period of training enhances the ability to counter ionic shifts during exercise for a given intensity. Training reduces and delays exercise‐related accumulation of extracellular K+ and lactate– as well as the influx of Na+ and Cl–. Furthermore, training reduces and delays the rate of decline of myoplasmic pH. These training‐induced changes are attributed to adaptations of the ion transport and H+‐buffer capacity of skeletal muscle. In addition, increments in the capacity of aerobic energy systems with training postpone the reliance on anaerobic metabolism and possibly contribute to the reduced exercise‐related ionic shifts relative to workload. Solid and dashed arrows denote after and before a period of training, respectively.


Figure 23. Training induces several adaptations involved in ion transport of skeletal muscle as shown in this figure. (a) Training upregulates the number of Na+/K+‐ATPase complexes to counter K+ and Na+ disturbances during exercise. (b) Training increases the content of monocarboxylate transporter isoforms 1 and 4 (MCT1/4) and Na+/H+ exchanger isoform 1 (NHE1) to augment the capacity for lactate– and H+ extrusion. A training‐induced increase in the content of H+ acceptors enhances the myoplasmic H+ buffer capacity and attenuates acidosis during intense exercise—in effect delaying or lowering the amount of ATP‐sensitive K+ channels (KATP) being in their open configuration to reduce K+ efflux via these channels. (c) Training enhances the ability of skeletal muscle to match metabolic demands during exercise (i.e., increased metabolic stability) and to resist metabolic perturbations (e.g., H+ accumulation and loss of adenosine nucleotides, such as ATP). Such adaptations may postpone the opening of chloride channel isoform 1 (ClC‐1) and KATP as well as counter reductions of glycolytic rate; the latter of which is important to sustain the activity of the Na+/K+‐ATPase and SERCA. (d) Proteins regulating sarcoplasmic reticulum Ca2+ handling adapt according to the training performed, in which endurance‐based training, such as aerobic training and speed endurance training, may reduce the content of SERCA isoforms and sarcolipin, whereas repeated sprint training may increase ryanodine receptor isoform 1 (RyR1) and SERCA content via an expansion of sarcoplasmic reticulum volume. Furthermore, training makes muscle fibers less susceptible to exercise‐mediated fragmentation of RyR1. Solid green and red arrows indicate an enhancement/upregulation or reduction/downregulation, respectively; dashed green and red arrows indicate a stimulation/activation or inhibition, respectively; dashed black arrows indicate a modulation.


Figure 24. Muscle interstitial (blue circles) and intracellular (red squares) concentrations of K+ ([K+]) during repeated intense exercise. Adapted from Mohr M, et al., 2004 495 and Lindinger MI, et al., 1995 435. Note that performance declines as bouts are repeated.
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Article Title: Muscle Ionic Shifts During Exercise: Implications for Fatigue and Exercise Performance
 

How to Cite

Morten Hostrup, Simeon Peter Cairns, Jens Bangsbo. Muscle Ionic Shifts During Exercise: Implications for Fatigue and Exercise Performance. Compr Physiol 2021, 11: 1895-1959. doi: 10.1002/cphy.c190024