. 2007 Nov 14;104(47):18537–18542. doi:10.1073/pnas.0709312104

AnRyr1^I4895T mutation abolishes Ca²⁺ release channel function and delays development in homozygous offspring of a mutant mouse line

Elena Zvaritch^*,Frederic Depreux^†,Natasha Kraeva^*,Ryan E Loy^‡,Sanjeewa A Goonasekera^‡,Simona Boncompagni^§,^¶,Alexander Kraev^*,Anthony O Gramolini^*,^‖,Robert T Dirksen^‡,Clara Franzini-Armstrong^§,Christine E Seidman^†,J G Seidman^†,David H MacLennan^*,^**

*Banting and Best Department of Medical Research, University of Toronto, Charles H. Best Institute, 112 College Street, Toronto, ON, Canada M5G 1L6;

^†Department of Genetics, Harvard Medical School and Howard Hughes Medical Institute, 77 Avenue Louis Pasteur, Boston, MA 02115;

^‡Department of Pharmacology and Physiology, University of Rochester School of Medicine and Dentistry, Rochester, NY 14642;

^§Department of Cell and Developmental Biology, University of Pennsylvania School of Medicine, Philadelphia, PA19104-6058;

^¶Center for Research on Aging, Università G. d'Annunzio, Chieti CH I-66013, Italy; and

^‖Department of Physiology, University of Toronto, Toronto, ON, Canada M55 1A8

^✉

**To whom correspondence should be addressed. E-mail:david.maclennan@utoronto.ca

Contributed by David H. MacLennan, October 1, 2007

Author contributions: E.Z., F.D., A.K., A.O.G., R.T.D., C.F.-A., C.E.S., J.G.S., and D.H.M. designed research; E.Z., F.D., N.K., R.E.L., S.A.G., and S.B. performed research; E.Z., A.K., A.O.G., R.T.D., C.F.-A., C.E.S., J.G.S., and D.H.M. analyzed data; and E.Z., A.K., A.O.G., R.T.D., C.F.-A., C.E.S., J.G.S., and D.H.M. wrote the paper.

Received 2007 Sep 7; Issue date 2007 Nov 20.

PMC Copyright notice

PMCID: PMC2141812 PMID:18003898

Abstract

A heterozygous Ile4898 to Thr (I4898T) mutation in the human type 1 ryanodine receptor/Ca²⁺ release channel (RyR1) leads to a severe form of central core disease. We created a mouse line in which the correspondingRyr1^I4895T mutation was introduced by using a “knockin” protocol. The heterozygote does not exhibit an overt disease phenotype, but homozygous (IT/IT) mice are paralyzed and die perinatally, apparently because of asphyxia. Histological analysis shows that IT/IT mice have greatly reduced and amorphous skeletal muscle. Myotubes are small, nuclei remain central, myofibrils are disarranged, and no cross striation is obvious. Many areas indicate probable degeneration, with shortened myotubes containing central stacks of pyknotic nuclei. Other manifestations of a delay in completion of late stages of embryogenesis include growth retardation and marked delay in ossification, dermatogenesis, and cardiovascular development. Electron microscopy of IT/IT muscle demonstrates appropriate targeting and positioning of RyR1 at triad junctions and a normal organization of dihydropyridine receptor (DHPR) complexes into RyR1-associated tetrads. Functional studies carried out in cultured IT/IT myotubes show that ligand-induced and DHPR-activated RyR1 Ca²⁺ release is absent, although retrograde enhancement of DHPR Ca²⁺ conductance is retained. IT/IT mice, in which RyR1-mediated Ca²⁺ release is abolished without altering the formation of the junctional DHPR-RyR1 macromolecular complex, provide a valuable model for elucidation of the role of RyR1-mediated Ca²⁺ signaling in mammalian embryogenesis.

