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. 2018 Nov 20;597(1):225–235. doi: 10.1113/JP276528

Elevation of extracellular osmolarity improves signs of myotonia congenita in vitro: a preclinical animal study

Kerstin Hoppe 1, Sunisa Chaiklieng 2,3, Frank Lehmann‐Horn 2, Karin Jurkat‐Rott 5, Scott Wearing 4, Werner Klingler 4,5,6,✉
PMCID: PMC6312412  PMID: 30284249

Abstract

Key points

  • During myotonia congenita, reduced chloride (Cl−) conductance results in impaired muscle relaxation and increased muscle stiffness after forceful voluntary contraction.

  • Repetitive contraction of myotonic muscle decreases or even abolishes myotonic muscle stiffness, a phenomenon called ‘warm up’.

  • Pharmacological inhibition of low Cl− channels by anthracene‐9‐carboxylic acid in muscle from mice and ADR (‘arrested development of righting response’) muscle from mice showed a relaxation deficit under physiological conditions compared to wild‐type muscle.

  • At increased osmolarity up to 400 mosmol L–1, the relaxation deficit of myotonic muscle almost reached that of control muscle.

  • These effects were mediated by the cation and anion cotransporter, NKCC1, and anti‐myotonic effects of hypertonicity were at least partly antagonized by the application of bumetanide.

Abstract

Low chloride‐conductance myotonia is caused by mutations in the skeletal muscle chloride (Cl−) channel gene type 1 (CLCN1). Reduced Cl− conductance of the mutated channels results in impaired muscle relaxation and increased muscle stiffness after forceful voluntary contraction. Exercise decreases muscle stiffness, a phenomena called ‘warm up’. To gain further insight into the patho‐mechanism of impaired muscle stiffness and the warm‐up phenomenon, we characterized the effects of increased osmolarity on myotonic function. Functional force and membrane potential measurements were performed on muscle specimens of ADR (‘arrested development of righting response’) mice (an animal model for low gCl– conductance myotonia) and pharmacologically‐induced myotonia. Specimens were exposed to solutions of increasing osmolarity at the same time as force and membrane potentials were monitored. In the second set of experiments, ADR muscle and pharmacologically‐induced myotonic muscle were exposed to an antagonist of NKCC1. Upon osmotic stress, ADR muscle was depolarized to a lesser extent than control wild‐type muscle. High osmolarity diminished myotonia and facilitated the warm‐up phenomenon as depicted by a faster muscle relaxation time (T 90/10). Osmotic stress primarily resulted in the activation of the NKCC1. The inhibition of NKCC1 with bumetanide prevented the depolarization and reversed the anti‐myotonic effect of high osmolarity. Increased osmolarity decreased signs of myotonia and facilitated the warm‐up phenomenon in different in vitro models of myotonia. Activation of NKCC1 activity promotes warm‐up and reduces the number of contractions required to achieve normal relaxation kinetics.

Keywords: Myotonia congenita, warm‐up phenomena, NKCC1

Key points

  • During myotonia congenita, reduced chloride (Cl−) conductance results in impaired muscle relaxation and increased muscle stiffness after forceful voluntary contraction.

  • Repetitive contraction of myotonic muscle decreases or even abolishes myotonic muscle stiffness, a phenomenon called ‘warm up’.

  • Pharmacological inhibition of low Cl− channels by anthracene‐9‐carboxylic acid in muscle from mice and ADR (‘arrested development of righting response’) muscle from mice showed a relaxation deficit under physiological conditions compared to wild‐type muscle.

  • At increased osmolarity up to 400 mosmol L–1, the relaxation deficit of myotonic muscle almost reached that of control muscle.

  • These effects were mediated by the cation and anion cotransporter, NKCC1, and anti‐myotonic effects of hypertonicity were at least partly antagonized by the application of bumetanide.

Introduction

Myotonia congenita is caused by mutations in the skeletal muscle chloride‐channel gene type 1 (CLCN1) and can be inherited either as an autosomal dominant (Thomsen's myotonia) or autosomal recessive (Becker's myotonia) trait (Lehmann‐Horn et al. 2004; Trivedi et al. 2014). Close to 130 different mutations are currently known, which might at least partly explain the variation of clinical phenotypes (Lossin & George, 2008). Otherwise, identical mutations can also cause a wide‐ranging and heterogeneous spectrum of phenotypes (Colding‐Jorgensen, 2005). Moreover, another mutation within the sodium voltage‐gated channel alpha subunit (SCN4A) gene was detected in clinically diagnosed myotonia congenita (Heine et al. 1993). Hence, mutations in CLCN1 and SCN4A can result in clinically indistinguishable myotonia.

Chloride (Cl−) conductance generates 70–80% of the resting muscle membrane conductance, stabilizing the resting membrane potential near the chloride equilibrium potetenial (Adrian et al. 1956; Adrian & Freygang, 1962; Byrant & Morales‐Aguilera, 1971; Steinmeyer et al. 1991). In Thomsen's and Becker's myotonia, however, decreased chloride conductance impairs electrical stability, with less current being required to trigger an action potential, resulting in membrane hyper‐excitability (Adrian & Bryant, 1974). Moreover, re‐polarization is impaired and the re‐excitation process is prevented (Adrian & Marshall, 1976; Skov et al. 2013). An experimental decrease of Cl− conductance to 20% is associated with a clear myotonic muscle behaviour, whereas a reduction to 50% does not cause myotonia (Furmann & Barchi, 1978).

