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Voltage sensing mechanism in skeletal muscle excitation-contraction coupling: coming of age or midlife crisis?

Voltage sensing mechanism in skeletal muscle excitation-contraction coupling: coming of age or... The process by which muscle fiber electrical depolarization is linked to activation of muscle contraction is known as excitation-contraction coupling (ECC). Our understanding of ECC has increased enormously since the early scientific descriptions of the phenomenon of electrical activation of muscle contraction by Galvani that date back to the end of the eighteenth century. Major advances in electrical and optical measurements, including muscle fiber voltage clamp to reveal membrane electrical properties, in conjunction with the development of electron microscopy to unveil structural details provided an elegant view of ECC in skeletal muscle during the last century. This surge of knowledge on structural and biophysical aspects of the skeletal muscle was followed by breakthroughs in biochemistry and molecular biology, which allowed for the isolation, purification, and DNA sequencing of the muscle fiber membrane calcium channel/transverse tubule (TT) membrane voltage sensor (Cav1.1) for ECC and 2+ of the muscle ryanodine receptor/sarcoplasmic reticulum Ca release channel (RyR1), two essential players of ECC in skeletal muscle. In regard to the process of voltage sensing for controlling calcium release, numerous studies support the concept that the TT Cav1.1 channel is the voltage sensor for ECC, as well as also being 2+ aCa channel in the TT membrane. In this review, we present early and recent findings that support and define the role of Cav1.1 as a voltage sensor for ECC. Keywords: Skeletal muscle, Excitation-contraction coupling, Charge movement, Voltage sensors, DHPR/Cav1.1, 2+ L-type voltage-gated calcium channel, Ca release, RyR1 Background transverse tubular (TT) system [6, 7]. The AP In skeletal muscle, electrical impulses carried by the axons depolarization activates skeletal muscle voltage-gated of motoneurons travel to the nerve endings at the muscle calcium channels (Cav1.1; also known as dihydropyridine endplate (the muscle synapse), where these electrical sig- receptors, DHPR) [8]. The Cav1.1 channels serve as the nals are converted into chemical signals that produce de- voltage sensing machinery for the process of TT polarizing postsynaptic potentials at the neuromuscular depolarization-induced calcium release from the sarco- junction sarcolemma of the muscle fiber [1, 2]. In all but a plasmic reticulum [9] via intracellular sarcoplasmic few “tonic” muscle fibers, these postsynaptic endplate po- reticulum (SR) calcium release channels, the type 1 ryano- tentials elicit a further depolarization of the muscle fiber, dine receptors (RyR1) [10]. This process that begins with carried out by skeletal muscle voltage-gated sodium chan- the muscle AP propagation and results in muscle contrac- nels, initiating and propagating the muscle action poten- tion is known as excitation-contraction coupling (ECC). tial [3–5]. The muscle action potential (AP) travels both This term was coined by Sandow in the 1950s [11]to in- longitudinally away from the fiber endplate along the clude these main events critical for muscle activation, well muscle fiber surface sarcolemma and radially into the fiber before the molecular identities or even the existence of via invaginations of the sarcolemma that form the the molecular players (Cav1.1 and RyR1) was identified or established. Since then we have accumulated an incredible amount of information concerning the structural aspects * Correspondence: mschneider@som.umaryland.edu Department of Biochemistry and Molecular Biology, University of Maryland and molecular and functional details of the ECC process. School of Medicine, 108 N. Greene Street, Baltimore, MD 21201, USA © The Author(s). 2018 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 2 of 20 The knowledge that the muscle contraction was con- discovery and early functional studies of the ECC voltage trolled by electrical signals was already established by sensor and its role and properties, largely carried out the pioneering work of Galvani, Volta, and Walsh [12, during the last quarter of the twentieth century. We 13]. Subsequent studies and discoveries by Nobili, then consider more recent molecular, structural, and Matteucci, Du Bois-Reymond, and Ringer, to mention mechanistic studies, as well as possible future directions. just a few of the pillars, formed the foundations for A detailed review of the cloning of the Cav1.1 and RyR, modern understanding of bioexcitability of muscle and and identification of their skeletal muscle isoforms as 2+ other excitable tissues [14]. The remarkable work by the ECC voltage sensor and skeletal muscle SR Ca re- Hodgkin, Huxley, and Katz [15–19] established the play- lease channel, respectively, is beyond the present scope ing field for the subsequent wave of functional studies and can be found elsewhere [38–42]. dealing with excitability in general, and with the TT volt- age sensor for EEC in particular, as we consider here. Intramembrane charge movement and ECC Voltage sensor charge movements were predicted by Introduction to excitation-contraction coupling Hodgkin and Huxley By the 1950s and 1960s, the processes that initiated and In their classic work on the membrane potential- accompanied skeletal muscle contraction had been stud- dependent ionic conductances underlying the nerve ied from several angles [20, 21]. Muscle biologists and axon action potential, Hodgkin and Huxley [15] pre- physiologists were working collectively trying to de- dicted that any voltage-sensitive process, such as + + cipher the details of the machinery that controls the voltage-dependent Na or K conductance, should be process of muscle contraction using state-of-the-art controlled by mobile charges that are trapped within the techniques from that period. These pioneers made im- membrane but can be displaced in response to changes portant fundamental contributions to understanding in electrical potential energy due to changes in trans- ECC, including the following. (1) Hodgkin and Horowicz membrane voltage. They further predicted that such [22, 23] proposed that the event that normally induces intramembrane charges should give rise to tiny charge muscle contraction is a change in membrane potential displacement currents in response to changes in trans- rather than the longitudinal spread of current along the membrane voltage (Vm), as the putative charges trapped fiber; they also showed that the development of tension within the membrane redistribute within the membrane was dependent on membrane potential and was de- in response to the change in electrical potential. How- scribed by a steep sigmoidal curve of tension as a func- ever, charge displacement currents were not detected by tion of membrane potential. (2) The experiments of Hodgkin and Huxley [15]. In fact, it took over two de- Huxley and Taylor [7] showing activation at the Z disk cades to prove the charge movement hypothesis of in frog muscle fibers and of Huxley and Straub showing Hodgkin and Huxley [15, 37]. In addition, at the time of local activation at the A-I band junction in lizard muscle Hodgkin and Huxley, and continuing well through the [24], together with the localization of the TT system at time of the early experimental studies characterizing the the Z disk in frog muscle [25] and at the A and I band functional properties of voltage sensor charge move- junction in lizard muscle [26], indicated that the trans- ments, the molecular identity of the voltage sensors was verse tubules (TT) of the skeletal muscle fibers form the not even known. Consequently, it was unknown at the network which conducts the surface depolarization radi- time whether the putative and subsequently measured ally into a muscle fiber to initiate contraction. (3) Inves- intramembrane charge movements were generated by tigations started by Ringer [27] and continued by (1) positive charges held near the inside of the mem- Heilbrunn [28], Kamada and Kinoshita [29], and others in- brane at the inside-negative resting potential, moving 2+ troduced the role of Ca as key regulator of striated outward during depolarization (Fig. 1a, right) and muscle activation. Further details of the complex action of returning inward after repolarization (Fig. 1a, left); (2) 2+ Ca on muscle contractile activation were eventually pro- negative charges positioned near the outside of the vided by Weber [30, 31] and Ebashi [32]; reviewed in more membrane at rest, moving inward during membrane detail by Endo [33]. (4) Robertson [26], Andersson-Cedegren depolarization and returning outward during repolariza- [34], Francini-Armstrong and Porter [35], and Peachey tion (Fig. 1a, left); or even (3) dipolar charges rotating in [36], using electron microscopy, described that the ultra- the membrane as the positive end moves outward and structure of transverse tubules (TT) and that the terminal the negative charge moves inward during depolarization cisternae of the SR are in close proximity to the TTs. and reverses this movement during repolarization This year (2018) is the 45th anniversary of the demon- (Fig. 1b). Subsequently, with the establishment of the stration of ECC voltage sensor charge movement [37]. In molecular identity, amino acid sequence, and predicted keeping with the theme of “coming of age/midlife crisis” or experimentally determined molecular structure of the of the ECC voltage sensor, here we will first review the ECC voltage sensor (Cav1.1) [38, 43], as well as those of Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 3 of 20 Fig. 1 Hypothetical mechanisms for a mobile charged intramembrane voltage sensor. a Voltage-dependent intramembrane charge movement could be generated by positive charges moving outward during depolarization, negative charges moving inward, or dipoles rotating. In each, the charge would return to its starting distribution on membrane repolarization, so the charge moved outward during membrane depolarization must equal the charge moving back on repolarization. b Cartoon illustration of current concept of intramembrane voltage sensors as positively charged amino acid residues on transmembrane alpha helices. As in a, the charge moved outward during depolarization (here on an alpha helix) will equal the charge returning on repolarization other voltage-sensitive channels, it is now accepted that were expected to occur over the Vm range where muscle positively charged transmembrane alpha helical “S4” seg- fiber contraction was activated, roughly between about − ments in membrane-spanning domains (Fig. 1b; consid- 50 and + 20 mV [23]. Thus, the putative muscle voltage ered in detail below) are the electrically charged sensor charge displacement current was predicted to be molecular components that serve as voltage sensors for an “extra” non-linear component of the total membrane both ECC and channel gating of plasma membrane and current. Formally, this extra current was “capacitative” in + 2+ + TT Na ,Ca , and K channels [44]. nature since whatever charge moved outward during depolarization was trapped within the membrane and was Voltage sensor charge movements were first detected in obliged to move back to its starting intramembrane loca- skeletal muscle fibers tion when the membrane was repolarized (Fig. 1). To ex- The first successful intentional measurement of the volt- tract the non-linear component from the total measured age sensor charge displacement currents predicted by membrane capacitative current for each “test” pulse (P) Hodgkin and Huxley was carried out on skeletal muscle applied from the holding potential (Fig. 2a,left), the same fibers. Assuming that the charge displacement currents depolarizing pulse (P) was superimposed on a negative would be small, Schneider and Chandler [37] voltage prepulse (ΔV pre) of larger absolute amplitude than clamped frog skeletal muscle fibers in the presence of the test pulse (Fig. 2a,right) [37]. In this way the blockers for each of the major ionic conductances (TTX same amplitude pulse (P) was now applied over a Vm + + + + for Na conductance, Rb replacing K for K conduct- range that was entirely negative to the initial holding ance, methanesulphonate for Cl− conductance). Under potential. This pulse served as the “control” pulse and these conditions, ionic currents were essentially absent. is assumed to contain only the linear capacitative However, the linear capacitative current, needed to current (Fig. 2b,right). charge the linear capacitance of the muscle fiber lipid bi- The currents for both the test and control pulses were layer membrane when the fiber membrane potential was recorded digitally during the experiment using a digital changed, still remained and obscured the putative volt- signal averaging device, which preceded the introduction age sensor charge movement. of laboratory computers and the use of PCs. Using the A strategy was needed for removing the linear capacita- digitized records of total current, the current recorded tive current in order to “unmask” the current carried by for the “control” depolarization (in the range negative to voltage sensor charge movement. The expected voltage the holding potential; Fig. 2b, right) was digitally sensor charge displacement current was anticipated to sat- subtracted from the current for the “test” depolarization urate at highly positive or highly negative membrane po- (over a voltage range positive to the holding potential; tentials, as all mobile charges were maximally displaced Fig. 2b, left) to give the “non-linear” membrane current during large depolarizations or hyperpolarizations (Fig. 1). (Fig. 2c). Assuming any remaining non-linear ionic Furthermore, the voltage sensor charge displacements current to be constant (i.e., time-independent) during Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 4 of 20 cd Fig. 2 Protocol for original recording of intramembrane charge movements. a Pulse protocol used to extract current carried by intramembrane charge movements. The same pulse (P) was applied either from the holding potential (test, left) or superimposed on a hyperpolarizing prepulse (Δ Vpre; control, right). The pulse over the control voltage range is assumed to cause no intramembrane charge movement. b The total current recordings for the pulses in a.Tracesin a and b were recorded as photos of oscilloscope display. c Difference between membrane current in the test pulse minus the current in the control pulse, obtained by digital subtraction of digital recordings of test and control currents using an analog to digital converter and a digital recording system. d Schematic diagram of the three microelectrode voltage clamp systems used in this experiment. All records obtained at the tendon termination of a muscle fiber in a frog sartorius muscle stretched to reduce contraction and bathed in solution to block or remove essentially all membrane conductance. From ref. [37], with modification the pulse, the time-dependent component of the amplitude. In contrast, the time course of the charge non-linear current (test−control) was taken to be the movement current after the pulse (i.e., during fiber repo- non-linear “charge displacement current” or “charge larization to the initial holding potential) did not notice- movement current” due to the voltage sensor movement. ably change in kinetics as the pulse depolarization was This current is the current in excess of the steady increased (Fig. 3a). As discussed further below, from the non-linear current during the pulse (Fig. 2c; upper original report, the charge movement kinetics and voltage dashed line during the pulse), and the current due to the dependence were generally in the range that would be ap- return of the voltage sensors was taken to be the current propriate for muscle contractile activation, so it was not below the initial and final zero current (Fig. 2c; dashed unreasonable to identify these charge movements with the line after the pulse) when the voltage sensors return to intramembrane movement of ECC voltage sensors in the their starting distribution after the pulse. These initial TT membrane [37]. With subsequent sophistication of ex- recordings of voltage sensor currents were made at the perimental and recording procedures, records with better end of a single muscle fiber in an isolated frog sartorius signal to noise and corresponding resolution of kinetic de- muscle using the three microelectrode voltage clamp tails were obtained (a) from frog individual muscle fibers system developed by Adrian et al. (Fig. 2d;[45, 46]). in a single Vaseline gap voltage clamp system which allowed fiber movement without movement artifacts in Critical steps: voltage sensor currents and charge the membrane current records during contractile activa- movements in skeletal muscle tion [47, 48], (b) from frog fibers studied in a double Vas- The original voltage sensor charge movement currents de- eline gap when stretched to eliminate mechanical tected during and after pulses to a range of membrane po- movement and the corresponding movement artifacts tentials