Keywords: calcium, central core disease, ryanodine receptor

Ryanodine receptor/Ca²⁺ release channels, represented by three genetically and pharmacologically distinct isoforms in mammals, play a key role in excitation-contraction (EC) coupling in all muscle cell types by releasing Ca²⁺ stored in the sarcoplasmic reticulum (SR) to generate a contraction (1,2). Type 1 ryanodine receptor (RyR1) predominates in skeletal muscle but is expressed at lower levels in other tissues (3–9). In skeletal muscle, RyR1 is located in the junctional terminal cisternae of the SR, where it assembles in ordered orthogonal arrays. It interacts with dihydropyridine receptor (DHPR) complexes, located in the transverse tubular membrane, and organizes them into tetrad arrays (10–13). Structural changes in the DHPR, produced upon plasma membrane depolarization, result in RyR1-mediated Ca²⁺ release from the SR (orthograde coupling) (14,15), whereas Ca²⁺ entry from extracellular spaces through the DHPR α1-subunit (Ca_V1.1) is enhanced by retrograde interaction with RyR1. RyR1 is a homotetramer with a subunit mass of ≈565 kDa. Six transmembrane sequences located in the C-terminal 700-aa residues of each of the four subunits form the Ca²⁺ release channel (16). A highly conserved hydrophobic sequence separating the last two transmembrane sequences in each of the four subunits forms the selectivity filter of the channel (17). RyR1 and DHPR interact with each other and with a large number of accessory proteins to form a supramolecular complex referred to as a Ca²⁺ release unit (CRU) (2,18,19).

The essential role of DHPR-RyR1 interaction in coordinating skeletal muscle function is highlighted by the fact that ablation of either protein in mice is birth-lethal (20–22), and that mutations in theRYR1 gene are causal of several debilitating skeletal myopathies such as central core disease (CCD) [Online Mendelian Inheritance in Man (OMIM) no.117,000], multiminicore disease (OMIM no.602,771), and nemaline rod myopathy (OMIM no.161,800), and mutations in bothRYR1 andCACNA1S encoding Ca_V1.1 are causal of malignant hyperthermia susceptibility (OMIM no.145,600). Disease mutations inRYR1 have been shown to lead to three distinct channel defects. In one class, referred to as leaky mutations, RyR1 channel activation is hypersensitive to electrical and drug stimulation (23–25). Excessive Ca²⁺ leakage and SR store depletion might contribute to abnormalities in intracellular Ca²⁺ homeostasis and compromise skeletal muscle function. The second class of mutations diminishes or abolishes Ca²⁺ release channel activity in response to activation (26,27). Mutations of this class are clustered in the C-terminal part of the protein and are referred to as EC uncoupling mutations (26). A third group of mutations compromise the stability of the protein complex and leads to a partial or almost-complete ablation of RyR1 in skeletal muscle (28,29).

The I4898T RyR1 mutation in the selectivity filter of the channel has been associated with the dominant inheritance of severe central core disease in at least seven unrelated families worldwide (24,30). Assessment of the effect of this mutation on RyR1 channel functionin vitro has led to inconsistent conclusions. Transient expression of the corresponding rabbit RyR1 mutation (I4897T) in HEK-293 cells displayed characteristics of a leaky channel (24). Increased ER/SR Ca²⁺ leak was also reported for B-lymphocytes (31) and myotubes (32) derived from CCD patients carrying the I4898T mutation. However, transient expression of the rabbit I4897T mutant in primary myotubes derived from RyR1-null (dyspedic) mice resulted in a loss of functional EC coupling (26), suggesting that the mutation disrupts Ca²⁺ permeation through the channel.

To investigate the pathogenic effect of the disease mutationin vivo, we generated a knockin mouse line carrying the analogous I4895T mutation in murine RyR1. Here, we show that homozygous I4895T mice are paralyzed and die at birth. The mutant Ca²⁺ release channel is unresponsive to electrical and drug stimulation despite the fact that channel assembly, targeting to SR-sarcolemmal junctions, DHPR tetrad alignment, and retrograde DHPR coupling remain intact. Homozygous mice exhibit abnormal myogenesis and a marked delay in dermatogenesis, skeleton formation, and cardiovascular development, suggesting a direct involvement of RyR1-mediated Ca²⁺ signaling in these processes.

Results

Generation and Viability of I4895T Knockin Mice.

