Skeletal muscle excitation-contraction coupling: who are the dancing partners?

The International Journal of Biochemistry & Cell Biology 48 (2014) 28–38

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The International Journal of Biochemistry & Cell Biology journal homepage: www.elsevier.com/locate/biocel

Review

Skeletal muscle excitation–contraction coupling: Who are the dancing partners? Robyn T. Rebbeck, Yamuna Karunasekara, Philip G. Board, Nicole A. Beard, Marco G. Casarotto, Angela F. Dulhunty ∗ John Curtin School of Medical Research, Australian National University, PO Box 334, Canberra, ACT 2601, Australia

a r t i c l e

i n f o

Article history: Received 28 September 2013 Received in revised form 29 November 2013 Accepted 4 December 2013 Available online 24 December 2013 Keywords: Excitation–contraction coupling Skeletal muscle Ryanodine receptor Dihydropyridine receptor Sarcoplasmic reticulum

a b s t r a c t There is an overwhelming body of work supporting the idea that excitation–contraction coupling in skeletal muscle depends on a physical interaction between the skeletal muscle isoform of the dihydropyridine receptor L-type Ca2+ channel and the skeletal isoform of the ryanodine receptor Ca2+ release channel. A general assumption is that this physical interaction is between “critical” residues that have been identified in the II–III loop of the dihydropyridine receptor alpha subunit and the ryanodine receptor. However, despite extensive searches, the complementary “critical” residues in the ryanodine receptor have not been identified. This raises the possibility that the coupling proceeds either through other subunits of the dihydropyridine receptor and/or other co-proteins within the large RyR1 protein complex. There have been some remarkable advances in recent years in identifying proteins in the RyR complex that impact on the coupling process, and these are considered in this review. A major candidate for a role in the coupling mechanism is the beta subunit of the dihydropyridine receptor, because specific residues in both the beta subunit and ryanodine receptor have been identified that facilitate an interaction between the two proteins and these also impact on excitation–contraction coupling. This role of beta subunit remains to be fully investigated as well as the degree to which it may complement any other direct or indirect voltage-dependent coupling interactions between the DHPR alpha II–III loop and the ryanodine receptor. © 2014 Published by Elsevier Ltd.

Contents 1.

2.

3.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Membrane geometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Alignment of essential proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Excitation contraction coupling proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. The dihydropyridine receptor and ryanodine receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.1. The dihydropyridine receptor alpha subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. The dihydropyridine receptor beta subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3. Dihydropridine receptor beta subunit binding to the ryanodine receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.4. Update on dihydropridine receptor beta subunit contribution to excitation–contraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.5. The binding site on the ryanodine receptor for the dihydropyridine receptor beta subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.6. Stac3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Additional triadic proteins that influence excitation–contraction coupling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Major junctional proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1. Calsequestrin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Triadin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.3. Junctin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author. Tel.: +61 2 6125 4491; fax: +61 2 6125 4761. E-mail address: [email protected] (A.F. Dulhunty). 1357-2725/$ – see front matter © 2014 Published by Elsevier Ltd. http://dx.doi.org/10.1016/j.biocel.2013.12.001

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Minor junctional proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1. Histidine rich Ca2+ binding protein (HRC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2. Junctophilins (JPs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.3. Mitsugumin-29 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.4. JP-45 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.5. Junctate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Additional regions of the ryanodine receptor that influence skeletal excitation–contraction coupling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3.2.

4. 5.

Abbreviations EC coupling excitation contraction coupling C2C12 a skeletal muscle cell line CSQ calsequestrin CSQ1 skeletal isoform of calsequestrin CSQ2 cardiac isoform of calsequestrin dihydropyridine receptor DHPR DHPR ␣ DHPR alpha subunit DHPR ␣1S skeletal isoform of the DHPR alpha subunit DHPR ␤ DHPR beta subunit DHPR ␤1a skeletal isoform of the DHPR beta subunit ER endoplasmic reticulum FRET fluorescence resonance energy transfer GK guanylate kinase domain HEK cells human embryonic kidney cells HRC histidine rich Ca2+ binding protein JP junctophilin skeletal isoform of junctophilin JP1 JP2 cardiac isoform of junctophilin JP3 skeletal isoform of junctophilin MAGUK membrane-associated guanylate kinases MG29 mitsigummin-29 MORN membrane occupation and recognition nexus domains Orai1 calcium release-activated calcium channel protein 1, located in the plasmalemma RyR ryanodine receptor RyR1 skeletal isoform of the ryanodine receptor RyR2 cardiac isoform of the ryanodine receptor third isoform of the ryanodine receptor first identified in RyR3 brain SH3 an Src homology 3 domain SOCE store operated Ca2+ entry protein–protein interaction domains first identified in SPRY tyrosine kinase spore lysis A (SplA) and mammalian RyRs SR sarcoplasmic reticulum STAC3 an Src homology 3 and cysteine rich domain 3 protein located in the T-tubule of skeletal muscle STIM1 stromal interaction molecule 1, component of store operated Ca2+ entry located in the sarcoplasmic reticulum membrane T cells T lymphocytes TRPC transient receptor potential cation channels Trisk95 the main triadin isoform associated with the skeletal junctional face membrane T-tubule transverse tubule 1. Introduction It is well established that excitation–contraction (EC) coupling in skeletal muscle depends on a physical interaction between the skeletal isoforms of the dihydropyridine receptor L-type Ca2+ channel (DHPR) and the ryanodine receptor Ca2+ release channel (RyR1). However the molecular mechanisms underlying the interactions between these proteins remain elusive despite many

Fig. 1. The membrane systems of a skeletal muscle fiber and the alignment of the two membrane proteins that are essential for skeletal EC coupling, i.e. the DHPR L-type Ca2+ channel in the surface membrane and the RyR1 Ca2+ channel in the SR (sarcoplasmic reticulum) membrane. The surface membrane surrounding the muscle fiber is shown invaginating into a transverse tubule and forming a triadic junction with the membrane surrounding the terminal cisternae of the SR Ca2+ store. The surface membrane component of the junction contains clusters of four DHPR molecules (each containing 5 subunits) which form a “tetrad” structure which is aligned every second RyR. The insert shows the tetrad structure with each of the four DHPR molecules in the tetrad interacting with each of the four subunits of the adjacent RyR1 tetramer.