Chloride conductance is important for countering the depolarizing effect of potassium accumulation in the transverse tubular system (T‐system) and for volume control of the T‐tubule (Gosmanov et al. 2002; Kristensen et al. 2006; Kristensen & Jeul, 2010). The T‐tubule is characterized by a long (∼20–30 μm in length) narrow shape and a tight opening (∼10–50 nm in diameter) on the sarcoplasmic surface. It consists of a regular arrangement along the myotubes and forms triads inside the cell in conjunction with the sarcoplasmic reticulum. These anatomic properties hinder the diffusion of potassium ions between the inner and outer extracellular space, resulting in a heightened potassium concentration in the T‐system during muscle activation (Almers, 1972; Adrian & Peachey, 1973). When exposed to hypertonicity, skeletal muscle fibres shrink to a new steady‐state volume (Blinks, 1965; Chinet, 1993). Assuming that chloride distribution is in equilibrium, cell shrinkage might cause hyperpolarization as a result of an increase in the intracellular cation concentration and higher permeability for potassium compared to sodium (van Mil et al. 1997). Indeed, hyperpolarization was detected in frog toe muscle fibres with exposure to increasing medium osmolarity (Gordon & Godt, 1970).

The ‘warm‐up’ phenomenon is a conspicuous and use‐dependent feature of low chloride‐conductance myotonia in which continued muscle activity reduces or even abolishes myotonic stiffness. Although well described clinically, the mechanism underlying the warm‐up phenomenon is currently unexplained (Novak et al. 2015). The cation and anion cotransporter, NKCC1, is widely expressed in skeletal muscle, particularly in the sarcolemmal membrane and T‐system. NKCC1 is activated by hyperosmolarity resulting mainly from K+ efflux and lactate production during muscle contraction (Delpire et al. 1994). In resting skeletal muscle, it accounts for 15% of K+ uptake and 23% of Na+ uptake and for >30% of K+ transport during muscle stimulation with either catecholamines or electrical stimulation (Lindinger et al. 2002). The chloride equilibrium potential in skeletal muscle is essentially set by the passive electrochemical equilibrium of chloride in accordance with the resting membrane potential, which in turn is mainly determined by the potassium equilibrium potential (Tang & Chen, 2011). However, there is emerging evidence to support the notion that, as a result of chloride import, NKCC1 might also influence the membrane potiential (Geukes Foppen et al. 2002; Geukes Foppen, 2004; Gallaher et al. 2009). Therefore, the present study aimed to clarify the role of increased osmolarity and the role of NKCC1 in different models of myotonia congenita.

Methods

Ethical approval

All experiments with mice were perfomed in vitro after death by cervical dislocation after CO2 narcosis, with a concentration of 35% within the transparent cage, for at least 2 min (Moody et al. 2014; Boivin et al. 2016). All animal experiments were carried out in accordance with the guidelines set out by the welfare committee of the University of Ulm (approval reference 37/97) and conformed with the principles and regulations described by Grundy, 2015.

Mice

Myotonic ADR and wild‐type (WT) mice were used as an animal model for low choloride‐conductance myotonia. To obtain ADR (adr/adr) mice, Balb/c × adr/+ females were mated with Balb/c × adr/+ males and raised in a pathogen‐free animal facility. The adr allele was verified using polymerase chain reaction analysis. The homozygous (adr/adr) offspring were distinguished from heterozygous mice (adr/+) by their myotonic phenotype, which was manifest from day 7 onward. All experiments were undertaken in agreement with the regulations of the local animal welfare committee (Ulm University).

Genetic screening

For PCR analysis, DNA was extracted from homogenized mouse tails tips (4–6 mm) and purified using a QIAamp DNA Mini Kit (Qiagen, Hilden, Germany) in accordance with the manufacturer´s protocol. PCR amplification was performed using 40–80 ng of genomic DNA in a final volume of 50 μL containing a mixture of: 1 μL of DNA (40–80 ng); 41.3 μL of distilled water; 5 μL of 10 × PCR buffer (15 mm MgCl2; Qiagen); 1 μL of primer ADR‐forward (50 pmol μL–1; Interactiva, Ulm, Germany); 0.5 μL of dNTP‐mixture (dGTP, dATP, dTTP, dCTP) (10 mm; Sigma‐Aldrich, Steinheim, Germany); 0.2 μL of Taq‐DNA‐polymerase = 1 U (Qiagen). For detection of the adr allele, genomic DNA was amplified using (from 5′‐ to 3′) CTG TCC AAC CTA AAC TCT CAA GC as forward primer and TCC TAC CGC ATC CTC AGC AA as reverse primer. PCR amplification was performed using a Techne Progene Thermocycler (Thermodux, Wertheim, Germany).

Double‐stranded DNA fragments were separated for agarose gel electrophoresis. The 1.8% agarose (Biozym, Oldendorf, Germany) was heated (100°C for 2 min) in 1 m 10 × TBE Butffer (Tris‐Borate 45 mm; EDTA 1 mm, pH 8.3; Merk, Darmstadt, Germany). After cooling to 50–55°C, 8 μL of 0.7 μg mL–1 of ethidiumbromide (Eurobio, Lesulis, France) was added and a horizontal gel was prepared in a mini‐chamber (Peplab, Erlangen, Germany). DNA probes were loaded with 1:5 final volume 6 × loading buffer (15% Ficoll, 0.1% Bromphenolblau; Merk). As a standard DNA marker, 10 ng from a 100 bp or a 1 kb marker (Eurogentec, Karlsruhe, Germany) was used. The conditions used for gel electrophoresis were 100 V at 50°C for 1 h. Nucleic acids were detected by UV illumination and the results were documented photographically. The presence of the adr allele resulted in a PCR product of ∼600 bp (calculated: 594 bp) for both homozygous (adr/adr) and heterozygous animals (adr/+), whereas no band was seen from the wild type (+/+) allele.

Induction of pharmacologically‐induced myotonia

Induction of myotonia was achieved via inhibition of low chloride channels by anthracene‐9‐carboxylic acid (9‐AC). 9‐AC is a ClC‐1 blocker and myotonia inducing effects were confirmed both in vitro and in vivo (Byrant & Morales‐Aguilera, 1971; Desaphy et al. 2013). Previous work revealed that 50 μm 9‐AC reduced gCl– by >90%. As a stock solution, 9‐AC was dissolved in DMSO, which itself causes an increase in osmolarity. For this reason, the optimum concentration of stock solution was prepared at 500 times the desired end concentration (i.e. 50 mm in 80% DMSO). The osmolarity of the final solution was checked and the pH was equilibrated to 7.4. 9‐AC stock solution was added to the solution to yield an end concentration of 100 μm.