TheRyr1^I4895T mutation was introduced into the murine gene by using homologous recombination in ES cells. The strategy for introduction and confirmation of theRyr1^I4895T mutation is described insupporting information (SI)Text andSI Fig. 5. The mutated allele is referred to as IT. IT/+ mice were viable and fertile and did not develop central cores in their skeletal muscle. Homozygous IT/IT mice survived gestation and were born at a predicted Mendelian frequency (24% IT/IT, 50% IT/+, 26% WT;n = 157). Homozygous IT/IT mice were readily identifiable among the neonates (Fig. 1A andB), because they retained a curved embryonic position, did not respond to stimuli, and were devoid of skeletal muscle contractions. Initially pink in color, they turned cyanotic within minutes after birth and died, most probably of asphyxia.

Fig. 1. — Gross morphology and histological abnormalities in IT/IT neonates. Gross lateral view of WT (A) and IT/IT (B) littermates. Note the smaller size, the curved embryonic posture, and the transparent skin of the IT/IT pup (B). (*C–F*) HE-stained tissue sections. (C andD) Sections through the skin layers at the cervical region. Arrow shows the dermis. BF, brown fat; SM, skeletal muscle. (Magnification: ×10.) (E andF) Sections of hind limb skeletal muscle. (Magnification: ×20.) (G andH) Electron micrographs of myotubes from E18 fetal diaphragm. Arrowheads show myofibril branchings in IT/IT myotubes (H). Triads, indicated with arrows, are more frequent in WT muscles (G).

Gross Morphology of IT/IT Neonates.

Severely reduced skeletal muscle bulk was a striking feature of IT/IT neonates. Tissues at all sites where skeletal muscles would normally be found were jelly-like and disintegrated easily during dissection. The diaphragm was essentially transparent, and all bones and joints were easily dislocated. The skin was smooth, transparent, and tightly followed body contours (Fig. 1B), unlike the wrinkled, well developed skin of WT siblings (Fig. 1A). Inspection of hematoxylin-and-eosin (HE)-stained tissue sections revealed a severely underdeveloped dermis with a very thin layer of relatively undifferentiated cells and poorly defined hair follicles (Fig. 1D), compared with WT (Fig. 1C). IT/IT neonates exhibited excessive deposits of brown adipose tissue in the cervical region (Fig. 1D). Brown fat hyperplasia has been reported in other murine models of skeletal muscle paralysis or ablation (20,33,34).

Skeletal Muscle Histology.

In WT neonates, skeletal muscle groups were well formed and were composed of elongated, aligned, and striated myofibers with nuclei located predominantly in the periphery (Fig. 1E andSI Fig. 6). By contrast, muscle patterning could be outlined only tentatively in IT/IT neonates, and muscles were composed of scarce, poorly aligned myotubes with no obvious cross striations and were embedded within an amorphous mass of connective tissue (Fig. 1F andSI Fig. 6). Some areas showed probable degeneration, with shortened myotubes containing central stacks of pyknotic nuclei.

Myotube formation was detected in the cervical and facial skeletal muscles as early as embryonic day (E) 14.5 in both IT/IT and WT embryos (SI Fig. 6A andB, respectively) and tended to progress to other skeletal muscles as a rostro-caudal wave. Although fusion of myoblasts and multinucleation was apparent at normal early times of embryogenesis, IT/IT myotubes failed to form myofibers. They appeared to become arrested at a stage of myogenesis in which nuclei were centrally located before their migration to the periphery of the myotube (SI Fig. 6).

Ultrastructural Organization of IT/IT Myotubes.

Electron microscopy (EM) analysis of skeletal muscle samples from E16–18 IT/IT fetuses revealed general myofibrillar disorder and paucity of intracellular membrane systems. Poor alignment of cross striations, myofibril splitting, and abrupt changes in the orientation of either a whole or a portion of a myofibril were the most obvious defects (Fig. 1H) when compared with WT (Fig. 1G). Similar abnormalities of the skeletal muscle development and organization were detected previously in the dysgenic (21,35) and dyspedic mouse models (22,36), suggesting that similar intracellular pathways might be affected in all three models.

Skeletal Abnormalities.