years of intense investigation. Although some critical residues have been identified, their partner binding residues in the RyR1 calcium release channel, or in potential linker proteins, have not been identified. Thus we have little knowledge of the actual molecular interactions or indeed the sequence of events that facilitate the transmission of the EC coupling signal. The signal originates in the surface membrane voltage sensor in the ␣1S subunit of the DHPR L-type Ca2+ channel. The message is transmitted somehow to the large cytoplasmic domain of RyR1 and then progresses within microseconds through the protein to the gating mechanisms that open the RyR1 pore and allow Ca2+ to flow into the cytoplasm and activate the contractile machinery. We first consider factors that are essential for EC coupling. 1.1. Membrane geometry The correct geometry of the muscle membranes is one essential component of EC coupling. This geometry is such that the plasma membrane or its transverse (t-) tubule invaginations come into very close association with the membranes of the internal sarcoplasmic reticulum (SR) Ca2+ store (Fig. 1). In regions of functional coupling, the junctional gap between the surface and SR membranes is only ∼10 nm wide and thus similar to the thickness of the membranes that form its borders. This narrow width means that the cytoplasmic domains of proteins in the membrane on either side of the junction come into such close proximity that they could interact with each other. In adult skeletal muscle the junctions usually have three components, with terminal expansions of SR on either side of a central transverse t-tubule element, and are thus known as

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triads. However different combinations of SR and t-tubule elements are seen in thin section electron micrographs, from simple diads (one SR and one t-tubule element) to heptads (four SR and three t-tubule elements) or greater ensembles (Takekura et al., 2003). In the heart, diads commonly form between the external surface or T-tubule membrane and terminal expansions of SR. Many recent studies of skeletal EC coupling use genetically modified skeletal muscle cell lines or myotubes from animal models. In these preparations peripheral couplings form on the external surface of the cell and are accessible to freeze-fracture electron microscopy which has been instrumental in defining the characteristic clusters of four ␣1S voltage sensor particles, termed “tetrads” in junctional regions of the surface membrane (Takekura et al., 2004). The arrangement of ␣1S particles within the tetrad reflects the four subunits of RyR1 located in the underlying SR membrane. A term “couplon” has been coined to describe a generic surface (or t-tubule)/SR junction containing the functional groupings of proteins that facilitate EC coupling (Franzini-Armstrong et al., 1999). 1.2. Alignment of essential proteins It has been known for more than 20 years that three proteins are essential for EC coupling, because knockout of any one of the proteins in mammals is embryonic lethal or animals die at birth. These proteins are the skeletal isoforms of the DHPR ␣1S and ␤1a subunits, which are located in surface/t-tubule membrane and cytoplasm respectively, and the RyR1 Ca2+ release channel which is embedded in the membrane of the internal SR Ca2+ store that directly opposes the surface/t-tubule membrane. The proper alignment of DHPR ␣1S and RyR1 on either side of the junctional gap (Fig. 1) is a further requirement for EC coupling as reviewed by Dulhunty et al. (2002) and Beam and Bannister (2010). The membrane spanning ␣1S subunit is the voltage sensor for EC coupling and the DHPR response to surface depolarization (an action potential in the muscle fiber in vivo), triggers opening of RyR1, Ca2+ release from the SR and contraction. A unique property of coupling between the DHPR and RyR1 in skeletal muscle is a bi-directional signaling between the two proteins (Nakai et al., 1996). “Orthograde coupling” is the EC coupling signal that passes from the DHPR to RyR1. However RyR1 also exerts a “retrograde” influence on the ␣1S subunit which enhances Ca2+ flow across the surface/t-tubule membrane (Nakai et al., 1996). Orthograde skeletal coupling is in itself unique, and in contrast to that in the heart and in many other excitable cells, it does not require a Ca2+ current through the DHPR. Skeletal EC coupling is defined as being independent of an influx of extracellular Ca2+ . In the heart, an influx of Ca2+ from the extracellular media, through the DHPR, is essential because cardiac RyR2 opening is triggered by a Ca2+ -dependent activation process. In the healthy heart, the origin of that Ca2+ is the current that flows through the DHPR. The fact that EC coupling in skeletal muscle occurs in the absence of an influx of external Ca2+ , as well as the lack of evidence for any other second messenger, has led to the conformational coupling hypothesis (Dulhunty et al., 2002; Beam and Bannister, 2010). Conformational coupling is strongly supported by structural evidence. In freezefracture replicas of plasma membrane, particles identified as DHPR molecules are clustered in groups of four (Fig. 1). The frequency and arrangement of tetrads and the spacing between the four DHPR particles reflect the geometry of the underlying RyR tetramer (viewed in thin section electron-microscopy and reconstructions from cryoelectron microscopy particle analysis). The tetrad distribution of particles, first described in fish, suggested that each DHPR is in register with one of the four subunits of every second RyR (Block et al., 1988). The tetrad:RyR ratio may be less in mammals as binding studies indicate that the DHPR:RyR ratio is ∼1:4 in extensor digitorum longus, or 1:8 in soleus muscle from 14 month old rats and

decreases with age to ∼1:12 in 28 month rat soleus (Renganathan et al., 1997). In tetrad-coupled RyRs, the precise geometric arrangement between the four DHPRs and the four subunits of the RyR is such that it would allow a physical interaction between the proteins. The mechanism that opens the uncoupled RyR1 during EC coupling is debatable, but could be either Ca2+ -activated Ca2+ release (Stern et al., 1997), or conformational coupling between neighboring RyRs (Yin et al., 2005). Also in support of the conformational coupling hypothesis is that, in knockout, chimeras or mutation studies with the three essential proteins (␣1S and ␤1a subunits of the DHPR and RyR1), abolition of skeletal EC coupling is generally accompanied by disruption of the tetrad structure. In these situations, large DHPR-like particles can be seen in regions of the freeze-fractured surface membrane that likely correspond to junctions, but clusters of groups of four are not seen. Conversely manipulations that restore tetrad formation, for the most part restore skeletal EC coupling (Takekura et al., 1994, 2004; Protasi, 2002; Schredelseker et al., 2005, 2009; Sheridan et al., 2006). There are of course exceptions to this general rule discussed in Section 3 below. The fact that both tetrad structure and EC coupling are frequently altered in parallel by genetic manipulations does complicate the interpretation of the results. This is because it is not clear whether the particular modification interrupts the geometrical arrangement of the proteins or because the manipulated residues or proteins contribute functionally to the information flow during EC coupling or both (Schredelseker et al., 2009).