using the pulse protocol in Fig. 2a are shown in during activation [49, 50], and (c) from whole cell voltage Fig. 3a [37]. The voltage sensor current amplitude both clamped mammalian short skeletal muscle fibers adhering during and after the pulses increased with increasing to a glass coverslip [51–54]. depolarization (Fig. 3a). The time course of the charge As embodied in the cartoons in Fig. 1, any hypothet- movement currents during the pulse became increasingly ical positive voltage sensor charges that moved outward rapid as the depolarizing pulses were increased in within the membrane during fiber depolarization (or any Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 5 of 20 ab Fig. 3 Initial characterization of intramembrane charge movement currents. a Non-linear difference currents between test pulses to the indicated membrane potentials and corresponding control pulses covering a membrane potential range negative to the holding potential. Same protocol as in Fig. 1. Pulse duration was decreased for the two largest depolarizations displayed. b Equality of charge moved during the “on” and “off” of different amplitude test pulse depolarizations. c Voltage dependence of non-linear charge moved for test pulses to the indicated membrane potentials. All records and graphs from ref. [37], with modification hypothetical negative voltage sensor charges that moved Non-linear charge movement (Q) had a sigmoidal de- inward) are expected to return to their initial intramem- pendence on test membrane potential, according to a brane location when the fiber was repolarized to its ini- two-state Boltzmann function: tial membrane potential. Thus, the amount of voltage sensor charge moved outward during depolarization was Q ¼ Q max=½ 1 þ expðÞ ðÞ −V þ Vh =k anticipated to equal the amount of voltage sensor charge moved inward during the repolarization. This “on/off” where Qmax is the maximum charge (per unit of linear equality of amounts of charge moving outward during capacitance), Vh is the mid-point, and k a measure of depolarization and inward during repolarization was the steepness (Fig. 3c, continuous line through symbols). expected signature of voltage sensor charge displace- While this procedure allows for an approximation of the ment currents (Fig. 3b). From the first report of mea- voltage dependence of the charge movement in ECC, it surements of the ECC voltage sensor currents in muscle may not be adequate to estimate total charge (i.e., total fibers, it was established that the on/off equality criter- number of elementary charges), especially if the charge ion was in fact fulfilled both for the charge movement moves in multiple sequential steps [55]. currents generated by test pulses to various membrane potentials (Fig. 3b) as well as by pulses of various dura- Voltage sensor charge moved predicts pulse durations tions to the same membrane potential [37]. needed to give detectable contraction An immediate question that arose after the first detec- Voltage dependence of charge movement, and its first tion of charge movement currents was whether the volt- interpretation age sensor currents detected in muscle fibers were in The measured amount of charge moved increased in a fact the control system for depolarization-induced con- sigmoidal manner as a function of increasing membrane tractile activation. Two early studies addressed this ques- depolarization from the resting holding potential and tion, using different pulse protocols to show that voltage approached saturation for the largest depolarizations sensor charge movement measurements can be used to used (Fig. 3c). The voltage dependence of charge moved closely predict the initiation of muscle contraction. First, was interpreted using a model in which a single uniform it was previously well-established that during prolonged population of intramembrane charges were each as- (10s of sec) fiber depolarization, fibers first contracted sumed to occupy one of two possible membrane and then became mechanically relaxed [23]. During locations that differed in energy by a fraction of the full similar prolonged voltage clamp depolarizations, muscle electrical potential energy across the membrane. voltage sensor charge displacement properties were also Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 6 of 20 modified [56]. Comparing the time course of recovery of decreased the time needed to reach contractile threshold charge movement after repolarization of fully depolar- during the test pulse (Fig. 4b). The shortening of the test ized fibers with the time for recovery of just-detectable pulse duration for just-detectable contraction could be contraction during repolarization of a depolarized fiber, predicted from the charge movement recordings as the it was found that charge recovery could predict the re- time to move the prepulse charge at the test pulse covery of contractile ability, implying a close relationship voltage. These studies demonstrated a close correlation between charge movement and contractile activation between the voltage sensor charge movement and [57]. Second, during voltage clamp depolarization of just-detectable contractile activation of muscle fibers. fully polarized fibers, the pulse duration required to pro- The charge required to attain a just-detectable contrac- duce a microscopically just-detectable contraction at dif- tion is here termed “pre-activating” charge since it must ferent depolarizations moved a constant amount of be moved in a step or sequence of steps prior to the voltage sensor charge [47, 48]. In this experiment, step(s) that actually activate contraction, but it does not non-linear capacitive currents (IQ(t)), charge movement itself activate contraction (discussed further below). (Q(t)), and the occurrence of just-detectable contraction were all monitored in the same single muscle fiber Voltage sensors control other membrane potential- (Fig. 4a). Contraction was elicited by test pulses to − 45, dependent processes − 35, and − 25 mV, but not by the smallest test pulse to As predicted by Hodgkin and Huxley [15], any − 55 mV (Fig. 4a). Furthermore, using a test pulse alone Vm-sensitive process was expected to involve a voltage (to − 32 mV) or together with two different amplitude sensor charge movement. Indeed, shortly after the initial prepulses (Fig. 4b, bottom), which alone did not produce measurements of intramembrane charge movement in detectable contraction, it was found that the prepulses skeletal muscle fibers, analogous charge displacement Fig. 4 Depolarizing pulses that produce a just-detectable muscle fiber movement displace a set (“threshold”) amount of charge, which can be termed “pre-activating” charge since it must move in order to attain detectable fiber activation. a Charge movement records during muscle fiber depolarization to indicated voltages. The dashed vertical lines indicate the pulse duration needed to give a microscopically just-detectable fiber movement for shorter pulses to the same voltage. No contraction was detected in at − 55 mV. b Pulse to − 32 mV applied alone or together with indicated prepulses, which move only pre-activating charge, since no contraction was detected during the prepulses alone. Charges moved for just-detectable fiber movement (height of black dots) in the test pulses were the same with or without prepulses at the pulse durations for just-detectable fiber contraction (dashed vertical lines). The prepulses decrease the pulse duration required to reach detectable fiber contraction during the test pulse, and this decrease was equal to the time to move the prepulse (pre-activating) charge at the test pulse voltage. Reproduced, with modification from ref. [47] Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 7 of 20 currents were monitored in squid axons [58]. Based on release from the SR (red arrow in Fig. 5; discussed in their properties, the intramembrane charge movements detail below). It is crucial to note that even though iso- 2+ detected in axons were identified as “gating currents” for lated RyR1/Ca release channels of skeletal muscle can + 2+ the axon Na channels. Following the initial work of be activated by elevated Ca [71, 72], it is well estab- 2+ 2+ Schneider and Chandler in muscle and of Armstrong lished that Ca entry via the Cav1.1 Ca channel 2+ and Bezanilla in axons, charge movement of voltage sen- current is not required for activation of RyR1 Ca re- sors has been used extensively to study channel gating lease during muscle fiber depolarization [73], where kinetics and putative voltage-dependent molecular rear- depolarization beyond the reversal