Bone and cartilage staining revealed a delay in ossification of the skeleton in IT/IT neonates (Fig. 2A–D). IT/IT neonates exhibited a dome-shaped skull and a persistent cervical kyphosis (Fig. 2B), characteristic of earlier embryonic stages. The delay was especially pronounced in the cranial part of the skull (Fig. 2D), heel bones (talus and calcaneus), and phalanges of the digits (Fig. 2B). A cleft secondary palate was detected in 80% of IT/IT neonates (11 of 14 mice). The rib cage of IT/IT mice was slightly excavated and laterally enlarged, but otherwise properly formed. It is noteworthy that the skeletons of IT/IT mice, although underdeveloped, did not exhibit malformations found in other models of skeletal muscle paralysis, such as micro- and retrognathia, absence of the deltoid tuberosity of the humerus, fused cervical vertebrae, or malformed rib cage with ribs extending perpendicular to the vertebral axis (21,22,33).

Growth Retardation and Receding Edema.

IT/IT neonates were smaller in stature and lighter than their WT littermates (Fig. 1A andB). Average newborn IT/IT body weight (1.12 ± 0.07 g,n = 35) was ≈20% lower than that of WT controls (1.42 ± 0.13 g,n = 40). The earliest sign of abnormal development in IT/IT embryos was a mild, local, subcutaneous edema (Fig. 2F) that was detected in the cervical region at E13.5–14 and receded by E16.5. A larger-scale receding edema was reported previously in dysgenic mice (20). Growth delay was observed in IT/IT embryos starting from E15 (compareFig. 2E andF). It is of interest that growth retardation has not been reported in other murine models of skeletal muscle paralysis or ablation.

Delay in Cardiogenesis.

The inspection of whole-body parasagittal sections of IT/IT neonates revealed a significant delay in cardiac development (Fig. 2G). The heart retained the early embryonic antero-posterior orientation in contrast to the oblique orientation that is characteristic of later-stage fetuses from approximately E14.5–15 and onward (37). Abnormal cardiac positioning precluded proper formation of the outflow tract, interfering with bending of the arterial arch and pulmonary trunk. The neonatal IT/IT heart retained a clearly visible interventricular groove; secondary blood vessels branching from the coronary artery were poorly developed or absent (data not shown). The size, shape, and wall thickness of the right and left ventricles remained approximately equal (Fig. 2H). An atrial septal closure defect with underdeveloped septum primum and poorly developed or absent septum secundum was a consistent feature in IT/IT neonatal hearts (Fig. 2H andI).

Triad Assembly.

The I4895T mutation could potentially affect folding and/or subcellular targeting of RyR1 to SR-sarcolemmal junctional (jSR) domains so that skeletal muscle paralysis might result from a disruption of the physical interaction between RyR1 and DHPR. Skeletal muscles from IT/IT fetuses were examined by EM to answer three questions: whether jSR domains form appropriate structural connections with transverse (T) tubules; whether RyR1 tetramers are targeted and form ordered arrays within the jSR membrane; and whether RyR1 establishes a normal structural relationship with the DHPR in plasmalemma/transverse (T) tubules.

The first two questions were answered by examining thin sections of diaphragm and limb muscles from E16–18 fetuses that, in WT, exhibited sufficiently differentiated myotubes and/or muscle fibers. In all muscles examined, some association between the SR and sarcolemma (peripheral couplings; data not shown) or between the SR and T tubules (dyads and triads;Fig. 3A–F) was found, although the frequency of associations was lower and more variable in IT/IT muscles (Fig. 3D–F). For example, in cross sections of E18 diaphragm, we counted 11.7 ± 7.1 triads/dyads (or junctions) per 100 μm² of cross sectional area in WT, versus only 2.6 ± 2.8 in the mutant (mean ± SD from 40 micrographs, two mice). In IT/IT muscles, there was evidence for retardation in the process by which triads reach their final adult position. At E16–18, junctions were not yet perfectly aligned along transverse planes in either WT (Fig. 3A–C) or mutant (Fig. 3D–F) muscles, but misalignment was more frequent in the mutants. This feature was observed previously in dysgenic (38) and dyspedic muscles (36).