2. Excitation contraction coupling proteins As is evident from the discussion in the previous section, the involvement of specific proteins, or residues within proteins, in EC coupling can be verified only in studies of voltage-activated contraction and Ca2+ release in intact myotubes. However whole cell studies do not reveal the molecular nature of the interactions between the individual proteins, their binding affinities or the gating properties of the single RyR channels. This information requires biochemical or electrophysiological approaches using the isolated proteins. While providing detailed information, these studies in turn suffer from the disadvantage that the proteins are removed from their native environment and cannot be subjected to their physiological activation process. Studies of the isolated proteins can reveal likely protein–protein interactions or interaction sites, which must then be verified in intact cells. Complementary data from both approaches is required to provide a detailed mechanistic picture.

2.1. The dihydropyridine receptor and ryanodine receptor The DHPR is composed of five subunits: the transmembrane ␣1 subunit, the membrane associated ␥ and ␦ subunits; the extracellular ␣2 subunit that is disulphide linked to the ␦ subunit and the intracellular ␤ subunit (Fig. 3 below). It has been clearly demonstrated that skeletal EC coupling requires the skeletal isoforms of both the ␣1 subunit (␣1S ) and the ␤ subunit (␤1a ) (Nakai et al., 1996; Gregg et al., 1996; Neuhuber et al., 1998; Coronado et al., 2004). The RyR1 ion channel protein is a homotetramer (Fig. 1) with specific isoforms expressed in skeletal muscle (RyR1 and RyR3) or the heart (RyR2). The three isoforms are products of three different genes, namely RyR1, RyR2 and RyR3. There are also tissue-specific or developmentally regulated splice variants of each gene. Skeletal EC coupling depends on expression of the ␣1S , ␤1a and RyR1 isoforms (Nakai et al., 1997).


Fig. 2. The ␣1a and ␤1a subunits of the DHPR and their likely interactions with one subunit of RyR1. The ␣1a subunit contains four membrane spanning repeats each containing six transmembrane helices in addition to a “pore loop”. Helices 5 and 6 line the pore, while helix 4 contains positively charged residues which allow it to “sense” the voltage across membrane and to respond to changes in voltage. The four repeats are connected by cytoplasmic loops for which some specific functions have been identified. The ␤ subunit contains a binding site for the I–II loop known as the beta interaction domain (BID). In skeletal muscle, the II–III loop of the ␣1S isoform contains sequences that are specific for the retrograde and orthograde interactions in skeletal EC coupling and for targeting the DHPRs into a tetrad formation which allows interactions between the DHPR and RyR. The III–IV loop also interacts with the RyR. Mutations in this loop lead to a malignant hyperthermia (MH) phenotype in which RyR1 is hypersensitive to activating stimuli (Weiss et al., 2004). Finally the C-terminal tail of the ␣1S subunit interacts with RyR1 (Tang et al., 2002). The distal C terminus of ␤1a (C-term) binds to RyR1 and the SH3 domain which can potentially interact with polyproline residues in the proximal C-terminal region and in the critical region of the II–III loop. The “?” is included to indicate other interactions that might yet to be specifically defined.

2.1.1. The dihydropyridine receptor alpha subunit Ten genes encode the various isoforms of the ␣1 subunit of the DHPR proteins and these are expressed in a range of excitable tissues. The ␣1 subunit is a canonical voltage gated ion channel with four membrane inserted repeats, each containing six transmembrane helices, the fourth of which (S4) has a number of positively charged residues which align with the membrane field and facilitate a voltage sensor function (Fig. 2). The role of the ␣1S subunit in skeletal EC coupling has been reviewed in detail (Dirksen, 2002; Protasi, 2002; Dulhunty et al., 2002; Dulhunty, 2006; Bannister, 2007; Beam and Bannister, 2010) and is outlined briefly here as background for the subsequent discussion. The ␣1S subunit contains isoform specific sequences that are critical for the extracellular Ca2+ -independent skeletal EC coupling. These sequences are located in the central portion of the cytoplasmic II–III loop which links the 2nd and 3rd repeats of the protein. Chimera studies with skeletal sequences inserted into a cardiac DHPR background have allowed the identification of a “critical region” for skeletal EC coupling between residues 720 and 765 (Nakai et al., 1998; Grabner et al., 1999; Beam and Bannister, 2010). Indeed skeletal EC coupling is suppressed by substitution of only four residues within this region with divergent residues found in equivalent positions in the cardiac loop (Kugler et al., 2004). The II–III loop, including the “critical region”, is intrinsically unstructured (Cui et al., 2009). The unstructured nature is consistent with a hypothesis that suggests that the loop could undergo a rapid structural change following charge movement within the voltage sensor, and thus underpin a rapid conformational coupling. The conformational coupling hypothesis has been tested in structural studies in which structural changes in RyR1 after treatment with ryanodine alters the arrangement of DHPR’s in tetrads (Paolini et al., 2004). A 2 nm shift in the ␣1S spacing within the tetrad indicated that the ␣1S –RyR1 complex acted as a physical unit. Further structural studies of ␣1S distribution are outlined below (Section 4 below). Other approaches have been utilized to identify the “critical region” and to verify the physical coupling hypothesis. For example fluorescent markers inserted into various parts of the loop

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Fig. 3. Components of the triad junction and couplon RyR1 complex. Two subunits of RyR1 are shown, each with their associated DHPR consisting of 5 subunits of which ␣1S binds to and ␤1a binds to RyR1. The positions of MG29 and Stac3 are hypothetical. On the other hand evidence for the locations of the other proteins is presented in the text. JP-45, junctin, junctate and Trisk95 (skeletal triadin) and JP1 are inserted into the SR membrane and interact either with cytoplasmic components including the RyR and DHPR, or with SR luminal proteins including CSQ and HRC. The red dots in the SR lumen indicate Ca2+ that is bound to CSQ and HRC.

