Muscle weakness in Ryr1I4895T/WT knock-in mice as a result of reduced ryanodine receptor Ca2+ ion permeation and release from the sarcoplasmic reticulum

Abstract

The type 1 isoform of the ryanodine receptor (RYR1) is the Ca(2+) release channel of the sarcoplasmic reticulum (SR) that is activated during skeletal muscle excitation-contraction (EC) coupling. Mutations in the RYR1 gene cause several rare inherited skeletal muscle disorders, including malignant hyperthermia and central core disease (CCD). The human RYR1(I4898T) mutation is one of the most common CCD mutations. To elucidate the mechanism by which RYR1 function is altered by this mutation, we characterized in vivo muscle strength, EC coupling, SR Ca(2+) content, and RYR1 Ca(2+) release channel function using adult heterozygous Ryr1(I4895T/+) knock-in mice (IT/+). Compared with age-matched wild-type (WT) mice, IT/+ mice exhibited significantly reduced upper body and grip strength. In spite of normal total SR Ca(2+) content, both electrically evoked and 4-chloro-m-cresol-induced Ca(2+) release were significantly reduced and slowed in single intact flexor digitorum brevis fibers isolated from 4-6-mo-old IT/+ mice. The sensitivity of the SR Ca(2+) release mechanism to activation was not enhanced in fibers of IT/+ mice. Single-channel measurements of purified recombinant channels incorporated in planar lipid bilayers revealed that Ca(2+) permeation was abolished for homotetrameric IT channels and significantly reduced for heterotetrameric WT:IT channels. Collectively, these findings indicate that in vivo muscle weakness observed in IT/+ knock-in mice arises from a reduction in the magnitude and rate of RYR1 Ca(2+) release during EC coupling that results from the mutation producing a dominant-negative suppression of RYR1 channel Ca(2+) ion permeation.

Figure 1.

Reduced in vivo muscle strength in IT/+ mice. (A) In vivo hanging task determination of upper body strength in WT (n = 13 mice; black bar) and IT/+ (n = 12 mice; gray bar) mice. (Left) Average hanging scores from 10 trials/mouse (refer to Materials and methods for details). (Right) Percentage of hanging task trials in which WT and IT/+ mice successfully escaped to one of the stanchion supports. (B) Average grip strength (five trials/mouse) assayed from WT (n = 8 mice; black bar) and IT/+ (n = 14 mice; gray bar) mice using a digital force gauge (GTX; Dillon) set to record the peak tensile force generated by mice gripping a metal grid while being pulled away until grip fails. *, P < 0.01; †, P < 0.05.

Figure 2.

Reduction of electrically evoked and 4-CMC–induced RYR1-mediated Ca²⁺ release in indo-1–loaded FDB fibers from IT/+ mice. (A) Representative indo-1 ratio traces for FDB fibers from WT (top) and IT/+ (bottom) mice during successive electrical stimulation (arrowheads) and the application of 500 µM 4-CMC (black bar). (B) Average (±SEM) peak electrically evoked Ca²⁺ transients in FDB fibers from WT (n = 33; black bar) and IT/+ (n = 32; gray bar) mice. (C) Average (±SEM) peak 4-CMC–induced Ca²⁺ responses in FDB fibers from WT (n = 61; black bar) and IT/+ (n = 98; gray bar) mice. (D) Expanded time course of the rising phase of the indo-1 ratio during the application of 500 µM 4-CMC in representative FDB fibers from WT and IT/+ mice. For clarity, each trace is truncated after reaching the peak response. (E) Average (±SEM) peak of the first derivative of the rising phase of 4-CMC–induced Ca²⁺ responses in FDB fibers from WT (n = 61; black bar) and IT/+ (n = 98; gray bar) mice. *, P < 0.01.

Figure 3.

Slowed rate of electrically evoked Ca²⁺ transients in mag-fluo-4–loaded FDB fibers from IT/+ mice. (A) Representative single electrically evoked Ca²⁺ transients elicited during a brief 0.1-Hz train of stimulation in FDB fibers from WT (left) and IT/+ (right) mice. Sampling rate was 10 kHz. (B) The first derivative of the mag-fluo-4 traces within the region marked by two arrows for the traces shown in A. (C) Box plot representation for the dataset of peak electrically evoked mag-fluo-4 transients (ΔF/F_max) in FDB fibers from WT (left) and IT/+ (right) mice. (D) Box plot representation for the dataset for the peak of the first derivative of electrically evoked mag-fluo-4 transients (d(ΔF/F)/dt) in FDB fibers from WT (left) and IT/+ (right) mice. (E) Average (±SEM) peak electrically evoked Ca²⁺ transients (ΔF/F_max) across FDB fibers from WT (n = 3; black bar) and IT/+ (n = 3; gray bar) mice (12–29 fibers/mouse). (F) Mean (±SEM) peak of the first derivative (d(ΔF/F)/dt) across FDB fibers from WT (n = 3; black bar) and IT/+ (n = 3; gray bar) mice (12–29 fibers/mouse).