Muscle dissection and preparation

ADR and WT mice, aged between 45–70 days, were killed by cervical dislocation after narcosis with CO2 for at least 2 min. The hind limbs were freely dissected and fixed on a thin layer of Sylgard (Dow Corning, Belgium) following removal of the skin. The gastrocnemius was subsequently dissected and used for force measurement. Hemidiaphragms were also harvested from the central tendon to ribs and sectioned parallel to the muscle fibres to form samples that were ∼7 mm in width. The hemidiaphragm was used for resting membrane potential (RMP) measurements because it is easily impaled. The hemidiaphragms of WT and ADR mice were placed in a chamber filled with 4.5 mm [K+]0 KR solution for internal microelectrode measurements.

Myographic registrations

Murine muscle samples were mounted in an organ bath and continuously bubbled with carbogen (95% O2, 5% CO2; MTI IndustrieGASE, Neu‐Ulm, Germany). For muscle contraction experiments, physiological Krebs‐Ringer solution was used comprising (mm): 118 NaCl, 0.8 MgSO4, 1.0 KH2PO4, 11.1 glucose, 25 NaHCO3 and 2.5 CaCl2. One set of experiments was conducted using Bretag comprising (mm): 107.7 NaCl, 0.69 MgSO4, 1.67 NaH2PO4, 5.05 glucose, 26.2 NaHCO3, 1.53 CaCL2, 9.63 Na‐gluconate and 7.6 saccarose. KCl concentrations were adjusted in the range 0–10 mm. pH was set at 7.4. The temperature was set to 25°C for murine samples. Each muscle specimen was attached to a highly sensitive force transducer (Model FT03; Glass Instruments, Quincy, MA, USA) coupled with a bridge amplifier and an analogue‐digital board (Digidata 1200B; Axon Instruments, Union City, CA, USA). Force signals were recorded using custom computer software (Delphi 1.0; Borland International, Scotts Valley, CA, USA). A pair of platinum electrodes was placed on the lateral parts of the muscle for electrical stimulation with supramaximal stimuli (25 V, 1 ms). To ensure optimum force development, muscle bundles were pre‐stretched to ∼150% of initial length. All muscle samples were allowed to equilibrate in the chamber solution for at least 15 min prior to measurement. For electrophysiological experiments, the test agents were added to the organ bath.

For single twitch experiments, muscle samples were stimulated at a frequency of 0.1 Hz. For each sample, a set of 20 successive twitches were recorded. The contraction and relaxation parameters evaluated were: twitch tension, defined as the peak contractile force (mN); time to peak contractile force (t peak), measured from the beginning of the pulse until the twitch reached maximum amplitude (ms); time to peak half‐life (t 1/2), determined as the time for the force to decrease to half of t peak value (ms); and time from 90% to 10% of t peak (t 90/10), defined as the time between 90% of t peak and the time when the force had decreased to its 10% value (ms).

In some experiments, the influence of Cl− on the resting potential and myotonic effects were removed. Therefore, Cl− was replaced by non‐permeant methanesulphonate instead of chloride. The Cl−‐free solution always contained (mm): 107.7 Na‐methanesulphonate, 0.69 MgSO4, 1.67 NaH2PO4, 26.2 NaHCO3 and 9.63 Ca‐gluconate. pH was set at 7.4.

Internal microelectrode measurements

Microelectrodes were formed on a horizontal two‐stage puller (DMZ Universal Puller; Zeitz Instruments, Munich, Germany) from thin‐wall borosilicate glass pipettes (outer diameter 1.5 mm, inner diameter 1.05 mm; Science Products, Hofheim, Germany). The electrodes were tapered to minimize access resistance and cell damage. The central bore of the electrodes was filled with 3 m KCl and had a resistance of 10 ± 3MΩ. A silver wire (length 5 cm, diameter 300 μm; Science Products) was chemically chlorated in Fe2Cl3 solution to obtain Ag/AgCl electrodes. The recording chamber was placed under a microscope (SMZ‐U stereomicroscope; Nikon Dornach, Germany). Head‐stages and pipette holders were fixed using manipulators (Leitz, Wetzlar, Germany). Membrane potentials were amplified (Axopatch 200B; Axon Instruments) and digitized (Digidata 1200 AD/DA converter; Axon Instruments) using Fetches 6.0 pClamp acquisition software (Axon Instruments). Proprietary software (pClamp 8.0; Axon Instruments) was used to determine the membrane potential in response to pharmacological exposure.

Membrane potential measurements

A 7 mm wide hemidiaphragm was mounted in a chamber containing iso‐osmolar 4.5 [K+]0 KR solution. The bath solution was buffered to pH 7.4 by bubbling with carbon (95% O2, 5%CO2). The small‐volume chamber of 500 μL allowed rapid exchange of the chamber solution. The chamber solution was changed in accordance with the protocol of the intervalence charge transfer experiments (Usher‐Smith et al. 2007). To block repetitive AP bursts from myotonic muscle, 500 nm TTX (Latoxan, Valence, France), a Nav1.4 channel blocker, was added to the chamber solution. After equilibration for 15 min, the glass microelectrodes were inserted into the muscle sample. An Ag/AgCl pellet was used for the bath electrode. RMP was continuously recorded in parallel with stepwise penetration of the muscle by the glass electrode.

Statistical analysis

All data are presented as the mean ± SD. The significance of differences between groups were evaluated using a Wilcoxon matched pairs signed‐rank test. P < 0.05 was considered statistically significant.

Results

Influence of osmolarity on myotonic acitivity

Experiments were performed under hyper‐ and hypo‐ osmotic conditions. 9‐AC, Cl−‐free and ADR muscle all showed a relaxation deficit (t 90/10) at physiological conditions (300 mosmol L–1) compared to WT. Under hypo‐osmotic conditions (250 mosmol L–1), the relaxation deficit (T90/10) was more than 100 times longer in ADR muscle than in WT. As shown in Fig. 1, elevation of extracellular osmolarity shortened t 90/10 in all models of myotonia. At 400 mosmol L–1, the relaxation time of myotonic muscle almost reached that of control (WT) muscle (Figs 1 and 2).