confirm the location of the critical region (Bannister et al., 2009; Beam and Bannister, 2010). Skeletal EC coupling is abolished by streptavidin binding to the ␣1S II–III loop, indicating that the binding of this large molecule prevents a depolarization-dependent conformational interaction that is required for skeletal EC coupling. In contrast, neither streptavidin binding to the N- or C-termini of ␤1a , nor to the N terminus of ␣1S affects skeletal EC coupling (Lorenzon and Beam, 2007). Fluorescence resonance energy transfer (FRET) indicates that the presence of RyR1 causes reorientation of multiple cytoplasmic domains of the ␣1S subunit including the critical region of the II–III loop (Polster et al., 2012). Curiously, the crowded junction between the DHPR and RyR1 (containing the cytoplasmic domains of the four ␣1S subunits with their N- and Cterminal tails, I–II, II–III and III–IV loops and the four ␤1a subunits, not to mention the massive cytoplasmic domain of RyR1), as well as the many other junctional-associated proteins (Fig. 3 below), extensively reviewed in (Hwang et al., 2012; Treves et al., 2009) is accessible to large molecules. Streptavidin has some access to regions around the C- and N-terminal ends of the ␤1a subunit and to the C- and N-terminal part of a ␣1S as well as the II–III loop (Lorenzon et al., 2004; Lorenzon and Beam, 2007). 2.1.2. The dihydropyridine receptor beta subunit Four genes encode various isoforms of the ␤ subunit of the DHPR and express soluble cytoplasmic proteins that belongs to the MAGUK (membrane-associated guanylate kinases) family of structural proteins (Karunasekara et al., 2009). The protein contains five domains, with two conserved structural domains: an Src homology 3 (SH3) – like domain and a guanylate kinase (GK) domain, which have both discrete as well as interdependent functions. The two domains interact with each other within each molecule and their interaction is important for the ability of the ␤ subunit to influence the kinetics of the Ca2+ current through the channel but not for its chaperone role (Chen et al., 2009). The large central GK domain is highly conserved between the different isoforms of the ␤ subunit and it binds with high affinity to the alpha interaction domain (AID) in the ␣1 subunit. The residues involved in this high affinity binding between the ␤ and ␣1 have been clearly defined and the structures of the domains with the proteins bound have been solved (Van Petegem et al., 2008). The high affinity ␣1S ␤1a interaction is necessary for ␤1a targeting to triads (Neuhuber et al., 1998). In contrast, targeting of ␣1S to triads depends primarily on residues in the ␣1S C-terminal domain, but requires ␤1a to be bound to ␣1S (Flucher et al., 2000). Ca2+ currents and charge movement are greatly reduced in myotubes from ␤1a knock-out mice, indicating a reduced ␣1S surface expression

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(Strube et al., 1996). Deletion of the ␤ interaction domain in the I–II loop of ␣1S results in ␣1S retention in the ER (Neuhuber et al., 1998). When ␤1a is expressed in ␣1S -null myotubes, the protein remains in the cytoplasm (Neuhuber et al., 1998). These observations indicate that the ␣1S ␤1a complex is important in the targeting of DHPRs to the surface membrane and to triad junctions in mammals. Curiously in the ␤1a null zebrafish, the lack of the ␤1a subunit does not prevent junctional surface membrane targeting of the DHPR ␣1S , although expression is reduced, tetrad formation appears to be largely absent (Schredelseker et al., 2005). FRET, cryoelectron microscopy and bimolecular fluorescence complementation studies have attempted to uncover the arrangement of ␤1a subunits within a tetrad cluster. FRET measurements indicate that N- and C-termini of individual ␤1a proteins are separated by 10 nm from each other (Leuranguer et al., 2006). These results are consistent with the idea that the N- and C-termini of ␤1a subunits are directed away from the center of tetrads and thus that the SH3 domain may point to the center of the tetrad and toward the central part of the RyR1 tetramer. Consequently, the C-terminal RyR1-binding domain of ␤1a would be in a position to interact with the clamp regions of the RyR tetramer, which extend further toward the t-tubule membrane than any other part of the RyR, and are in the best position to interact with ␣1S (Samso et al., 2005; Serysheva et al., 2008). The most recent model of the arrangement of ␤1a within DHPR tetrads, based on bimolecular fluorescence complementation, suggests that the N- and C-termini of individual ␤1a subunits are separated by 50% (Cheng et al., 2005). This remaining Ca2+ release is consistent with residual binding. Earlier results showed that deletion of the 35 residue C-terminal tail of ␤1a caused a 5-fold reduction in the voltage-activated Ca2+ transient without altering the Ca2+ current through ␣1S or charge movement (Beurg et al., 1999). We have used affinity chromatography to show that, as suggested by the EC coupling and binding studies, a peptide with the sequence of the C-terminal 35 residues of ␤1a can pull down the full length native RyR1 (Rebbeck et al., 2011). In addition, both the full length ␤1a subunit and the C-terminal peptide increase RyR1 channel activity at a half maximal concentration of 450-600pM (Rebbeck et al., 2011). We found that the C-terminal 35 residues of of ␤1a adopt a nascent ␣-helix in which 3 hydrophobic residues align to form a hydrophobic surface that binds to RyR1. Mutation of the hydrophobic residues (L496, L500, W503), reduces peptide binding to RyR1 by ∼85% and prevents activation of RyR1 channels in bilayers (Karunasekara et al., 2012). Earlier mammalian EC coupling studies implicated an upstream hydrophobic heptad repeat in ␤1a C-terminal tail binding to RyR1 (Sheridan et al., 2004). Later work suggested that this was not the case in the zebrafish model, where EC coupling was independent of the heptad repeat residues (Dayal et al., 2010). Consistently, we found that the hydrophobic heptad repeat did not contribute to RyR1 activation by the ␤1a Cterminal domain (Karunasekara et al., 2012). It will be important to revisit the effect of the heptad repeat on mammalian voltage activated Ca2+ release EC coupling before its contribution to EC coupling can ruled out. However these consistent reports from voltage-activated Ca2+ release in intact cells, as well as from biochemistry and lipid bilayer electrophysiology suggest that the ␤1a subunit may be intimately involved in the EC coupling process, and that its involvement is facilitated by the high affinity binding of three hydrophobic residues in the ␤1a C-terminal tail to RyR1 residues including K3495 KKRR · R3502 . However, it is unlikely that the K3495 KKRR · R3502 residues form the binding partner for the hydrophobic surface in ␤1a . It is most likely that the charged sequence is intimately involved in a hydrophobic binding pocket within RyR1.

2.1.4. Update on dihydropridine receptor beta subunit contribution to excitation–contraction As discussed, early experiments in mammalian myotubes provided compelling evidence for a role of ␤1a in skeletal EC coupling (Beurg et al., 1999; Cheng et al., 2005; Coronado et al., 2004; Sheridan et al., 2004; Strube et al., 1996). Recent experiments indicate that the ␤1a -null zebrafish mutant “relaxed” lacks skeletal EC coupling and has reduced expression of the ␣1S , lacks charge movement and DHPR tetrad structures in the freeze-fractured surface membrane (Schredelseker et al., 2005). The relaxed mutant has been used to determine which functions could be restored by transfection with ␤ isoforms in general and which were specific for transfection with ␤1a (Schredelseker et al., 2009). Expression of ␣1S and its targeting to junctional membrane, as well as charge movement, were restored by all ␤ isoforms examined, which ranged from the mammalian skeletal ␤1a , or the closely related mammalian cardiac/neuronal isoform (␤2a ), to the distantly related housefly (M. domestica) isoform (␤M ). In contrast, tetrad formation and skeletal EC coupling were restored only with the skeletal ␤1a isoform. The authors concluded from this study that EC coupling could not proceed because ␣1S was not correctly positioned to communicate with RyR1 with ␤ isoforms other than ␤1a , and that ␤1a is a key requirement for proper alignment of the two proteins. It is worth noting however that tetrad formation does not necessarily go handin-hand with skeletal type EC coupling as ␤3 expression restores