Fig. 3. — Normal triad structure and DHPR tetrad organization in the muscles of E18 IT/IT fetuses. Electron micrographs of WT (*A–C*) and IT/IT (*D–F*) diaphragm muscle. Triads have the same general features in both cases. Their orientation varies between longitudinal and transverse. Electron-dense feet are periodically distributed in the junctional gap. Long arrows indicate the long axis of the muscle cell. (G andH) Arrays of tetrads in the plasmalemma of IT/IT and WT limb muscles. Yellow dots mark the center of a tetrad and emphasize the pattern. The examples of complete and incomplete tetrads in both groups are shown in the smaller boxes beneathG andH.

The basic structure of the junctions was essentially the same in mutant and WT muscle. Within the triads, “feet,” representing the cytoplasmic domains of RyR tetramers, formed long ordered rows in the junctional gap between SR and T tubules (Fig. 3A–F). In the SR lumen, a fine, electron-dense network representing the calsequestrin polymer was present in both WT and IT/IT muscle.

The structural relationship between DHPR and RyR1 was explored by freeze fracture, which reveals the DHPR as clusters of large particles in the cytoplasmic leaflet of the plasmalemma/T tubules. Diaphragm and limb muscles from E16–18 fetuses were examined. Particle clusters were much easier to find in WT than in IT/IT muscle. They consisted of specific groupings containing a mixture of complete four-particle tetrads and incomplete tetrads with three or two appropriately located particles (Fig. 3G,H, and the smaller boxes beneath each). By “dotting” the apparent center of tetrads, an arrangement of small particle groups into ordered arrays with an orthogonal symmetry could be recognized in both WT and mutant muscles. The spacing between the dotted centers of tetrads was not significantly different between WT (48.2 ± 6.9 nm) and IT/IT (47.6 ± 9.7 nm) embryos (mean ± SD). The results of quantitative analysis of particles and their positioning are reported inSI Table 1. WT and IT/IT muscles did not differ significantly in any of the measured parameters, although WT muscles had a slightly higher percentage of four-particle tetrads.

EM and freeze-fracture analyses demonstrated that the mutant protein is properly assembled and targeted to jSR and that it retains the ability to form ordered arrays and organize DHPR tetrads within these junctions.

Functional Activity.

The next question we asked was whether I4895T homotetramers support bidirectional DHPR-RyR1 signaling and Ca²⁺ release during EC coupling. Skeletal muscle paralysis in IT/IT mice could result from severe SR Ca²⁺ leak through mutant Ca²⁺ release channels that leads to near-complete SR Ca²⁺ store depletion. Alternatively, the mutation could produce a Ca²⁺ permeation defect that results in failure of release from a full-complement Ca²⁺ store, a condition referred to as EC uncoupling.

To distinguish between these possibilities, we compared SR Ca²⁺ release channel function in intact indo-1 acetomethoxy ester (AM)-loaded WT and IT/IT myotubes (Fig. 4 and representative traces inSI Fig. 7). Resting indo-1 ratios were similar for WT (0.62 ± 0.01,n = 68) and IT/IT (0.67 ± 0.02,n = 81) myotubes. WT myotubes exhibited robust SR Ca²⁺ release in response to both electrical stimulation and application of 4-CMC (0.5 mM), whereas IT/IT myotubes were completely unresponsive to stimulation (Fig. 4A Left). The lack of SR Ca²⁺ release was apparently not the result of severe SR Ca²⁺ leak, because both the rate of SR Ca²⁺ leak into the myoplasm after SERCA blockade with 30 μM cyclopiazonic acid (CPA) (Fig. 4B) and SR Ca²⁺ store capacity, assessed by using a rapid-release mixture (ICE; 10 μM ionomycin, 30 μM CPA, and 100 μM EGTA/0 Ca²⁺) (Fig. 4A Middle), were similar in WT and IT/IT myotubes. As a positive control for the latter assay, a significant reduction in the ICE-induced Ca²⁺ release was observed after pretreatment of WT myotubes with a submaximal concentration (10 μM) of CPA (Fig. 4A Right).