potential for L-type 2+ 2+ rangements in a variety of voltage-sensitive channels Ca current [74]orin zeroCa external with EGTA 2+ [59–62] and even in membrane potential-dependent en- [75], which eliminates inward Ca current, does not zymes, pumps, and receptors [63–65]. Over the years, alter muscle activation. Indeed, “skeletal” type of ECC 2+ various pulse protocols (“P/n”,+/− P) have been devised is defined as being Ca influx-independent ([76]; see to extract the non-linear capacitative current (the further discussion below). “charge movement current” or “gating current”) from the total capacitative current [44, 66, 67], in addition to Monitoring and characterizing TT membrane 2+ the P test–P control protocol (Fig. 2a, b) developed for depolarization-induced SR Ca release the initial measurements of muscle voltage sensor An important experimental distinction exists between 2+ charge movement [37]. the two Ca channels regulated by the TT Cav1.1 volt- 2+ age sensor. L-type Ca current can be monitored dir- 2+ A multi-tasking Ca channel: the ECC voltage sensor ectly using the same voltage clamp circuit as used for 2+ controls two distinct Ca channels in two different monitoring voltage sensor charge movement [69, 70]. In 2+ membranes contrast, SR Ca release occurs across the SR mem- Activation of the TT voltage sensor within the Cav1.1 brane, which is not part of the electrical circuit for 2+ molecule controls two different TT voltage-sensitive Ca current flow between the cytoplasm and bathing solu- channels [68]. First, Cav1.1 voltage sensor movement tion that is monitored by the voltage clamp circuit. Con- 2+ 2+ leads to opening of the ion conducting Ca channel sequently, SR Ca release cannot be monitored by the within the Cav1.1/ECC voltage sensor molecule itself voltage clamp system. A second experimental measuring 2+ [69, 70]. This allows Ca influx across the TT mem- system and analysis procedure is needed to calculate SR 2+ brane and into the cytoplasm (blue curved arrow in Ca release. 2+ 2+ Fig. 5), which is manifested as L-type inward Ca The first step in determining SR Ca release is to 2+ current across the voltage clamped TT system. Second, monitor the free myoplasmic Ca concentration during the Cav1.1 voltage sensor movement promotes opening a voltage clamp depolarization [77, 78] (Fig. 6a, b), or 2+ 2+ of the SR RyR1/Ca release channel [8, 9], allowing Ca during an action potential or train of action potentials 2+ 2+ Fig. 5 Cav1.1 (pale blue) serves as voltage sensor for two different Ca channels: its own intramolecular Ca channel in the TT membrane 2+ (current illustrated in blue) and the RyR1 Ca release channel (tan) in the SR membrane (current illustrated in red). Cartoon representation of 2+ 2+ the simplest gating mechanism. RyR1 Ca channel is directly controlled by molecular coupling of Cav1.1 to RyR1. Note that Ca influx via 2+ 2+ the TT Cav1.1 Ca channel is not needed for activation of the RyR1 SR Ca release channel Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 8 of 20 ab 2+ 2+ Fig. 6 Rate of Ca release from the SR during muscle fiber depolarization calculated from the myoplasmic Ca transients measured experimentally 2+ 2+ in individual muscle fibers. a Measured Ca transients (top) and corresponding calculated time course of rate of Ca release from the SR (middle) for 2+ voltage clamp depolarizations to indicated membrane potentials (bottom). The rate of Ca release reaches an early peak and then 2+ declines appreciably during continued depolarization. Reproduced, with modification from ref. [78]. b Time course of recovery of Ca release following an initial inactivating pulse, followed at various times by a repeat application of the same pulse. After a lag of about 100 ms, the early peak begins to recover and is fully recovered by 600 ms. However, at 600 ms recovery, the release wave form is still smaller than in the initial pulse and recovers much more slowly, indicating recovery from a second process, which was attributed to 2+ 2+ 2+ recovery from SR Ca depletion. Reproduced, with modification from ref. [83]. c Ca transients (top) and rate of SR Ca release 2+ (bottom) calculated from the measured Ca time courses for a single action potential or for a train of action potentials. Release in the second and later action potentials is considerably reduced compared to the release in the first action potential. Reproduced, with modification from ref. [79] (Fig. 6c)[79] using a calcium-sensitive indicator dye and amplitude and duration, with a variable time interval be- appropriate optical apparatus [77, 80–82]. However, the tween the two pulses (Fig. 6b). For test pulses applied 2+ measured myoplasmic free Ca transient represents shortly after the conditioning pulse, the time course of 2+ 2+ only a small fraction of the total Ca released during the Ca release completely lacked the early peak 2+ the fiber depolarization. A much larger fraction of the (Fig. 6b). Importantly, the inactivation of Ca release 2+ 2+ released Ca is bound to endogenous myoplasmic Ca during a 20–50-ms pulse does not appear to be due to 2+ binding sites (troponin C, parvalbumin, SR Ca pump) modification of the voltage sensor since charge move- 2+ or transported back to the SR. Taking the Ca binding ment is not modified during these pulses, as judged by properties of these binding sites and transport into con- the criteria Qon = Qoff, and Q kinetics are not modified 2+ 2+ sideration, the Ca release flux (rate of Ca release) after an inactivating prepulse [84]. can be calculated [78]. An important first result of such 2+ calculations was the conclusion that Ca release is not Pre-activating and activating components of charge 2+ maintained during a step depolarization or during a movement for SR Ca release train of action potentials, but instead declines during a In theory, “pre-activating” (also termed “sub-threshold” 20–50-ms step depolarization (Fig. 6a, middle records) or “threshold”) charge movement would be generated by [78] or during a 100-Hz train of actin potentials (Fig. 6c, charge-generating molecular transitions that precede the 2+ lower records) [79]. A slower phase of decline of release actual SR Ca channel opening event in the signaling during longer duration voltage clamp depolarizations pathway from charge movement to RyR1 activation. In 2+ also was observed and was attributed to Ca depletion contrast, the “activating” charge would coincide with 2+ from the SR [83, 84], but the faster developing decline of and determine the actual opening of the SR Ca release 2+ Ca release during a voltage clamp pulse or train of channel. As described above, the pulse duration needed 2+ APs appears to reflect inactivation of SR Ca release. to produce a microscopically just-detectable fiber move- Another important feature of the inactivation is its re- ment for various voltages was found to be the pulse dur- covery, as demonstrated using a double-pulse protocol ation that produced the same constant (= “threshold”) [83]. Here a first conditioning pulse of fixed amplitude amount of charge moved at each voltage, including dur- and duration is followed by a second pulse of the same ing the stepped-on pulse pattern (Fig. 7, inset voltage Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 9 of 20 protocol) [50]. This observation implicates the threshold preceded by the same prepulse, which moved a substan- (or pre-activating) charge as being a precursor that tial amount of the pre-activating charge, but did not ac- 2+ moves before the charge component that actually causes tivate detectible Ca release (Fig. 7, right inset). Using 2+ Ca release and the subsequent contractile activation. these data, it was possible to systematically separate the 2+ 2+ Using Ca -sensitive dyes and the Ca release calcula- charge movement that was predominantly precursor 2+ tion described in the preceding section, together with (“pre-activating”) for producing SR Ca release (and the stepped-on pulse protocol (e.g., Fig. 7, lower left was moved in the prepulse) from charge that was pre- 2+ inset), it became possible to relate the activating compo- dominantly activating for Ca release (and was moved 2+ 2+ nent of charge movement to the rate of SR Ca release, during the test pulse) (Fig. 7, inset right). Ca transients the process directly downstream of voltage activated were simultaneously measured for the same pulses, and 2+ charge movement, and thus directly controlled by the the Ca release time course was calculated for each test voltage sensor [50]. pulse (Fig. 7, inset top left). Over a wide range of test pulse amplitudes and durations (Fig. 7, voltage