tetrad structures but not EC coupling (Dayal et al., 2013). See also Section 4 below. The functional importance of the individual ␤1a domains in EC coupling has been elegantly dissected by systematically swapping ␤3 domains with those from ␤1a (Dayal et al., 2013). This demonstrated that both the SH3 and ␤1a C-terminal domains were required to fully restore charge movement and intracellular Ca2+ release in the “relaxed” model. From this work it was suggested that a putative SH3 C-terminal interaction (via a PXXP) motif could be important for voltage-sensing, supporting an earlier hypothesis regarding the role of the SH3 domain (Karunasekara et al., 2009). However, when the available crystal structures of the core of cardiac ␤2a and neuronal ␤3 are extrapolated to ␤1a (Van Petegem et al., 2004), the SH3 domain is occluded and is unlikely to interact with polyproline sequences. More direct experimental evidence of a polyproline/SH3 domain interaction is required before a compelling argument can be mounted for its involvement in EC coupling. 2.1.5. The binding site on the ryanodine receptor for the dihydropyridine receptor beta subunit The K3495 KKRR · R3502 sequence in RyR1 that has been implicated in ␤1a binding (Section 2.1.3) is separated by only 9 residues from the Alternatively Spliced, ASI (3481 DAQSG3485 ) residues, and overlaps with the greater ASI region domain (T3471 -G3500 ) that substantially influences RyR1 activity (Kimura et al., 2007). The absence of the ASI residues in the juvenile isoform of RyR1 (ASI(−)) causes an increase in amount and rate of Ca2+ released from the SR following depolarization, without substantially changing the amount of stored Ca2+ (Kimura et al., 2009). Significantly, the juvenile ASI(−) RyR1 isoform is over-expressed in adults suffering from myotonic dystrophy type 1 (DM1) and the resultant abnormal Ca2+ handling by this inappropriate isoform is likely to contribute to the myopathy seen in DM1 patients (Kimura et al., 2005). The greater ASI region, including the K3495 KKRR · R3502 residues, is thus a hot-spot for the regulation of RyR1 channel opening and for the gain of EC coupling, and it is significant that the ␤1a subunit may also bind in this region. As mentioned previously, an essential aspect of skeletal EC coupling is the proper alignment of ␣1S and RyR1. Any disruption of the alignment can abolish EC coupling, as occurs when the ␤1a subunit is not expressed (Neuhuber et al., 1998; Coronado et al., 2004; Schredelseker et al., 2005, 2009) and with removal of other triadic proteins (Section 3). The difficulty in separating an alignment role from the transduction role has hampered progress in identifying the protein interactions that play a role in conveying the depolarization signal from ␣1S to RyR1 once the proteins are correctly tethered. 2.1.6. Stac3 The new kid on the EC coupling block is Stac3. First reported in 2012 (Bower et al., 2012), the protein was identified as a nutritionally regulated gene from an Atlantic salmon subtractive hybridization library with highest expression in skeletal muscle where it is required for myogenic differentiation. Stac3 is a 360 residue protein that contains an SH3 domain and cysteine rich domain. There are no reports of the structure of the entire protein or its membrane association. It is localized to the t-tubule system, but it is not clear if it occupies the entire t-tubule or is confined to triads (Nelson et al., 2013). Nor is it known whether Stac3 spans the membrane or is localized on the external or cytoplasmic surface. Stac3 participates in EC coupling in zebrafish muscles and a mutation in human Stac3 is linked to the native American myopathy (Horstick et al., 2013). Homozygous Stac3 knockout mice die at birth (Reinholt et al., 2013). Contractility and Ca2+ release in cultured myotubes from Stac3 knockout mice is restored by 4chloro-m-cresol, indicating that the SR Ca2+ store and RyR1 are

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functional, but that EC coupling is disrupted (Nelson et al., 2013). It is possible, based on evidence thus far that Stac3 plays an active role in signal transmission during EC coupling. However it is equally possible that it is a structural protein that helps maintain the integrity of the T-SR junction and thus maintains the function of the couplon. The potential role of Stac3 in EC coupling is a significant finding and its role in EC coupling will be revealed with further characterization of the protein and its interaction with other proteins in the triad junction. 3. Additional triadic proteins that influence excitation–contraction coupling 3.1. Major junctional proteins Three proteins, apart from the DHPR and RyR1, are present in significant amounts in the triad junction and influence skeletal EC coupling. These are calsequestrin (CSQ), the predominant Ca2+ binding protein in the SR lumen, and two SR membrane inserted proteins, triadin and junctin, that bind to RyR1 and to CSQ (Fig. 3). The sarcoplasmic, endoplasmic reticulum Ca2+ ATPase (SERCA) also appears in substantial quantities in junctional face membrane fractions. SERCA regulates the amount of Ca2+ available for release and can thus influence the amplitude of voltage-activated Ca2+ release. However there is no evidence to suggest that it plays a role in the chain of events linking the voltage sensor to RyR1 activation in skeletal EC coupling and will not be considered further here. 3.1.1. Calsequestrin CSQ influences depolarization-induced Ca2+ release in two ways. Firstly through a direct action on the open probability of RyR1, that proceeds though junctin (Wei et al., 2009a,b; Protasi et al., 2011) in a manner that is yet to be fully defined. We recently showed that triadin was not involved in transmitting signals from CSQ1 to RyR1, while junctin alone served this function in RyR1 channels in lipid bilayers where RyR1 is not under the influence of the DHPR (Wei et al., 2009a,b). Secondly, the Ca2+ binding capacity of CSQ also determines the Ca2+ load that can be maintained within the SR, and thus influences the amount of Ca2+ that can be released during EC coupling (Launikonis et al., 2006). It is notable that CSQ1-null mice develop a phenotype akin to that seen in malignant hyperthermia (Dainese et al., 2009), which has been attributed to an altered coupling between the DHPR and RyR1 (Eltit et al., 2012). The mechanisms underlying the ability of CSQ to influence EC coupling are yet to be clearly defined. 3.1.2. Triadin The two “anchoring” proteins, triadin and junctin have also been reported to impact on EC coupling. Of these two proteins, triadin has been the subject of more investigation and much controversy in recent years. Deletion of three acidic residues in the pore loop of RyR1 not only abolished binding of the major triadassociated skeletal muscle isoform of triadin (Trisk95) to RyR1, but also abolished voltage-dependent Ca2+ release in myotubes expressing the mutant RyR channels (Goonasekera et al., 2007). Our conclusion was that Trisk95 is a major contributor to EC coupling. Subsequent work with knockdown of Trisk95 in skeletal C2C12 myotubes indicated that triadin facilitated, but was not essential for, depolarization-induced Ca2+ release (Wang et al., 2009). Curiously, however, over-expression of Trisk95 also reduces the efficacy of depolarization-induced Ca2+ transients (Rezgui et al., 2005; Fodor et al., 2008). There is major structural rearrangement of triads in skeletal muscles of mice lacking either Trisk95 or junctin or both (Oddoux et al., 2009; Boncompagni et al., 2012). The structural effect of removing Trisk95 could explain the effects on EC coupling. On the other hand, there is evidence that changes in EC coupling