Fig. 4. — Functional properties of RyR1 in WT and IT/IT myotubes. (A) Histogram representation of maximal Ca²⁺ responses after either electrical stimulation (*Left*), 4-CMC (0.5 mM) application (*Center*), or addition of ICE (*Right*). (B) Summary of average time-to-peak (TTP) increase in cytosolic Ca²⁺ after application of 30 μM CPA. (C andD) Distribution of maximal L-current conductance (G_max) and Ca²⁺ release [(ΔF/F)_max] values obtained from WT, IT/IT, and dyspedic (Dys) myotubes. Each myotube is represented by a filled circle. Two populations ofG_max values were observed in IT/IT myotubes: those withG_max values similar to Dys myotubes (below dotted line) and those withG_max values larger than that of Dys myotubes (above dotted line). A single population of (ΔF/F)_max values was observed for all IT/IT myotubes. (E) Average voltage dependence of L-current density in WT (circles), lowG_max IT/IT (inverted triangles), highG_max IT/IT (upright triangles), and Dys (squares) myotubes. (F) Average voltage dependence of Ca²⁺ release in WT (circles), IT/IT (triangles), and Dys (squares) myotubes.

EC coupling in skeletal muscle involves a unique bidirectional signaling interaction between RyR1 and DHPR proteins, with the DHPR triggering SR Ca²⁺ release (orthograde coupling) and RyR1 enhancing the Ca²⁺ conducting activity of the DHPR (retrograde coupling) (15,39,40). Paralyzed RyR1-null (dyspedic) mice lack both orthograde and retrograde DHPR-RyR1 coupling (Fig. 4C–F). Representative L-currents and voltage-gated Ca²⁺ transients in WT and IT/IT myotubes are shown inSI Fig. 7. IT/IT myotubes lacked sigmoidal voltage-gated Ca²⁺ release (Fig. 4D andF andSI Table 2), consistent with the absence of DHPR-RyR1 orthograde coupling. Evidence that the very small Ca²⁺ transients observed in IT/IT myotubes reflect Ca²⁺ influx through L-channels during depolarization is based on the fact that these transients exhibited a U-shaped voltage dependence (Fig. 4F), closely followed the time course of the integral of the ionic current (SI Fig. 7D), and were blocked by the addition of extracellular Cd²⁺ (0.5 mM) and La³⁺ (0.2 mM) (data not shown). Similar results were observed after homozygous expression in dyspedic myotubes of the analogous mutation in rabbit RyR1 (41,42). However, unlike dyspedic myotubes, the failure of orthograde coupling in IT/IT myotubes was not due to the complete absence of RyR1 expression or impaired interaction between DHPR and RyR1, because retrograde enhancement of DHPR channel activity was observed in ≈70% (31 of 45;SI Table 2) of IT/IT myotubes (data points above the dotted line inFig. 4C and upright triangles inFig. 4E). Interestingly, a smaller proportion of IT/IT myotubes (14 of 45;SI Table 2) exhibited L-current density similar to that of dyspedic myotubes (data points below the dotted line inFig. 4C and inverted triangles inFig. 4E). The absence of retrograde coupling in a subset of IT/IT myotubes may indicate a delay in differentiation of the cultured myotubes, consistent with the delay in IT/IT muscle development presented inFigs. 2 and3 andSI Fig. 6.

Discussion

In earlier studies, the human RyR1 mutation, I4898T, was linked to a severe form of central core disease (24,30). We have generated a knockinRyr1^I4895T mouse line carrying the analogous mutation in murine RyR1. As in many mouse models, the I4895T mutant mouse line does not display all of the characteristics of the human mutant. In this case, heterozygous mice do not display the severe form of central core disease seen in human carriers. However, analysis of the homozygous mutants has provided us with the opportunity to define several aspects of structure–function relationships of the mutant RyR1 proteinin vivo and to uncover key aspects of mammalian development.

The I4895T RyR1 is expressed in the mutant mouse line as a full-length protein. Although total skeletal muscle membrane protein is reduced in the IT/IT mutant, the ratio of RyR1 protein/total membrane protein is approximately the same in mutant and WT muscles (SI Fig. 5D). The mutant RyR1 is targeted to the jSR and forms appropriate orthogonal arrays of feet structures in the gap between the SR and the T tubule. Freeze-fracture studies demonstrate that I4895T RyR1 homotetramers retain the ability to organize DHPR molecules into arrays of tetrads.