Figure 1. Example original mechanographic registrations of gastrocnemius muscle.

Figure 1

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A, representative original registration of WT muscle. B, ADR muscle showing ‘warm up’. The strips were electrically stimulated with 1 ms pulses at 0.1 Hz after 20 min incubation and 300 mosmol L–1. C, representative registration of ADR muscle in 500 mosmol L–1 showing ‘absent myotonia’.

Figure 2. Anti‐myotonic effects of high osmolarity.

Figure 2

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A, average relaxation time (t 90/10) of 20 twitches in WT (n = 20), ADR (n = 13), 9‐AC (n = 15) and Cl− free (n = 17) muscle were shown. Increasing extracellular osmolarity siginificantly decreased t 90/10 in all models of myotonia. +Significant difference vs. control. *Significant difference vs. 300 mosmol L–1. B, relative peak force (t force) in WT (n = 20), ADR (n = 13), 9‐AC (n = 15) and Cl− free (n = 17) muscle are shown. Increasing extracellular osmolarity significantly reduced t force in the WT, 9‐AC and the Cl−free muscle. +Significant difference vs. control. *Significant difference vs. 300 mosmol L–1. Supplementary preliminary data at 340 mosmol L–1 (n = 3) in WT and ADR are also shown.

Twitch force of myotonic muscle was diminished with increasing extracellular osmolarity. There was a significant reduction in twitch force in 9‐AC and Cl−‐free muscle starting at 350 mosmol L–1. The mean values compared to the physiological osmolarity were 67.0 ± 0.7 in 350 mosmol L–1 vs. 79 ± 5.2 mN in 400 mosmol L–1 and 49.4 ± 5.0 mN in 350 mosmol L–1 vs. 66.1 ± 6.2 mN in 400 mosmol L–1, respectively.

Facilitation of the warm‐up phenomenon by high osmolarity

High osmolarity also influenced the warm‐up phenomenon. From 20 consecutive twitches, t 90/10 of WT muscle remained stable over a series of contractions. Contractions of 9‐AC, Cl−‐free and ADR muscle showed relaxation deficits in several first twitches at physiological osmolarity. The full warm‐up twitch occurred after 20 twitches in 9‐AC and C−‐free solution and at 13.0 ± 3.7 twitches in ADR. The strong myotonic activity in ADR was clearly illustrated by an increase in t 90/10 by a factor of 500 in some twitches. Elevation of osmolarity caused an earlier onset of the warm‐up phenomenon, which was most pronounced in the 9‐AC model. However, the Cl−‐free and the ADR models did not clearly reveal enhancement of the warm‐up. Instead, the results suggest that, once a threshold is exceeded, myotonia is absent in these two models (Fig. 3). At 400 mOsm, the full warm‐up phenomenon was reached at twitch 1.17 ± 0.11, 5.14 ± 1.01 and 3.0 ± 0.53 in Cl−‐free, 9‐AC and ADR muscle. Hence, elevated osmolarity facilitates the warm‐up phenomenon in low gCl− myotonia.

Figure 3. Influence of osmolarity on the warm‐up phenomenon.

Figure 3

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During a succession of single twitch contractions, the strips were electrically stimulated with 1 ms pulses at 0.1 Hz after 20 min of incubation in 250, 300, 350, 400 and 550 mosmol L–1. Data were obtained from the first 15 twitches of contraction of Cl−‐free (n = 30) (A), 9‐AC (n = 15) (B) and ADR‐muscle (n = 13) (C). High osmolarity facilitates the warm‐up in the 9‐AC model.

Influence of osmolarity on RMP and proportion of myotonic fibres

Elevation of osmolarity resulted in a significant depolarization of the RMP in WT and ADR mice tissue (Fig. 4). Membrane depolarization of WT was increased compared to ADR; however, this effect was not statistically significant. Hyperosmolaritiy significantly reduced the proportion of myotonic fibres in ADR mice.

Figure 4. Influence of osmolarity on RMP and proportion with myotonia.

Figure 4

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The RMP was obtained from ADR hemidiaphragm as mean values of 25–30 fibres per animal at 250 mosmol L–1 (n = 13), 290 mosmol L–1 (n = 17), 330 mosmol L–1 (n = 10), 360 mosmol L–1 (n = 11), 395 mosmol L–1 (n = 14) and 430 and 500 mosmol L–1 (n = 6). The proportion with myotonia are illustrated as the percentage of muscle fibres from 75 at 250 mosmol L–1, from 210 at 290 mosmol L–1, from 75 at 360 mosmol L–1 and from 70 at 430 and 500 mosmol L–1. *Significantly different vs. 250 mosmol L–1.

Effects of NKCC1 in low gCl− myotonia

Under hyperosmotic conditions, the cation cotransporter NKCC1 is activated and is a key mediator of volume regulation. Experiments were conducted using bumetanide, a potent NKCC1 inhibitor. In all models of myotonia, the anti‐myotonic effects of hypertonicity were at least partly antagonized by application of bumetanide (Fig. 5). In ADR muscle, bumetanide increased t 90/10 to values comparable to those of physiological osmolarity. The average twitch forces at 400 mosmol L–1 with and without bumetanide to 9‐AC, Cl−‐free and ADR muscle were 70.7 ± 13.6 mN vs. 39.0 ± 4.3 mN (n = 8), 37.1 ± 5.4 mN vs. 29.7 ± 4.3 mN (n = 7) and 35.5 ± 5.8 mN vs. 15.4 ± 1.8 mN (n = 9), respectively.

Figure 5. Anti‐myotonic effects of high osmolarity are partially antagonized by the specific NKCC1 inihibitor bumetanide.