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with Trisk95 knockout are due to dissociation of another regulatory protein and not the absence of Trisk95 (Eltit et al., 2011). Nevertheless the majority of reports indicate a contribution of Trisk95 to EC coupling, although the molecular nature of this contribution is yet to be deciphered. 3.1.3. Junctin Most work addressing junctin’s role in EC coupling has been in cardiac models where junctin is a significant anti-arrhythmic factor (Yuan et al., 2007; Fan et al., 2008). We suggest that this depends on junctin’s role in anchoring cardiac CSQ2 to the cardiac RyR2, because the effect of junctin knockout on luminal Ca2+ sensitivity (Altschafl et al., 2011) reflects the effect of CSQ dissociation on RyR1 activity (Dulhunty et al., 2012). The role of junctin in skeletal muscle has received little attention and results are inconsistent. Junctin in C2C12 myotubes appears to maintain SR Ca2+ store size (Wang et al., 2009), possibly through its interaction with CSQ1 (Wei et al., 2009a,b). Conversely, studies in junctin-null mice indicate that interactions between junctin and CSQ have little influence on junctional SR structure or Ca2+ cycling (Boncompagni et al., 2012). From a different perspective, reversible polymerization of CSQ2 requires junctin and is critical for SR Ca2+ homeostasis (Lee et al., 2012). It is likely that junctin does play a part in skeletal EC coupling because (a) both CSQ1 and CSQ2 are expressed in skeletal muscle (Paolini et al., 2011), (b) CSQ1 polymerizes at lower luminal Ca2+ concentrations than CSQ2 (Wei et al., 2009a,b) and (c) reversible polymerization of CSQ is implicated in skeletal EC coupling (Launikonis et al., 2006). However the molecular nature of junctin’s contribution remains to be determined. 3.2. Minor junctional proteins An increasing number of proteins are being discovered that are localized to the skeletal muscle t-tubule/SR junctions and appear to influence EC coupling (Fig. 3) and extensively reviewed in (Treves et al., 2009). The most notable of these proteins are summarized in the following sections. 3.2.1. Histidine rich Ca2+ binding protein (HRC) HRC is Ca2+ binding protein located within the SR. Although first identified in skeletal muscle, most information on HRC comes from cardiac muscle where a S96A mutation in HRC is associated with ventricular arrhythmia (Arvanitis et al., 2008). HRC binds to the cardiac isoform of triadin (triadin-1) at in vivo luminal [Ca2+ ]s of ∼0.1–1.5 mM, and also binds to SERCA although more so at nonphysiological luminal [Ca2+ ]s of 0.01–1.0 ␮M (Arvanitis et al., 2007). Nevertheless, overexpression of HRC in the heart disrupts SR Ca2+ uptake and depresses relaxation possibly via the interaction with SERCA2a (Fan et al., 2004). Knockout of HRC dysregulates SR Ca2+ cycling and a severe pathology under pressure-overload cardiac stress suggests an essential role in maintaining cardiac function through interactions with SERCA (Park et al., 2013). The role of HRC in skeletal muscle EC coupling has not yet been explored. However, as HRC binds to Trisk95 (Lee et al., 2001; Sacchetto et al., 2001), and Trisk95 is implicated in EC coupling (Section 2.1.2 above), HRC may well communicate intra-SR Ca2+ load to the RyR1 during EC coupling through Trisk95. 3.2.2. Junctophilins (JPs) There are three junctophilin genes: skeletal muscle contains both JP-1 and JP-2 mRNA, the heart contains JP-2 mRNA, and the brain contains JP-3 mRNA (Nishi et al., 2000). JPs bind to the ttubule membrane through N-terminal MORN motifs (membrane occupation and recognition nexus domains) and bind to the SR membrane via their C-terminal transmembrane domain. JP1 is coimmunoprecipitated with RyR1 (Phimister et al., 2007), while JP1