The RyR1–DHPR complex, referred to as a CRU (2,8,12), has two important functional characteristics: orthograde activation by the DHPR of Ca²⁺ release channel function through RyR1 and enhancement of retrograde entry of Ca²⁺ through the DHPR by interaction with RyR1. Our studies clearly show that mutant RyR1 retains the capacity for enhancement of retrograde Ca²⁺ entry, but completely lacks voltage- and 4-CMC-induced Ca²⁺ release, despite the fact that SR Ca²⁺ content was similar to that of WT myotubes. The lack of an effect on either SR Ca²⁺ store content or the rate of SR Ca²⁺ leak after SERCA blockade provides strong evidence that I4895T channels do not promote SR Ca²⁺ leak, but rather exhibit a defect in Ca²⁺ permeation/gating. Because disrupted Ca²⁺ permeation through the Ca²⁺ release channel seems not to invoke significant perturbation of RyR1 structure or the higher level of CRU organization, any deleterious effects of the mutation can be related directly to the loss of RyR1-mediated Ca²⁺ signaling.

An important feature of homozygous IT/IT pups is a general delay in embryogenesis. The delay becomes apparent from approximately E15.5–16 onward, even thoughRyr1 transcripts are detected as early as E9.5 in somites (43) and E11.5 in the cerebral cortex (44). In our study, the only early sign of defective IT/IT embryo development was a mild and receding subcutaneous edema occurring at approximately E13.5 and suggesting possible abnormalities in vascular development. The mild effect of the mutation at early embryonic stages before E15.5–16 might indicate that RyR1-mediated Ca²⁺ signaling is dispensable or redundant during these stages.

By contrast, the activity of RyR1 is crucial for later stages of embryogenesis. In IT/IT mice, skeletal muscle development is arrested at a stage in which myoblasts are fused to form myotubes, but nuclei remain stacked and centrally located. EM studies demonstrate a delay in triad formation, misaligned sarcomeres, and branching of the short and sparse myofibrils, consistent with impairment of late stages of myofibrillogenesis. Thickening and wrinkling of the skin, a hallmark of an E15.5–16 mouse embryo (37), does not take place. Ossification of the neurocranium and the skeleton is also delayed after E16. The I4895T mutation has critical effects on the development of the cardiovascular system. The IT/IT late embryonic and neonatal heart fails to attain its normal oblique orientation, instead retaining its early embryonic antero-posterior disposition. This was seen to affect proper development and bending of the outflow tract and would, no doubt, cause circulatory problems in the IT/IT fetuses. Ventricular chambers do not form appropriately. An atrial septal defect with underdeveloped or absent septum secundum, observed in the neonatal IT/IT heart, is another indicator of the developmental delay. A low level of expression of RyR1 in neonatal mouse hearts has been reported (45), but its role in cardiac development has never been explored.

To the best of our knowledge, the effect of the RyR1-mediated Ca²⁺ release on dermatogenesis and ossification has not been reported previously. Developmental retardation in these tissues, as well as in muscle tissues, might be rooted in their common origin in the differentiating layers of somites: the dermatome, myotome, and sclerotome (37). It appears that all three layers of somites are affected by the RyR1 mutation.

Previous studies have demonstrated that E15.5–16 provides a critical window for murine embryogenesis in which innervation of skeletal muscle and other organs takes place, neuromuscular junctions are formed, and fetuses gain synchronous skeletal muscle contractility and voluntary movements (37,46). Our observation that embryonic development in IT/IT mice is delayed after E15.5, a time frame that coincides with innervation, suggests that RyR1-mediated Ca²⁺ signaling might contribute to nerve-induced signal transduction events that direct murine embryonic development. Earlierin vitro studies inXenopus and murine myogenic cells suggested a role of the RyR1-mediated Ca²⁺ release in myogenesis (47–49). These data were further supported by the demonstration of impaired myogenesis in both dyspedic (22) and dysgenic mouse models (20,21). However, interpretation of these data were hampered by the fact that elimination of either Ca_V1.1 or RyR1 could disrupt other protein–protein interactions, which was required for myogenesis, but were not dependent on the Ca²⁺ release function of RyR1. The fact that embryonic defects in theRyr1^I4895T mouse line can be correlated with the specific loss of RyR1 Ca²⁺ permeation makes the line a valuable model for elucidation of the role of RyR1-mediated Ca²⁺ signaling in mammalian embryogenesis.