Figure 5

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Data represent mean values of 20 single twitches of Cl–‐free (n = 7), 9‐AC (n = 8) and ADR (n = 10) gastrocnemius muscle. Application of 100 μm bumetanide significantly reversed the anti‐myotonic effects. [Color figure can be viewed at wileyonlinelibrary.com]

NKCC1 inhibition under hyperosmotic conditions

Membrane depolarization and a reduction of a proportion of fibres with myotonia was observed under hyperosmotic conditions. To confirm that NKCC1 plays an important role on the anti‐myotonic effect, the RMP was recorded in ADR hemidiaphragm after 15 min of exposure to bumetanide under hyperosmotic conditions. Figure 6 demonstrates the RMP under hyperosmotic conditions in ADR fibres. After exposure to bumetanide, the membrane depolarization was significantly suppressed at 350, 395, 420 and 500 mosmol L–1. These results indicate that NKCC1 activation plays an important role on membrane depolarization in hypertonicity.

Figure 6. Inhibition of membrane depolarization in hyperosmotic solution by bumetanide.

Figure 6

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Increasing osmolarity caused membrane depolarization in ADR hemidiaphragm. Addition of 50 μm bumetanide to ADR‐muscle (ADR_bumetanide, n = 8) significantly prevented the membrane depolarization, particularly, under hyperosmotic conditions. *Significant difference vs. 290 mosmol L–1.

Discussion

Currently, the accepted explanation for the generation of myotonia is that K+ exits the fibre during voluntary action potentials and subsequently accumulates in the T‐tubules. The resulting K+ increase depolarizes the resting membrane potential (Adrian & Bryant, 1974; Adrian & Marshall, 1976; Wallinga et al. 1999; Fraser et al. 2011) and is suggested to open voltage‐gated Na+ channels, which finally trigger the increased excitability (Burge & Hanna, 2012; Skov et al. 2013). In healthy muscle, the K+ accumulation is balanced by ClC‐1 mediated Cl− currents, which accounts for 70–80% of the resting muscle membrane conductance. Myotonic muscle is a result of decreased Cl− conductance that is incapable of balancing the K+ accumulation, which might explain the small but significant depolarization of the resting membrane potential of ADR mice compared to WT (Mehrke et al. 1988). By contrast, elimination of Cl− conductance by 9‐AC or Cl−‐free solution resulted in hyperpolarizing responses of up to 20 mV (Dulhunty, 1978; Betz et al. 1984; Aickin et al. 1989). Moreover, iso‐osmotic removal of external Cl− results in intracellular Cl− depletion and immediate cell shrinkage (Hamann et al. 2010).

Increasing osmolarity results in cell shrinkage accompanied by a sustained depolarization in healthy skeletal muscle fibres (van Mil et al. 1997; Ferenczi et al. 2004; Pickering et al. 2009; Lindinger et al. 2011). Electrophysiological measurements of cell resting potential from muscle in Cl−‐free solutions suggested that cellular volume reduction proportionally increases the concentrations of intracellular ions following simple predictions of the Goldman–Hodgkin–Katz equation (Adrian, 1956). Similarly, all intracellular ions were conserved in Cl− containing solutions, which permit passive Cl− redistribution. Finally, a significant and maintained increase in [Cl−]intracellular/[Cl−]extracellular and a consequent divergence of the equilibrium potentials for Cl− and K+ were reported to accompany cellular shrinkage in Cl− containing solutions (Ferenczi et al. 2004). Further investigations suggested that persistent activity of NKCC maintains [Cl−]intracellular/[Cl−]extracellular above its electrochemical equilibrium, thereby stabilizing steady‐state resting potential following muscle fibre shrinkage (Ferenczi et al. 2004). Finally, the results of the present study suggest that increased intracellular [Cl−] as a result of cellular shrinkage and increased activity of the NKCC1 might counterbalance decreased Cl− conductance, at least in part. However, depolarization of resting membrane potential failed to reach the level of WT mice. Moreover, hypertonicity was suggested to promote the regenerative closure of KIR, a inward indirected K+ channel (Geukes Foppen et al. 2002). This might result in increased K+ levels in the tubule and subsequent depolarization. Although this depolarization might trigger myotonia, a K+‐induced depolarization of the membrane potential was also reported to render the muscle unexcitable and lead to a loss of force if the membrane potential depolarizes above −60 mV (Juel, 1986; Clausen & Everts, 1991; Cairns et al. 1995; Carins & Dulhunty, 1995; Pedersen et al. 2003).

Inhibition of NKCC1 cotransporter induced accelerated reduction in peak tetanic force during electrical stimulation in healthy soleus muscle, suggesting a key role of NKCC1 in ion regulation during muscle fatigue (Gosmanov et al. 2002). An increased 86Rb+ (a reliable 42K+ analogue for the NKCC1) uptake via the NKCC1 cotransporter was measured during exercise and electrical stimulation of muscle (Wong et al. 1999; Gosmanov et al. 2003). Moreover, after training, a relatively higher content (14% vs. 27%) of NKCC1 cotransporter was detected in soleus and plantaris rat muscle (Gosmanov et al. 2002). Collectively, these data strongly suggest that NKCC1 activity increases in exercising muscle and may play a role in preventing a rise of [K+]0 and thereby postpones K+‐induced muscle fatigue.

NKCC1 co‐transport also plays an important role in volume regulation under hyperosmotic conditions (Zhao et al. 2004). Thus, the NKCC1 cotransporter has been suggested to maintain cellular volume during exercise in non‐contracting but hyper‐osmotically stimulated skeletal muscle, thereby preventing cellular shrinkage, contractility and metabolism (Zhao et al. 2004; Hoppe et al. 2013). Finally, the NKCC1 cotransporter was suggested to have a dual function. In contracting muscle, it participates in the reuptake of K+ to reduce the subsequent rise in [K+]0 and postpone muscle fatigue. In non‐contracting muscle, it regulates muscle volume and prevents shrinkage of muscle cells (Gosmanov et al. 2002). However, whether these mechanisms might influence contractility and the warm‐up phenomenon of myotonic muscle is a focus of the present study.