protocol), 2+ 2+ Peak SR Ca release due to a pulse is proportional to the the peak rate of Ca release during the test pulse was amount of activating charge moved by the pulse found to increase linearly with the charge that was 2+ To relate Ca release to charge movement, charge moved by the test pulse (Fig. 7). The linear relationship movement was determined for a range of test pulse am- had a small positive charge value for the extrapolation to plitudes and durations, with each test pulse immediately zero peak rate of release (x intercept), indicating a small amount of pre-activating charge that was not moved during the subthreshold prepulse, but was instead moved during each test pulse, presumably the initial charge moved during the test pulse. These results dem- onstrated a close relationship between the extent of acti- 2+ vation of SR Ca release by a pulse and the amount of activating charge that moved during the same pulse [50]. Minimal model for voltage sensor control of a coupled 2+ RyR1 Ca release channel Figure 5 presents a cartoon of the functional states of 2+ the voltage sensor/L-type Ca channel in the TT mem- 2+ brane (top) and the RyR1 Ca release channel in the SR membrane (bottom) for a hypothetical minimal (two-s- tate) model [85] for regulation of RyR1 by its directly coupled TT voltage sensor(s). In this highly simplified gating scheme, fiber depolarization (top, left to right) 2+ Fig. 7 The peak rate of Ca release evoked by various depolarizing causes the mobile voltage sensor charges within the TT test pulses is linearly related to the amount of “activation” membrane voltage sensor protein (Cav1.1), which are intramembrane charge moved by the same pulse. Each test constrained to remain within the TT membrane, to re- pulse was immediately preceded by a depolarizing prepulse (see spond by generating intra-membrane movement (de- pulse schematic in lower left inset), which by itself did not 2+ tected by charge movement measurements [86]). In the activate Ca release. Many of the test pulses were too short to establish the ionic current baseline for calculating charge moved minimal scheme of Fig. 5, the voltage sensor charge during the test pulse, so Qon could not be measured for short movement obligatorily induces the opening of the RyR1 test pulse durations due to uncertainty regarding the level of 2+ Ca channel directly coupled to the voltage sensor(s) ionic current remaining during the test pulse. Charge moved by 2+ (bottom; left to right), resulting in a release of Ca from the test pulse was consequently determined as Qoff–Q pre. The the SR, as detected by monitoring the total increase in test pulse charge represents an upper estimate of the activating 2+ charge moved during the test pulse; the x intercept on the cytoplasmic and transported Ca . In this minimal graph is interpreted as representing the amount of pre-activating model, there are only two states of the voltage sensor (i.e., precursor) charge still present in the test pulse (i.e., pre-activating and its directly coupled RyR1 unit: voltage sensor rest- charge not moved by the prepulse). The lower right inset gives the ing/RyR1 closed (left) or voltage sensor active/RyR1 charge movement current records obtained using various duration test open (right). In addition, in this minimal scheme, the pulses to 0 mV, each immediately following the prepulse to − 50 mV. 2+ 2+ 2+ The upper left inset gives the Ca release calculated from the Ca L-type Ca channel in the TT membrane is also open transient recorded simultaneously with the charge movement records when the voltage sensor is active [8]. shown in the lower right inset for each of the same test pulses. The minimal model (Fig. 5) already raises several basic Reproduced, with modification from ref. [50] issues regarding the TT voltage-dependent gating of Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 10 of 20 RyR1. First, intramembrane charges (transmembrane charge movement, it is necessary to consider inactivation 2+ positively charged S4 alpha helices in the voltage sensor, of the SR Ca release channel, which occurs during the Cav1.1) move outward during TT depolarization fiber depolarization (Fig. 6)[83, 87]. The inactivation 2+ (i.e., away from the RyR). However, the resulting mo- process obscures the time course of Ca release channel 2+ lecular rearrangements in the voltage sensor accompany- activation, reducing Ca release to a fraction of the ing or following charge movement could bring other peak value reached earlier during the pulse (Fig. 6a, b) domains of the Cav1.1 voltage sensor (alpha 1) sub- [87]. One approach eliminated the effects of inactivation unit, or beta subunit either toward or away from the during a pulse by pre-inactivating the fibers (Figs. 6b RyR1, so at present we do not know whether the and 8a). Here a large “inactivating” prepulse, sufficient 2+ voltage-sensitive step constitutes the removal of an to produce maximal inactivation of RyR1 Ca release, inhibition to RyR1 opening, or the application of a was immediately followed by a brief repolarization (to positive factor for voltage-dependent RyR1 activation. return all charge to the resting state and to turn off the 2+ This basic issue awaits high-resolution molecular non-inactivating Ca release) and then by a test structure-function studies of the Cav1.1/RyR1 inter- depolarization during which activation of both the 2+ action in various functional states. Second, the gating “non-inactivating” component of Ca release and the scheme in Fig. 5 includes a major simplification. If charge movement (which does not inactivate in a each of the two ellipses shown in Fig. 5 represents few hundred ms time window: see above) were mon- one of the 4 Cav1.1 α1 subunits coupled to a single itored (Fig. 8a, b)[83, 88]. Using this approach it RyR1 molecule, then each Cav1.1 (i.e., each ellipse) was found that both the voltage dependence (Fig. 8c) 2+ should have four S4 segment charged transmembrane and time course (Fig. 8b) of the non-inactivating Ca α helices, one in each of the four transmembrane do- release very closely agreed with the voltage depend- mains of each Cav1.1, rather than the single charged ence and time course of (Q/Qmax) [88], where helix shown for each Cav1.1 in the simplified cartoon Qmax is the maximum charge, moved during a large in Fig. 5. depolarization This remarkable finding is consistent A clear shortcoming of the minimal scheme in Fig. 5 with a reaction scheme in which the voltage depend- is that all of the voltage sensor charge movement is dir- ence and kinetics of the non-inactivating component 2+ ectly involved in the closed to open transition of the of SR Ca releaserateiscontrolledbyfouridentical RyR1 to which it is coupled (i.e., all of the charge moved and independent voltage sensors [88]. Each RyR1 is “activating” charge movement, as defined above). This channel would be controlled by four identical voltage property clearly does not agree with the experimental sensors and is open when and only when all four characterization of the control system in muscle fibers. voltage sensors are in the active conformation, giving An appreciable fraction of the total charge movement rise to release being proportional to the fourth that is recorded from a fiber is pre-activating charge, power of Q/Qmax. which moves during depolarization, but prior to the If one “voltage sensor” is one Cav1.1 molecule, the charge for the actual activation step in the control voltage sensors could be independent (i.e., in different 2+ mechanism for Ca release (above) [50], yet is an es- molecules), and the RyR1 could require all four sential prerequisite for depolarization-activated SR Cav1.1s to be active in order for RyR1 opening. In- 2+ Ca release. In the following sections, we will exam- deed, early freeze fracture studies using electron mi- ine two functional studies that consider more compli- croscopy (EM) revealed that Cav1.1 forms four cated models for voltage sensor/RyR1 interaction(s), ordered clusters (tetrads) and the space between tet- 2+ as well as other changes occurring in the individual rads suggested an overall 1:2 ratio of tetrads to Ca components, leading to the need for increased com- release channels [89], corresponding to the observa- plexity and refinement of the minimal gating scheme tion that only half of the RyR1 homotetramers in the in Fig. 5. triad junction were coupled to Cav1.1 tetrads. How- ever, each Cav1.1 molecule contains four positively 2+ Model for RyR1 Ca release activation requiring charged S4 transmembrane helices, as described simultaneous activation of four identical but independent below. One possibility is that in the Cav1.1-RyR1 voltage sensors complex in a functioning muscle fiber, interaction be- In a preceding section and Fig. 7, we considered the em- tween the Cav1.1 and RyR1 gives rise to the condition 2+ pirical (linear) relationship between the peak rate of Ca that only one of the four S4 helices in each Cav1.1 is release produced by a given pulse and the activating physically able to move, resulting in a single mobile charge moved in the same pulse [50]. In order to next charged group per Cav1.1, with four independent 2+ relate the time course of Ca release activation during voltage sensors (one in each Cav1.1 of the tetrad) per 2+ TT depolarization to the time course of voltage sensor coupled RyR1 homo tetramer Ca release channel. Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 11 of 20 ab Fig. 8 Empirical evidence for a model for activation of each RyR1 in which four identical and independent voltage sensors must all be in the 2+ 2+ active configuration for the SR Ca release channel to open after pre-inactivation of the RyR1 channel. a Ca transients (top) and calculated rate 2+ of SR Ca release (middle) for the pulse protocol at the bottom. Pulses to various test pulse amplitudes were each preceded by the same inactivating prepulse, followed by brief return to the holding potential to reset the voltage sensors and close the RyR1 channel and then the 2+ 4 2+ various depolarizations (bottom). b Wave form of Ca release during the test pulses closely follows Q . c Voltage dependence of Ca released follows the fourth power of charge moved (dashed curve). From ref. [88], with modification Allosteric mechanisms for RyR channel control by TT justify the added transitions of the allosteric model, add- Cav1.1 voltage sensor itional experimental data are required and were intro- 2+ In the preceding model, the Ca release channel opens duced [92]. Also note that each “voltage sensor” when and only when a certain enabling configuration of considered here, as in the four independent voltage sen- the voltage sensors was achieved. An allosteric model sor models above, is an entire Cav1.1, containing a [90, 91] for gating of each RyR1 homo-tetramer by four charged transmembrane “S4” helix in each of its four independent DHPRs provides an alternative approach transmembrane domains. (Fig. 9a, b). In the allosteric system, each RyR1 channel can open when any number of its coupled voltage sen- Molecular components and mechanisms in ECC sors is active [92]. However, RyR1 opening becomes in- Cav1.1: the TT voltage sensor creasingly likely as more voltage sensors are active Membrane depolarization of the TT system, during an (Fig. 9b). In this model, each RyR1 channel has two AP or voltage clamp step depolarization, is detected by states, closed (C) and open (O). Lateral transitions rep- Cav1.1 channels, the TT voltage sensor. Cav1.1 channels resent opening or closing transitions of the RyR1. Each were initially identified using electrophysiological ap- voltage sensor has two states, inactive or active (− or +, proaches (charge movement and ionic currents) [37, 56, respectively in cartoon). Vertical transitions represent 69, 70, 93]. Cav1.1 channels are principally expressed in changes in the activation (+) or deactivation (−) status of the membrane of the TT system of adult skeletal muscle each of the four voltage sensors, as indicated by the four fibers and are members of a diverse family of 2+ circles with + or − representing the four voltage sensors voltage-dependent Ca channels. Molecular details of 2+ controlling each RyR1 [92]. It should be noted that re- the voltage-gated Ca channels from skeletal muscle moving the open states O through O , which are open were first identified by binding, purification, and recon- 0 3 and have less than four active voltage sensors, and the stitution [94]. Using molecular biology techniques, their transitions to and from each of these states removes the amino acid sequences were determined by cDNA clon- possibility of opening without movement of all four volt- ing and sequencing [43]. Contrasting with their promin- age sensors, and thereby reduces the allosteric model to ent functional status, TT voltage sensor (Cav1.1 the four independent voltage sensor model. Thus, the channels) was somewhat apart from the saga of allosteric model includes the four voltage sensor models, structure-function studies for other types of ion chan- 2+ which already fit the data for non-inactivating release nels [95]. The skeletal muscle Ca channel complex is a very closely [88], as a subset of possibilities. In order to hetero-tetramer, comprised of a main pore-forming α1 Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 12 of 20 Fig. 9 Allosteric model for gating of each coupled RyR1 by four different voltage sensors. a Cartoon of a single RyR1 homo-tetramer controlled by four independent voltage sensors, with all four voltage sensors in the inactive configuration (left) or with one voltage sensor in the activated 2+ configuration (right). b Kinetic reaction scheme for the RyR1 Ca channel gating process. Channel opening becomes more likely as more voltage sensors become active, which is the basis for the allosteric effect. However, the channel can open with any number of voltage sensors active. The subscript i on C or O is the number of voltage sensors active. Forward rate constant k for Ci→ Oi increases with i; the backward rate constant k for Oi→ Ci decreases with i. Note that f < 1. Elimination of states O to O from this scheme reduces it to the model proposed in Fig. 8. From ref. 0 3 [92], with modification 2+ subunit, complexed with β, α2δ, and γ subunits [38, 39, of Ca channels and their structural examination [95], 96] (Fig. 10a) and other ligands (i.e., STAC3) [97]. The which is changing with the molecular interpretation of Cav1.1 α1 subunit consists of a single polypeptide chain, their function. Recent cryo-EM studies at a resolution of with four highly homologs but non-identical intramem- 3.6 Å [100] revealed more details about the molecular brane domains (I–IV), each containing six transmem- architecture of the Cav1.1 channel of skeletal muscle with brane (TM) alpha helical segments (S1–S6), shown in its complete set of auxiliary subunits (Fig. 10a). The cen- cartoon representation in Fig. 10b [43, 96], as well as tral α1-subunit of CaV1.1 has a core structure and is asso- amino and carboxyl terminals. While the organization of ciated with an extracellular α2δ-subunit, an intracellular the TM domains of the Cav1.1 α1 subunit has a strong β-subunit, and a 4-TM γ-subunit (Fig. 10a). pseudo fourfold symmetry in the plane of the TT mem- brane, the intracellular structure Cav1.1, including the 2+ RyR1, the SR Ca release channel in skeletal muscle single β subunit is highly asymmetrical, which could 2+ 2+ RyR1-dependent SR Ca release via the Ca release have important implications for Cav1.1-RyR1 coupling. channel RyR1 initiates muscle contraction. The RyR1 is Segments S1–S4 of each transmembrane domain of the a colossal protein of approximately 2.3 MDa assembly of α1 subunit form a voltage-sensing domain (VSD) [44, four identical subunits [101, 102]. Each subunit contains 98], whereas segments S5 and S6 from all four intra- 2+ an intramembrane region, located within the C-terminal membrane domains contribute to the Ca -conductive region and representing ca. 20% of the total protein, plus pore (Fig. 10b and ribbon diagram in Fig. 11a [98]). The a cytoplasmic region that represents 80% of the total fourth TM helical segment (S4) contains a number of protein, and is known as the foot region (Fig. 11b, blue positively charged amino acids (Arg and Lys), separated structure; note that the spatial scale is about five times by two hydrophobic residues (Fig. 10b)[43, 98]. During compressed to Fig. 11a, so Cav1.1 appears much smaller changes in TT membrane potential, the S4 segments are in Fig. 11b than a); [103–107]. The cytoplasmic region believed to rearrange, moving outward across the plane of the RyR1 channel (280 Å × 280 Å × 120 Å) is continu- of the TT membrane, in response to membrane ous with the transmembrane region (120 Å × 120 Å × depolarization, establishing the determinants for voltage 60 Å; Fig. 11b)[41]. The RyR1 SR transmembrane re- sensitivity [9, 99]. Historically, there was a considerable 2+ gion forms the Ca release channel [106–109]. time gap between recognition of the biological importance Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 13 of 20 ab Fig. 10 Architecture and membrane topology of the Cav1.1. a Side view of a cryo-EM reconstruction