and JP2 interact with L-type Ca2+ channel (DHPRs) in skeletal muscle (Golini et al., 2011). JPs contribute to maintaining the structural integrity of the T-tubule/SR junction and to appropriate apposition of the DHPR and RyR1. This is suggested by the fact that proteolysis of JP1 and JP2 lead to separation of the membranes and disrupt skeletal EC coupling (Murphy et al., 2013). RyR2 coimmunoprecipitates with JP2 in myocyte lysates (van Oort et al., 2011) and co-localizes with RyR2 in nanoscale confocal microscopy (Jayasinghe et al., 2012). This is likely to also be the case with skeletal isoforms, as JP1 binds to RyR1 and binding is enhanced by oxidizing conditions (Phimister et al., 2007). The fact that JPs interact with both the DHPR and RyR1 raises the possibility that they also contribute directly to the flow of depolarization information from the DHPR to RyR1. 3.2.3. Mitsugumin-29 The 29 kDa membrane protein, MG29, is found in intracellular membranes of skeletal muscle, kidney and brain (Shimuta et al., 1998; Satoh et al., 2012). The protein contains four transmembrane domains and is located in the t-tubule membrane of mature skeletal muscle fibers (Shimuta et al., 1998; Brandt et al., 2001). A functional association between MG29 and RyR1 was discovered when the proteins were co-expressed in Chinese hamster ovary cells (Pan et al., 2004). Co-expression led to cell death due to depletion of intracellular Ca2+ stores. MG29 added to RyR1 channels in lipid bilayers, substantially increased channel activity. Muscles from MG29 knock out mice are characterized by swollen t-tubules, poorly aligned triads and vacuolated SR (Nagaraj et al., 2000; Pan et al., 2002) and demonstrate enhanced fatigability. The SR Ca2+ store in these muscles is rapidly depleted and refills more slowly than in muscle from wild type mice (Pan et al., 2002). These studies indicated a role for MG29 in store operated Ca2+ entry (SOCE), which may be simply to retain the normal triad structure (Treves et al., 2009), or may involve a more complex association with other components of SOCE, such as STIM1 and Orai1. The functional significance of MG29 interactions with RyR1 and its potential role in EC coupling and/or in the store-independent Ca2+ entry pathway, remains to be determined. 3.2.4. JP-45 The 45 kDa junctional protein, JP-45, is found in skeletal muscle junctional face membrane (Anderson et al., 2003). JP-45 contains a single transmembrane segment and binds to calsequestrin via its luminal C-terminal domain and to the DHPR ␣1S subunit through its cytoplasmic N-terminus (Anderson et al., 2003). JP-45 elutes from heparin-agarose columns with RyR1 (Zorzato et al., 2000), perhaps through its association with CSQ1 which binds to RyR1 via junctin and possibly triadin (Section 3.1.1 above). JP-45 binds to the C-terminus of the DHPR ␣1S and to a region within the I–II loop that also binds the ␤1a subunit (Anderson et al., 2006). This association raises the possibility that JP-45 may assist in targeting ␣1S into the surface membrane and perhaps in tetrad formation. Curiously, there are parallel increases in DHPR ␤1a subunit and JP-45 expression in aging mammals along with a decline in ␣1S expression (Delbono, 2011). The various associations with JP-45 would allow it to facilitate signaling between CSQ1 and the DHPR and to modulate EC coupling in response to Ca2+ levels in the SR. Indeed a role of JP-45 in EC coupling is indicated (a) by its overexpression, which suppresses voltage-activated Ca2+ transient without altering the DHPR Ca2+ current (Gouadon et al., 2006) and (b) a JP-45 variant which alters EC coupling by reducing the sensitivity of the ␣1S voltage sensor to depolarisation (Yasuda et al., 2013). 3.2.5. Junctate The genetic and structural relationship to junctin and to aspartyl-␤-hydroxylase has stimulated interest in the 33 kDa


junctate. It is a moderate affinity, high capacity Ca2+ binding protein (Treves et al., 2000) which has one ER/SR membrane spanning domain and is expressed in a range of tissues including the heart and skeletal muscle (Treves et al., 2000; Dinchuk et al., 2000; Hong et al., 2001). Junctate is a transcript of the BAH gene which encodes four different proteins through complex alternative splicing (Treves et al., 2009). These proteins are: (i) the aspartyl-␤-hydroxylase enzyme; (ii) humbug; (iii) junctin (3.13 above); and (iv) junctate. Humbug is a truncated form of aspartyl-␤-hydroxylase, lacking a catalytic domain, but has with Ca2+ binding properties similar to those of junctate (Treves et al., 2000; Dinchuk et al., 2000; Hong et al., 2001). Junctate over-expressed in HEK cells causes a proliferation of ER/plasma membrane couplings, where junctate is associated with inositol 1,4,5-trisphosphate receptors and TRPC channels. SOCE in these cells is enhanced and is influenced by the short cytoplasmic N-terminal domain of junctate, while Ca2+ stores are regulated by the luminal C-terminal Ca2+ binding domain (Treves et al., 2004, 2009). In addition, junctate is a structural component of ER junctions with the plasma membrane in T cells where Orai1 and STIM1 cluster (Srikanth et al., 2012). Junctate over-expression in skeletal muscle leads to a small increase in Ca2+ store load and larger increases in Ca2+ release through RyR1 and in SOCE (Divet et al., 2007). Junctate binding to RyR1 has not been reported, but remains possible. Furthermore, since TRPC3 can bind to RyR1 (Kiselyov et al., 2000), junctate may interact with RyR1 in a junctate/TRPC3/RyR1 complex.

4. Additional regions of the ryanodine receptor that influence skeletal excitation–contraction coupling The chimera approach that has been so successful in identifying the “critical region” in the ␣1S II–III loop for EC coupling has so far failed to reveal any discrete residues in RyR1 that are “essential” for skeletal EC coupling, although numerous lengthy sequences containing several hundred residues have been found to contribute to the coupling process. Chimeras have been constructed with regions of the cardiac RyR2 replaced by corresponding regions of RyR1. The chimeras have been expressed in “dyspedic”, RyR1null, myotubes. An initial study showed that both EC coupling and retrograde signaling were restored by skeletal residues 1635–2559, while retrograde signaling alone was restored by skeletal residues 2659–3730 (Nakai et al., 1998). Two hybrid studies reveal a weak interaction between the “critical region” of the II–III loop and skeletal residues 1835–2154 (Proenza et al., 2002). Disappointingly however a RyR2/RyR1 chimera with the skeletal sequence in residues 1835–2154 demonstrated only weak skeletal EC coupling (Proenza et al., 2002). Subsequent studies using a RyR1-null mouse skeletal cell line (1B5) transfected with RyR2/RyR1 chimeras found that the overlapping skeletal residues 1635–3720 restored both EC coupling and tetrad formation (Protasi et al., 2002). Curiously, skeletal residues 1635–2559 were more effective in restoring EC coupling than tetrads, while skeletal residue 2659–3720 were more effective in restoring tetrads than skeletal EC coupling. Finally skeletal residues 1837–2154 restored EC coupling to a greater degree than tetrad structures. The conclusion from this study is that that multiple regions of RyR1 may interact with ␣1S /␤1a and that the regions responsible for tetrad formation do not correspond exactly to the ones required for functional coupling. In the same vein, a later study found that RyR1/RyR3 constructs having RyR1 residues 1–1681, restored DHPR tetrad arrays to wild type numbers but only partly restored skeletal EC coupling and failed to enhance DHPR Ca2+ currents in retrograde signaling (Sheridan et al., 2006). It was noted that the divergent D2 domain (RyR1 residues 1272–1455) was instrumental in tetrad formation and EC coupling