Previous data have demonstrated that the warm‐up phenomenon occurs when an increase in [K+]° can not be accommodated in the T‐tubular lumen (i.e. AP‐propagation block) (Birnberger & Klepzig, 1979). However, the activation of contractile filaments results in an accumulation/efflux of acidic metabolites resulting in an expansion of the T‐tubular volume (Eisenberg & Gilai, 1979; Usher‐Smith et al. 2007). Osmolarity will first increase in the T‐tubular compartment, then the metabolites and K+ will slowly diffuse outward to the interstitial space and blood (Hess et al. 2005; Shorten & Soboleva, 2007). The findings of the present study clearly demonstrate that increased osmolarity results in a reduction of myotonic stiffness, sustained membrane depolarization and a marked suppression in the proportion of fibres with myotonia. Indeed, osmotic stress primary leads to activation of NKCC1. In the absence of gCl−, the inward transport of Cl− via NKCC1 is essential for osmotic homeostasis. Although the NKCC1 transporter carries no net current, it has a depolarizing influence on the resting potential by enhancing Cl− influx (Geukes Foppen et al. 2002; Russell, 2002; Wu et al. 2013). In non‐myotonic muscle, NKCC1 accounts for more than 30% of K+ import during muscle stimulation (Wong et al. 2001; Zhao et al. 2004). Bumetanide prevented the membrane depolarization and reversed the anti‐myotonic effect of high osmolarity. This effect was more pronounced in muscle from ADR mice compared to pharmacologically‐induced myotonic muscle. Hence, NKCC1 overexpression in low gCl– myotonia is very probable and NKCC1 activation is one key mechanism of the warm‐up phenomenon. Earlier in vitro data recorded from human intercostal muscle suggested that loop diuretics might worsen outcome (Kwiecinski et al. 1988). However, the concentration used in the study by Kwiecinski et al. (1988) was 1 mM, which is 10 to 30 times higher than therapeutic doses. The findings of the present study suggest that loop diuretics worsen the myotonic syndrome and should be avoided or at least dosed sparsely in patients with low gCl– myotonia. Moreover, the data of the present study support the clinical observation that a nutritional increase of serum osmolarity (e.g. water deprivation, carbohydrate rich meals) has anti‐myotonic effects.

Additional information

Competing interests

The authors declare that they have no competing interests.

Author contributions

WK, KH, FLH and SC were responsible for intellectual content and the study design. WK, SC, KH, FLH and KJR were responsible for collection, analysis and interpretation of data. KH, SC, WK and FLH were responsible for drafting the manuscript and graphical representation of data. KJR, SW and WK were responsible for the critical evaluation of the manuscript. All authors approved the final version of the manuscript submitted for publication. All persons designated as authors quality for authorship and all those who qualify for authorship are listed.

Funding

FL‐H is endowed Senior Research Professor of Neuroscience of the non‐profit Hertie Foundation. SC was supported by a governmental scholarship (‘Land Baden‐Württemberg’) for the promotion of young scientists and the German Academic Exchange Service (DAAD). We also thank the Deutsche Gesellschaft für Muskelkranke (DGM) for the grant to FL‐H and KJR for research on myotonia.

Acknowledgements

This work is dedicated to Professor Dr Dr hc Frank Lehmann‐Horn, who passed away in May 2018. His enthusiasm, helpfulness and his bright mind are not forgotten. Parts of this work were performed by Sunisa Chaiklieng and presented at the 36th European Muscle Conference in Heidelberg, Germany.

Biography

Kerstin Hoppe received her MD in medicine at University of Wuerzburg, Germany. She completed her specialization as a clinical anaesthesiologist at the University of Ulm, Germany. During her time there, she was trained as a post‐doc in the neurophysiological muscle laboratory of Werner Klingler. Her research focuses on muscle diseases and immunology.

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Edited by: Michael Hogan & Michael Shipston