of the Cav1.1 and its subunits at 3.6 Å (Protein Data Bank (PDB) 5GJW) [100]. The Cav1.1 α1 subunit is highlighted in blue. Auxiliary β-, α2δ-, and γ-subunits are colored in green, red, and yellow, respectively. This model was created in Chimera [174]. The asterisk symbol indicates the missing II–III loop sequence in the cryo-EM structure. b Membrane topology of the Cav1.1 α1 subunits. The α1 subunit is composed by an interconnect array of four homologous (but not identical) domains, each domain consisting of six transmembrane domains, S1–S6. S1–S4 from each domain form a voltage sensor domain, whereas S5 and S6 from all four domains form the pore domain. Intracellular loops connect the domains; the loops II–III and I–II are important for ECC, as indicated ab Fig. 11 Structural details of the Cav1.1 α1 subunit and the RyR1. a Shown are cryo-EM reconstructions of the Cav1.1 a subunit, side view (top) and upper view (bottom) (PDB 5GJW) [100]. The auxiliary subunits, the cytoplasmic tails, and loops are not shown. Each domain is color coded, positively charged residues are indicated in red, negatively charged residues are shown in blue. The dashed ellipse indicates the location of S1–S4 from domain I and the black circle shows the corresponding pore domain (S5–S6) from domain I. Note that each voltage sensor domain (S1–S4) is not in close proximity to its corresponding pore domain (S5–S6). b Side (top) and upper (bottom) views of cryo-EM reconstruction of the RyR1 (PDB, 5TAL) [175] and four superimposed Cav1.1 α1 subunits (PDB 5GJW) [100], forming a tetrad. This model was created in Chimera [174] using the cryo-EM maps with relative location of the Cav1.1 subunits as suggested by Samsó [118] Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 14 of 20 2+ RyR1s are arranged in a regular array within the ter- reversal potential, where Ca influx is greatly sup- minal cisternae of the junctional SR [89, 110, 111]. Simi- pressed or eliminated [74]. Second, the frequency of oc- 2+ 2+ larly, Cav1.1 channels are clustered in groups of four (or currence of Ca sparks, elementary intracellular Ca tetrads) in the TT membrane that is adjacent to the signals [129, 130], in frog skeletal muscle fibers can be 2+ junctional SR [89, 112], with a Cav1.1 tetrad facing every used as a measure of activation of “microscopic” Ca other RyR1 in the junctional SR RyR1 array (Fig. 11b). release events, which combine at high frequencies dur- For symmetry and simplicity, each Cav1.1 molecule ing fiber depolarization to produce the “macroscopic” 2+ composing a tetrad is believed to be oriented in the Ca transient observed during depolarization [131, same coordinated position relative to the subunits of its 132]. These events are extremely infrequent in resting fi- apposed RyR1homo tetramer (see Fig. 11b)[89, 112]. bers, but increase tremendously in frequency during Since these interfaces take place at alternate RyR1s, half depolarization, so much so that spark frequency can of the RyR1s are “uncoupled” with Cav1.1s, and half are only be monitored experimentally during large depolar- coupled to Cav1.1s [113]. The location of RyR1 (coupled izations by using depolarized fibers, and restoring only a and uncoupled) determines the organization of the small fraction of the release units by brief repriming re- Cav1.1 channels in the juxtaposed TT membrane, creat- polarizations [130, 133–135]. These types of experiments ing a “checkerboard” array of coupled and uncoupled indicate that in functioning muscle fibers, the RyR1 SR 2+ RyR1s that produces the Cav1.1 lattice organization Ca release channels are essentially fully off when the [114]. Depolarization-induced activation of RyR1 is be- voltage sensors are in the resting condition, but turn on lieved to be mediated via direct or indirect interactions strongly and rapidly during the AP or voltage clamp with TT voltage sensors [8, 115–117]. However, despite depolarization that activates the voltage sensors [132, extensive biophysical and ultrastructural studies, the 135, 136]. The simplest hypothesis is that under normal molecular basis for TT voltage sensor function and the conditions in mature functioning muscle fibers, RyR1s chemical mechanisms that support TT voltage regulated coupled to TT voltage sensors are locked in the off con- 2+ RyR1 SR Ca release, as well as the orientation of the figuration due to an inhibitory influence of the voltage Cav1.1 tetrads relative to the RyR1 have remained un- sensor in its resting configuration [137]. Movement of clear. In Fig. 11b, the location of the tetrads is based on the voltage sensor into the active configuration during reference [118], whereas the relative orientation is depolarization removes this “lock” on RyR1 opening, and arbitrary. the RyR1 channels open and, more slowly, inactivate The RyR1 components, including SR luminal seg- (Fig. 6a)[78]. When the voltage sensor returns to the ments, transmembrane domains, and large cytosolic do- resting configuration at the end of the depolarization, mains, and their interaction with the Cav1.1 channels, the RyR1 is relocked. In this scenario, all the other li- SR luminal proteins, and accessory proteins, metabolites, gands that modulate RyR1 channel activity in isolated and ions, as well as post-translational modifications, membrane or protein preps may or may not also simi- allow the RyR1s to be fine-tuned by numerous mecha- larly modulate the RyR1 in a functioning fiber, but the nisms [71, 107, 109, 119–128]. However, the TT voltage TT voltage sensor serves as master regulator determin- sensor (CaV1.1) is believed to be the “ligand” that ing whether or not the channel can open at all. uniquely enables RyR1 opening in functioning skeletal muscle fibers. Dysgenic muscle and restoration of skeletal vs. cardiac The TT voltage sensor is the master regulator of RyR1 ECC Despite the large number of modulatory interactions Advances in biochemistry, molecular biology, and that influence RyR1 activation, it is difficult to pharmacology allowed the identification of the molecular overemphasize the importance of the TT voltage sensor, components that are essential for ECC. One crucial dis- and/or some component(s) directly coupled to it, in the covery was the characterization of a naturally occurring physiological regulation of RyR1 in skeletal muscle. In- “knock out” of the Cav1.1 α1 subunit (“dysgenic” mouse; deed, the voltage sensor can be considered as the master [138]). This model demonstrated that myotubes derived 2+ ligand for controlling SR Ca release in normal mature from the dysgenic mice lacked ECC and intramembrane muscle fibers. First, no other component, except possibly charge movement; the expression of α1s subunit (skel- 2+ Ca influx into the TT/SR gap, would be anticipated to etal muscle isoform) in these cells restored “skeletal” 2+ change drastically during the few millisecond action po- type of ECC, which is independent of Ca influx [139]. 2+ tential which raises cytoplasmic Ca sufficiently to The expression of the cardiac isoform (α1c subunit) of 2+ cause a twitch contraction. However, Ca influx is not the Cav1 channels did not restore skeletal ECC [140]. needed for skeletal muscle activation [73], which is The inability of the cardiac isoform [140] and of other 2+ 2+ maintained for depolarizations well beyond the Ca Ca channel subtypes [141, 142] to rescue the ECC Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 15 of 20 allowed the investigation of essential elements for skel- essential for the skeletal muscle function. Skeletal ECC etal muscle ECC via chimeric channels. was not experimentally evident if Cav1.1/RyR1 were not in the membrane, or forming tetrads. Components of Cav1.1 that are needed for activating RyR1 Discrete S4 voltage sensor (i.e., S4, I–S4, IV) models Chimeric channels made with α1 subunits of Cav1.1 and In voltage-gated sodium channels, a channel structurally Cav1.2 demonstrated that a region in the intracellular and evolutionary similar to Cav1.1 [39], studies using mu- loop between the second and third domains (II–III tagenesis and voltage-clamp fluorometry revealed that the loop), specifically, the region spanning residues 720– four VSDs, each linked to a partial pore-forming region, 764/5, was important for this function (Fig. 10b)[143, may be differentially and allosterically coupled to the pore 144]. Interestingly, in the cryo-EM structure of Cav1.1 opening to various degrees of involvement in the control the structure of the II–III loop is undefined (dashed line of voltage dependence and gating [163]. VSDs I–III acti- in Fig. 10a,[100]), whereas the I–II and III–IV/C-ter- vate in parallel and sufficiently rapidly to modulate Na minal regions are defined and appear in Fig. 10a. While channel opening, whereas VSD IV activates more slowly the identification of the Cav1.1 regions that are critical and initiates fast inactivation [163]. In Cav1.2, a 2+ for ECC has been more active, perhaps due to the voltage-gated Ca channel expressed in cardiac cells, smaller size of the Cav1.1 channel, the identification of site-directed fluorophore labeling, and voltage clamp binding domains in the RyR1 for the II–III loop has fluorometry of individual VSDs showed differential func- been less fruitful. Only a few reports, where deletions of tion for each domain; VSDs II and III exhibited large segments of the RyR1 successfully altered ECC, voltage-dependent and kinetic characteristics compatible concluded that several regions of the RyR1 are involved with channel activation [164]. However, the cardiac iso- in the interaction with Cav1.1 [145–147]. Recent ap- form (α1c subunit) is unable to support RyR1 activation in proaches are revisiting the role of the loop II–III and its myotubes lacking α1s [140, 144]. 