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when expressed together with constructs that were alone only partially effective (Sheridan et al., 2006). This study indicated again that both structural and functional aspects of skeletal EC coupling are influenced by several broad regions of RyR1. It was hoped that point mutations in RyR1 associated with “core” myopathies might point to RyR1 residues that support EC coupling. Such mutations interrupt Ca2+ release in response to depolarization through a variety of mechanisms including effects on RyR1 Ca2+ permeability, effects on RyR1 open probability, as well as effects on the EC coupling process (Hwang et al., 2012; Van Petegem, 2012; Lanner et al., 2010; Dirksen and Avila, 2002; Kimura et al., 2007). Thus the mutated residues are often those that are important in setting aspects of RyR1 function in general, rather than those involved in receiving the signal from the DHPR. Although some of these mutations, and other mutations unrelated to disease, do impact directly on EC coupling (Dirksen and Avila, 2002; Kimura et al., 2007; Goonasekera et al., 2007), they have not thus far revealed individual residues that bind to the “critical region” in the ␣1S II–III loop. Smaller regions of RyR1 that bind to parts of the recombinant ␣1S II–III loop in vitro have been identified in regions encompassing residues 1076–1112 (Leong and MacLennan, 1998; Altafaj et al., 2005) which overlaps with second of three SPRY domains (SPRY2) in RyR1 S1085-V1208 (Cui et al., 2009; Tae et al., 2009a,b). SPRY domains are structural ␤-sandwich domains that are generally involved in protein–protein interactions and were first identified in, and named after, the fungal Dictyostelium discoideum tyrosine kinase spore lysis A (SplA) and mammalian RyRs (Tae et al., 2009a,b). The RyR1 SPRY 2 domain is contained within residues 1–1681 that are implicated in DHPR tetrad arrays (Sheridan et al., 2006). However the N-terminal part of the II–III loop that is primarily involved in binding to the SPRY2 domain in vitro (Cui et al., 2009) is not directly involved in EC coupling (Wilkens et al., 2001) and mutation of three critical II–III loop binding residues in RyR1 does not alter EC coupling (Tae et al., 2011). Nevertheless, the SPRY2 domain is located on the clamp domain (Peralvarez-Marin et al., 2011), close to the T-tubule membrane and DHPR tetrads. Thus it remains possible that the SPRY2 domain contributes to EC coupling through protein–protein interactions that are yet to be identified. It is worth noting that early attempts to show a direct interaction between isolated RyR1 and DHPR ␣1S proteins using protein overlay and affinity chromatography techniques failed (Brandt et al., 1990). Similarly, cross-linking studies have either failed to show an association between the two proteins (Shoshan-Barmatz et al., 1995) or demonstrated a low level of interaction (Murray and Ohlendieck, 1997). Curiously however, in the latter study, the ␤1a subunit was not immune-detected with the ␣1S /RyR1 complex. This was surprising given the established high affinity interaction between ␣1S and ␤1a (see Section 2.1.2 above) and the presence of ␤1a in the preparations. The apparent lack of ␤1a in the cross-linked ␣1S /RyR1 complex was attributed by the authors to alterations in the antibody binding site as a result of the cross-linking. Thus it is likely the DHPR␣1S subunit and RyR1 in the high molecular weight complex were in fact linked through ␤1a . It is also likely that other junctional proteins (Section 2.2 above) with scaffolding and/or transduction roles were present in the protein complex.

5. Conclusions Although the ␣1S subunit II–III loop and RyR1 are essential for skeletal EC coupling, there is no evidence that they bind to each other in vivo. The coupling process is likely to be through other proteins and there is emerging evidence that a part of this coupling is in fact through the DHPR ␤1a subunit (Beurg et al., 1999; Cheng et al., 2005; Coronado et al., 2004; Karunasekara et al., 2009,

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2012; Rebbeck et al., 2011). This possibility is supported by the fact that small clusters of residues involved in binding of ␤1a to RyR1 have been identified in the C-terminal tail of the ␤1a subunit and in RyR1 (Karunasekara et al., 2012; Rebbeck et al., 2011). However, at this stage the relationships between the ␣1S II–III, the ␤1a subunit and the RyR1 K3495 KKRR · R3502 residues remain unclear. Indeed, many questions remain, including the degree to which any ␤1a /RyR1 interaction may complement other potential interactions between ␤1a , ␣1S and RyR1. A component of skeletal EC coupling remains when the ␤1a C-tail/RyR1 interaction is disrupted (Beurg et al., 1999), indicating that there are other contributing interactions. Are these interactions between other regions of ␤1a and RyR1 or between ␣1S and RyR1 or through yet to be identified pathways? The involvement of multiple domains of RyR1 in the bidirectional coupling process (Sheridan et al., 2006) is indicative of an extraordinarily complex interaction process and it is clear that the details of these interactions provide a continuing challenge. It is possible that (a) an essential link in the chain of events linking the II–III loop to RyR1 remains to be discovered, and/or (b) there is a functional redundancy in the link between ␣1S and RyR1 that ensures survival, if not perfect coupling, when one of the components is compromised.

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Mild cold induced thermogenesis: are BAT and skeletal muscle synergistic partners?

Rem uncouples excitation-contraction coupling in adult skeletal muscle fibers.

Pericytes are Essential for Skeletal Muscle Formation.

Effects of 4-aminopyridine on the excitation-contraction coupling in frog and rat skeletal muscle.

Valinomycin and excitation-contraction coupling in skeletal muscle fibres of the frog.

Charge movement and the nature of signal transduction in skeletal muscle excitation-contraction coupling.

Effects of amrinone on the electromechanical coupling in frog skeletal muscle fibres.

Effect of myoplasmic pH on excitation-contraction coupling in skeletal muscle fibres of the toad.

Skeletal Muscle Microalterations in Patients Carrying Malignant Hyperthermia-Related Mutations of the e-c Coupling Machinery.

Regions of the skeletal muscle dihydropyridine receptor critical for excitation-contraction coupling.

Spontaneous calcium transients manifest in the regenerating muscle and are necessary for skeletal muscle replenishment.

Muscle sounds are emitted at the resonant frequencies of skeletal muscle.

Training-induced skeletal muscle adaptations are independent of systemic adaptations.

Skeletal muscle metaboreceptor exercise responses are attenuated in heart failure.

The dancing heart.

Coupling between skeletal muscle fiber size and capillarization is maintained during healthy aging.

Cardiac-type excitation-contraction coupling in dysgenic skeletal muscle injected with cardiac dihydropyridine receptor cDNA.

Characterization of a multiprotein complex involved in excitation-transcription coupling of skeletal muscle.

[K+]i-mediated excitation-transcription coupling.

Enantiomeric effects on excitation-contraction coupling in frog skeletal muscle by a chiral phenoxy carboxylic acid.

Ca2+-dependent regulations and signaling in skeletal muscle: from electro-mechanical coupling to adaptation.

Membrane Cholesterol in Skeletal Muscle: A Novel Player in Excitation-Contraction Coupling and Insulin Resistance.

Increased optical transparency associated with excitation--contraction coupling in voltage-clamped cut skeletal muscle fibres.

Excitation-contraction coupling in intact frog skeletal muscle fibers injected with mmolar concentrations of fura-2.