References

  1. Adrian RH (1956). The effect of internal and external potassium concentration on the membrane potential of frog muscle. J Physiol 133, 631–658. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Adrian RH & Freygang WH (1962). The potassium and chloride conductance of frog muscle membrane. J Physiol 163, 61–103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Adrian RH & Peachey LD (1973). Reconstruction of the action potential of frog satorius muscle. J Physiol 235, 103–131. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Adrian RH & Bryant SH (1974). On the repetitive discharge in myotonic muscle fibres. J Physiol 240, 505–515. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Adrian RH & Marshall MW (1976). Action potentials reconstructed in normal and myotonic muscle fibres. J Phyiol 258, 124–143. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Aickin CC, Betz WJ & Harris GL (1989). Intracellular chloride and the mechanism for its accumulation in rat lumbrical muscle. J Physiol 411, 437–455. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Almers W (1972). Potassium conductance changes in skeletal muscle and the potassium concentration in the transverse tubules. J Physiol 225, 33–56. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Betz WJ, Caldwell JH & Kinnamon SC (1984). Physiological basis of a steady endogenous current in a rat lumbrical muscle. J Gen Physiol 83, 175–192. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Birnberger KL & Klepzig M (1979). Influence of extracellular potassium and intracellular pH on myotonia. J Neurol 222, 23–35. [DOI] [PubMed] [Google Scholar]
  10. Blinks JR (1965). Influcence of osmotic strength on cross‐section and volume of isolated single muscle fibres. J Physiol 177, 42–57. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Boivin GP, Bottomley MA, Dudley ES, Schiml PA, Wyatt CN & Grobe N (2016). Physiological, behavioral, and histological responses of male C57BL//6N mice to different CO2 chamber replacement rates. J AM Assoc Lab Anim Sci 55, 451–461. [PMC free article] [PubMed] [Google Scholar]
  12. Burge JA & Hanna MG (2012). Novel insights into the pathomechanis of skeletal muscle channelopathies. Curr Neurol Neurosci Rep 12, 62–69. [DOI] [PubMed] [Google Scholar]
  13. Byrant SH & Morales‐Aguilera A (1971). Chloride conductance in normal and myotonic muscle fibres and the action of monocarboxylic acids. J Physiol 219, 393–383. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Cairns SP, Flatman JA & Clausen T (1995). Relation between extracellularl [K+], membrane potential and concentration in rat soleus muscle: modulation by the Na+‐K+ pump. Pflugers Arch 430, 909–915. [DOI] [PubMed] [Google Scholar]
  15. Carins SP & Dulhunty AF (1995). High‐frequency fatigue in rat skeletal muscle: role of extracellular ion concentrations. Muscle Nerve 18, 890–898. [DOI] [PubMed] [Google Scholar]
  16. Chinet A (1993). Ca(2+)‐dependent heat production by rat skeletal muscle in hypertonic media depends on Na(+)‐Cl– cotransport stimulation. J Physiol 461, 689–703. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Clausen T & Everts ME (1991). K(+)‐induced inhibition of contractile force in rat skeletal muscle: role of active Na(+)‐K+ transport. Am J Physiol Cell Physiol 261, C799–807. [DOI] [PubMed] [Google Scholar]
  18. Colding‐Jorgensen E (2005). Phenotypic variability in myotonia congenital. Muscle Nerve 32, 19–34. [DOI] [PubMed] [Google Scholar]
  19. Delpire E, Rauchmann MI & Beier DR (1994). Molecular cloning of and chromosome localization of a putative basolateral Na+‐K+‐2Cl− cotransporter from mouse inner medullary collecting duct (mIMCD‐3) cells. J Biol Chem 269, 25677–25683. [PubMed] [Google Scholar]
  20. Desaphy JF, Costanza T, Carbonara R & Conte Camerino D (2013). In vivo evaluation of antimyotonic efficacy of β‐adrenergic drugs in a rat model of myotonia. Neuropharmacology 65, 21–27. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Dulhunty AF (1978). The dependence of membrane potential on extracellular chloride concentration in mammalian skeletal muscle fibres. J Physiol 276, 67–82. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Eisenberg BR & Gilai A (1979). Structural changes in single muscle fibres after stimulation at a low frequency. J Gen Physiol 74, 1–16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Ferenczi EA, Fraser JA, Chawla S, Skepper JN, Schweining CJ & Huang C‐LH (2004). Membrane potential stabilization in amphibian skeletal muscle fibres in hypertonic solutions. J Physiol 555, 423–438. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Fraser JA, Huang CL & Pedersen TH (2011). Relationships between resting conductances, excitability, and t‐system ionic homeostasis in skeletal muscle. J Gen Physiol 138, 95–116. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Furmann RE & Barchi RL (1978). The pathophysiology of myotonia produced by aromatic carboxylic acids. Ann Neurol 4, 357–365. [DOI] [PubMed] [Google Scholar]
  26. Gallaher J, Bier M & Siegenbeek van Heukelom J (2009). The role of chloride transport in the control of the membrane potential in skeletal muscle – theory and experiment. Biophys Chem 143, 18–25. [DOI] [PubMed] [Google Scholar]
  27. Geukes Foppen RJ, van Mil HG & Siegenbeek van Heukelom J (2001). Osomolarity influences bistability of membrane potential under hypokalemic conditions in mouse skeletal muscle: an experimental and theoretical study. Comp Biochem Physiol A Mol Integr Physiol 130, 533–538. [DOI] [PubMed] [Google Scholar]
  28. Geukes Foppen RJ, van Mil HG & van Heukelom JS (2002). Effects of chloride transport on bistable behaviour of the membrane potential in mouse skeletal muscle. J Physiol 542, 181–191. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Geukes Foppen RJ (2004). In skeletal muscle the relaxation of the resting membrane potential induced by K(+) permeability changes depends on Cl(−) transport. Pflugers Arch 447, 416–425. [DOI] [PubMed] [Google Scholar]
  30. Gosmanov AR, Lindinger MI & Thomason DB (2002). Riding the tides: K+ concentration and volume regulation by muscle Na+‐K+‐2Cl− cotransport activity. News Physiol 542, 181–191. [DOI] [PubMed] [Google Scholar]
  31. Gosmanov AR & Thomason DB (2002). Insulin and isoprtenerol differentially regulate mitogen‐activated protein kinase‐dependent Na+‐K+2Cl− cotransporter activity in skeletal muscle. Diabetes 51, 615–623. [DOI] [PubMed] [Google Scholar]
  32. Gosmanov AR, Schneider EG & Thomason DB (2003). NKCC activity restores muscle water during hyperosmotic challenge independent of insulin, ERK, and p38 MAPK. Am J Physiol Regul Integr Comp Physiol 284, R655–R665. [DOI] [PubMed] [Google Scholar]