Figure 4.

Intracellular Ca²⁺ store content is similar in FDB fibers from WT and IT/+ mice. (A) Representative Fura-FF traces in FDB fibers from WT and IT/+ mice during successive applications of 500 µM 4-CMC and ICE (10 µM ionomycin, 30 µM CPA, and 100 µM EGTA/0 Ca²⁺ Ringer’s solution). (B) Average (±SEM) peak Fura-FF responses to 4-CMC in FDB fibers from WT (n = 10; left) and IT/+ (n = 13; right) mice. (C) Average (±SEM) peak rate of change in Fura-FF ratio (F₃₄₀/F₃₈₀) during 4-CMC application in FDB fibers from WT (n = 10; left) and IT/+ (n = 13; right) mice. (D) Average (±SEM) peak ICE responses in FDB fibers from WT (n = 10; left) and IT/+ (n = 13; right) mice. †, P < 0.05.

Figure 5.

L-type Ca²⁺ currents and SR Ca²⁺ release flux in muscle fibers from WT and IT/+ mice. (A) Voltage dependence of L-type Ca²⁺ current density. (B) Normalized Ca²⁺ conductance derived from the data shown in A (V_1/2, k, and G_max were −4.38 ± 1.58 mV, 4.66 ± 0.28 mV, and 154 ± 14.6 SF⁻¹ for WT and −4.57 ± 1.41 mV, 5.33 ± 0.20 mV, and 162 ± 15.3 SF⁻¹ for IT/+, respectively). Mean membrane capacitance was 5.06 ± 0.25 nF (WT) and 4.72 ± 0.38 nF (IT/+). (C) Voltage-activated changes in fura-2 fluorescence ratio. All mean values above −20 mV were significantly (P < 0.05) different, in contrast to the parameters V_1/2 and k, which were not significantly different (−18.5 ± 1.29 mV and 5.15 ± 0.24 mV for WT and −21.6 ± 1.24 mV and 5.17 ± 0.30 mV for IT/+, respectively). (D) Normalized peak Ca²⁺ release flux. Values from the different experiments were averaged after normalization to the maximum. V_1/2 and k were not significantly different (−10.0 ± 1.9 mV and 7.54 ± 0.56 mV for WT and −12.2 ± 1.19 mV and 8.54 ± 0.46 mV for IT/+, respectively). The absolute maximal values (assuming internal EGTA concentration to be 40% of the pipette concentration) at +50 mV were 64.6 ± 17.5 and 49.9 ± 9.4 mM/s, respectively. Current and fluorescence signals were simultaneously recorded in isolated interosseus fibers (WT, filled circles and continuous lines, n = 15; IT/+, open circles and dashed lines, n = 14).

Figure 6.

Single-channel recordings of coexpressed WT and IT channels. (A) Representative single-channel currents were recorded at −35 mV (left traces) or 0 mV (right traces), with openings represented by downward deflections from the closed state (c–) in symmetrical 250 mM KCl and 2 µM cis Ca²⁺ (left traces) and after the subsequent addition of 10 mM trans Ca²⁺ (right traces). Three traces of Group 3 channels recorded in the absence of 10 mM trans Ca²⁺ (left) indicate the absence of well-defined K⁺ conductance. Group 3 channels lack a Ca²⁺ current at 0 mV in the presence of 10 mM trans Ca²⁺. (B) Current–voltage relationships of Group 1 (•) and Group 2 (○) channels in 250 mM of symmetric KCl plus 10 mM trans Ca²⁺. (C) The boxed region indicated in B is expanded to highlight the negative shift in the reversal potential of Group 2 channels. (D) Proposed tetramer assembly model based on the assumption of independent WT and IT subunit assortment (left) and the expected reduction in fractional RYR1 Ca²⁺ conductance in muscle cells obtained from IT/+ and IT/IT mice (right), provided that Group 1 channels arise from tetramers with only one or fewer IT subunits, Group 2 channels from tetramers with two WT and two IT subunits, and Group 3 channels from tetramers with three or more IT subunits.