Methods

Detailed information is provided asSI Materials and Methods. All animal care protocols were approved by and conformed with the guidelines of the institutional animal care and use committees at all universities involved in this study.

Generation of the I4895T Knockin Mouse Line.

TheRyr1^I4895T mutation was introduced into the murine gene by using homologous recombination in ES cells. The PGK-Neo^r cassette was excised from the mouse genome by mating the mutant mice with E2A-Cre transgenic mice. HeterozygousRyr1^I4895T mice were intercrossed to obtain homozygous (IT/IT) mice. The procedures used for genotyping, RNA and microsomal protein preparation, protein analysis, Western blotting, histological, and skeletal stainings are described inSI Materials and Methods.

Electron Microscopy.

Diaphragm and limb muscles from E16–18 embryos were fixed with 3.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.2, and then prepared for thin sectioning and freeze fracture essentially as described in ref.11. The location of tetrads was estimated by the dotting approach (13).

Functional Measurements in Myotubes.

Skeletal myotubes from primary cultures of myoblasts isolated from limb muscles of newborn WT and IT/IT mice were prepared as described in ref.40. Intracellular Ca²⁺ measurements in intact myotubes were obtained from indo-1 AM-loaded myotubes bathed in a normal rodent Ringer's solution (50). Electrically evoked Ca²⁺ transients were elicited by using field stimulation (8 V, 20 ms) applied every 10 s. Agonist-induced Ca²⁺ release was triggered by the local addition of 0.5 mM 4-chloro-m-cresol (4-CMC). The rate of SR Ca²⁺ leak was assessed after blockade of SERCA pumps by application of 30 μM CPA in a Ca²⁺-free Ringer's solution. The magnitude of SR Ca²⁺ store content was assessed after the application of a release mixture designed to dump SR Ca²⁺ stores rapidly (ICE; 10 μM ionomycin, 30 μM CPA, and 100 μM EGTA/0 Ca²⁺). Perforated voltage clamp measurements of L-type Ca²⁺ channel activity and intracellular Ca²⁺ release were conducted as described in ref.50.

Supplementary Material

Supporting Information

pnas_0709312104_index.html^{(19.8KB, html)}

Acknowledgments

We thank Dr. A. Nagy (Samuel Lunenfeld Research Institute, Mount Sinai Hospital, Toronto) for the gift of a floxed PGK-Neo^r cassette and a PGK-TK cassette; Dr. P. Leder (Harvard Medical School) for the gift of the embryonic stem cell line, 129S6/SvEv Tc1; Dr. H. Westphal (National Institutes of Health, Bethesda), for the gift of E2A-Cre transgenic mice; Dr. V. Sorrentino (University of Siena, Siena, Italy) for the gift of RyR1-specific polyclonal antibodies; Dr. David Conner (Harvard Medical School) for carrying out blastocyte injections; Dr. A. Russ Tupling (University of Waterloo, Waterloo, ON, Canada) for discussions of this project; and Ms. Golnar Ziaeian for expert technical assistance. This work was supported by Canadian Institutes of Health Research Grants MT 3399 and MOP 49493 (to D.H.M.); a Muscular Dystrophy Association grant (to A.O.G. and D.H.M.); National Institutes of Health Grants AR044657 (to R.T.D.) and AR052354 (to R.T.D. and C.F.-A.); a National Institutes of Health Dental and Craniofacial Grant (to R.E.L.); National Heart, Lung, and Blood Institute and National Institutes of Health grants (to J.G.S. and C.E.S.); and a Howard Hughes Medical Institute grant (to C.E.S.).

Footnotes

The authors declare no conflict of interest.

This article contains supporting information online atwww.pnas.org/cgi/content/full/0709312104/DC1.

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