  33. Gordon AM & Godt RE (1970). Some effects of hypertonic solutions on contraction and excitation‐contraction coupling in frog skeletal muscles. J Gen Physiol 55, 254–275. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Grundy D (2015). Principles and standards for reporting animal experiments in The Journal of Physiology and Experimental Physiology. J Physiol 593, 2547–2549. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Hamann S, Herrera‐Perez JJ, Zeuthen T & Alvarez‐Leefmans FJ (2010). Cotransport of water by the Na+‐K+2Cl– cotransporter NKCC1 in mammalian epithelial cells. J Physiol 588, 4089–4101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Heine R, Pika U & Lehmann‐Horn F (1993). A novel SCN4A mutation causing myotonia aggravated by cold and potassium. Hum Mol Genet 2, 1349–1353. [DOI] [PubMed] [Google Scholar]
  37. Hess TM, Kronfeld DS, Williams CA, Waldron JN, Graham‐Thiers PM, Greiwe‐Crandell K, Lopes MA & Harris PA (2005). Effects of oral potassium supplementation on acid‐base status and plasma ion concentration of horses during endurance exercise. Am J Vet Res 66, 466–473. [DOI] [PubMed] [Google Scholar]
  38. Hoppe K, Lehmann‐Horn F, Chaiklieng S, Jurkatt‐Rott K, Adolph O & Klinler W (2013). In vitro muscle contracture investigations on the malignant hyperthermia like episodes in myotonia congenita. Acta Anaesthesiol Scand 57, 1017–1023. [DOI] [PubMed] [Google Scholar]
  39. Juel C (1986). Potassium and sodium shifts during in vitro isometric muscle concentration, and the time course of the ion‐gradient recovery. Pflugers Arch 406, 458–463. [DOI] [PubMed] [Google Scholar]
  40. Kristensen M, Hansen T & Juel C (2006). Membrane proteins involved in the potassium shifts during muscle activity and fatigue. Am J Physiol Regul Integr Comp Physiol 290, R766–R772. [DOI] [PubMed] [Google Scholar]
  41. Kristensen M & Juel C (2010). Potassium‐transporting proteins in skeletal muscle: cellular location and fibre‐type differences. Acta Physiol 198, 105–123. [DOI] [PubMed] [Google Scholar]
  42. Kwiecinski H, Lehmann‐Horn F & Rüdel R (1988). Drug induced myotonia in human intercostals muscle. Muscle Nerve 11, 576–581. [DOI] [PubMed] [Google Scholar]
  43. Lehmann‐Horn F, Rüdel R & Jurkat‐Rott K (2004). Nondystrophic myotonias and periodic paralyses In: Myology, eds. Engel AG. & Franzini‐Armstrong C, 3rd edn., pp. 1257–1300. McGraw‐Hill, New York, NY. [Google Scholar]
  44. Lindinger MI, Hawke TJ, Lipskie SL, Schaefer HD & Vickery L (2002). K+ transport and volume regulation response by NKCC in resting rat hindlimb skeletal muscle. Cell Physiol Biochem 12, 279–292. [DOI] [PubMed] [Google Scholar]
  45. Lindinger MI, Leung M, Trajcevski KE & Hawke TJ (2011). Volume regulation in mammalian skeletal muscle: the role of sodium‐potassium‐chloride cotransporters during exposure to hypertonic solutions. J Physiol 589, 2887–2899. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Lossin C & George AL Jr. (2008). Myotonia congenital. Adv Genet 63, 25–55. [DOI] [PubMed] [Google Scholar]
  47. Mehrke G, Brinkmeier H & Jockusch H (1988). The myotonic mouse mutant ADR: electrophysiology of the muscle fibre. Muscle Nerve 11, 440–446. [DOI] [PubMed] [Google Scholar]
  48. Moody CM, Chua B & Weary DM (2014). The effect of carbon dioxide flow rate on the euthanasia of laboratory mice. Lab Anim 48, 298–304. [DOI] [PubMed] [Google Scholar]
  49. Novak KR, Norman J, Mitchell JR, Pinter MJ & Rich MM (2015). Sodium channel slow inactivation as a therapeutic target for myotonia congenita. Ann Neurol 77, 320–332. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Pedersen TH, Clausen T & Nielsen OB (2003). Loss of force induced by high extracellular [K+] in rat muscle: effect of temperature, lactic acid and beta2‐agonist. J Physiol 551, 277–286. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Pickering JD, White E, Duke AM & Steele DS (2009). DHPR activation underlies SR Ca2+ release induced by osmotic stress in isolated rat skeletal muscle fibres. J Gen Physiol 133, 511–524. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Russell JM (2002). Sodium‐potassium‐chloride cotransport. Physiol Rev 542, 181–191. [DOI] [PubMed] [Google Scholar]
  53. Shorten PR & Soboleva TK (2007). Anomalous ion diffusion within skeletal muscle transverse tubule networks. Theor Biol Med Model 4, 18–26. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Skov M, Riisager A, Fraser JA, Nielsen OB & Pedersen TH (2013). Extracellular magnesium and calcium reduce myotonia in ClC‐1 inhibited rat muscle. Neuromuscul Dis 23, 489–502. [DOI] [PubMed] [Google Scholar]
  55. Steinmeyer K, Klacke R, Ortland C, Gronemeier M, Jockusch S, Gründer H & Jentsch TJ (1991). Inactivation of muscle chloride channel by transposon insertion in myotonic mice. Nature 354, 304–308. [DOI] [PubMed] [Google Scholar]
  56. Tang CY & Chen TY (2011). Physiology and pathophysiology of ClC‐1: mechanisms of a chloride channel disease, moyotonia. J Biomed Biotechnol Article ID 685328. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Trivedi JR, Cannon SC & Griggs RC (2014). Nondystrophic myotonia: challenges and future directions. Exp Neurol 253, 28–30. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Usher‐Smith JA, Fraser JA, Huang CL & Skepper JN (2007). Alterations in triad ultrastructure following repetitive stimulation and intracellular changes associated with exercise in amphibian skeletal muscle. J Muscle Res Cell Motil 28, 19–28. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Van Mil HG, Geukes Foppen RJ & Siegenbeek van Heukelom J (1997). The influence of bumetanide on the membrane potential of mouse skeletal muscle cells in isotonic and hypertonic media. Br J Pharmacol 120, 39–44. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Wallinga W, Meijer SL, Alberink MJ, Vliek M,Wienk ED & Ypey DL (1999). Modelling action potentials and membrane currents of mammalian skeletal muscle fibres in coherence with potassium concentration changes in the T‐tubular system. Eur Biophys J 28, 317–329. [DOI] [PubMed] [Google Scholar]
  61. Wong JA, Fu L, Schneider EG & Thomason DB (1999). Molecular and functional evidence for Na+‐K+2Cl− cotransporter expression in rat skeletal muscle. Am J Physiol Regul Integr Comp Physiol 277, R154–R161. [DOI] [PubMed] [Google Scholar]
  62. Wong JA, Gosmanov JR, Schneider EG & Thomason DB (2001). Insulin dependent, MAPK dependent stimulation of NKCC activitiy in skeletal muscle. Am J Physiol Regul Integr Comp Physiol 281, R561–R571. [DOI] [PubMed] [Google Scholar]
  63. Wu F, Mi W & Cannon SC (2013). Bumetanide prevents transient decreases in muscle force in murine hypokalemic periodic paralysis. Neurology 80, 1110–1116. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Zhao H, Hyde R & Hundal HS (2004). Signalling mechanisms underlying the rapid and additive stimulation of NKCC activity by insulin and hypertonicity in rat L6 skeletal muscle cells. J Physiol 560, 123–136. [DOI] [PMC free article] [PubMed] [Google Scholar]

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