2+ association with the adaptor protein STAC3 on ECC Similarly, the voltage dependence and timing of Ca [148]. These new results support the notion that the II– entry via Cav1.1, as well as the voltage dependence and 2+ III linker plays a role in ECC. timing of TT voltage-dependent RyR1 Ca release, are ex- The Cav1.1 I–II loop is the site for interaction with the pected to be functions of the α1-subunit of Cav1.1, which β1a subunit (Fig. 10b)[98]. The β1a subunit is important also contains four highly similar but non-identical VSDs, for several aspects of ECC. The β1a subunit is needed for I–IV [43, 96, 98]. Evidence for a differential role of each the functional expression of Cav1.1 α1 subunit [149]and VSD in Cav1.1 channel operation using chimeric studies is crucial for enhancement of Cav1.1 α1 triad expression (interchange of VD SI region, Cav1.1↔Cav1.2) and alter- [150], assembly of Cav1.1 α1in tetrads [151, 152], and native splicing of VSD IV of Cav1.1 suggests that VSDs I elicitation of Cav1.1 α1 charge movement [153]. and IV control the activation kinetics and voltage depend- Skeletal-type ECC is reduced in muscle cells lacking the ence, respectively [165–167]. Because VSD I and VSD IV expression of β1a [150] and is rescued by expression of appear to be linked to the slow activating Cav1.1 ionic β1a [154]. The use of chimeric constructs of β1a [155] current, it was hypothesized that VSDs I and IV do not 2+ with other β subunits, as well as the use of synthetic pep- contribute to the more rapid Cav1.1-dependent SR Ca tides [156], allowed the identification of the C-terminal re- release [168, 169] (see Fig. 12). In support of this hypoth- gion of β1a as an important domain for possible esis, a functional study of a mutation causing malignant interaction with RyR1 during TT voltage-dependent SR hyperthermia susceptibility (R174W) in S4 of VSD I of 2+ Ca release. Cav1.1 revealed that this mutation reduces Cav1.1 ionic 2+ Thus, several sites in the Cav1.1 α1 subunit, in current, but does not affect Cav1.1-dependent SR Ca re- addition to the II–III loop, contribute to the overall lease [170](Fig. 12). Wu and colleagues [171]used a Cav1.1/RyR interaction. These include the loop I–II and mouse model for hypokalemic periodic paralysis with a β-subunit [157, 158], and more indirectly, the III–IV targeted Cav1.1 R528H mutation in S4 of VSD II. Muscle loop [159] and the C-terminal domain of the Cav1.1 α1 fibers from the Cav1.1 R528H homozygous mouse exhib- 2+ [141] (Fig. 10)b. ited impaired depolarization-induced Ca release, sug- Another important advance in the characterization of gesting that VSD II could participate in Cav1.1-dependent 2+ the molecular players of the ECC and their interactions SR Ca release [171](Fig. 12). These results represent was the generation of a RyR1-knockout mouse (the dys- compelling but still indirect evidence of the role of each pedic mouse [160]), which allowed for the expression of VSD in Cav1.1. Currently, the contribution of the individ- various RyR constructs and different Cav1.1 α1/RyR1 ual VSDs to the voltage dependence of Cav1.1 pore open- 2+ combinations [161, 162]. These approaches identified ing and activation of RyR1 Ca release is unknown. Note that the skeletal Cav1.1 α1-subunit and RyR1 are that the Cav1.1 R528H mutation also introduces a “gating Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 16 of 20 Fig. 12 Voltage sensing domains of Cav1.1. From left to right, S4-I, S4-II, S4-III, and S4-IV. The positively charged residues of the S4 segments are labeled in red for each domain. Negatively charged amino acids are shown in blue. The depicted transmembrane helices are from the structure of the cryo-EM of Cav1.1 (PDB 5GJW) [100]. These models were created using Chimera [174]. Refs. for VDSI [165, 170], VDSII [165, 171], VSD III [165, 167], and VSD IV [165–167] pore” or “omega” current, which is normally not present near-atomic level, have provided in depth functional and in the wild-type channel and is responsible for the anom- structural details of the ECC process. However, it is clear alous depolarization seen in hypokalemic periodic paraly- that new approaches are needed to continue to explore sis [171]. Details regarding the gating pore current have the intricacies of ECC. Some of the many remaining un- been reviewed in [172, 173]. answered questions regarding ECC include the follow- ing: Are all four VSDs (I–IV) needed to activate the 2+ Future perspectives RyR1 Ca channel, or is only a subset of charges in- How does the propagated electrical impulse spreading volved? If so, which VSDs are coupled to Cav1.1 pore 2+ along the TT system produce Ca release? The Schneider opening? Which VSDs contribute the voltage sensor ele- and Chandler hypothesis that the excitatory signal passes ment(s) for electromechanical coupling between the 2+ from the TT to the SR membrane by way of charges mov- Cav1.1 and RyR1 Ca release? Which residues are moved ing in the TT membrane connecting with the junctional during Cav1.1 channel activation? Which residues are 2+ 2+ feet of the SR Ca release channel initiated the path to moved for RyR1 Ca release channel activation? How far answer this question. Yet, it is still unknown how this do they move within the membrane electric field? And movement of charge, originating in Cav1.1, transfers a sig- what are the molecular determinants that mediate the 2+ nal across to the RyR1 (feet) to trigger Ca release. This electromechanical coupling between the VSD and RyR1 2+ question is particularly fascinating because of the as yet Ca release? These are some of the interesting questions unknown molecular structure-function relationship be- for current and future investigation. tween these components in two different membrane sys- tems, the TT (Cav1.1 voltage sensors) and the SR (RyR1 2+ Abbreviations Ca release channels). Cryo-EM has revealed amazing 2+ AP: Action potential; Ca 1.1: Voltage-dependent Ca channel skeletal muscle details of the structure of the Cav1.1 (in a closed configur- isoform 1.1; Cryo-EM: Cryogenic electron microscopy; DHPR: Dihydropyridine ation) and of the RyR1 (in closed and ligand-induced open receptor; ECC: Excitation-contraction coupling; EGTA: Ethylene glycol-bis(2- aminoethylether)-N, N, N′,N′-tetraacetic acid; IQ(t): Non-linear capacitive conformations). The next generation of high-resolution currents; k: A measure of steepness in the Boltzmann function; Q(t): Charge cryo-EM, together with electrophysiological assays using movement; Qmax: Maximum charge per unit of linear capacitance; chimeric constructs or site-directed mutagenesis, may Qoff: Charge moved during the repolarization phase of the test pulse; 2+ Qon: Charge moved during the beginning of a test pulse; RyR1: Ca release provide a more comprehensive molecular picture of the channel type 1; SR: Sarcoplasmic reticulum; TT: Transverse tubules; Vh: The interaction between Cav1.1 and RyR1 in their respective mid-point in axis voltage for ½ of Qmax; VSD: Voltage sensor domain membranes. Acknowledgements Conclusions We thank all the contributors of the cited work and express appreciation to Electrophysiological studies and more recently, the solu- the authors in the field of skeletal muscle ECC that, due to space constrains, tion of the structures of the Cav1.1 and the RyR1 at were not referenced in our manuscript. Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 17 of 20 Funding 20. Sandow A. Excitation-contraction coupling in skeletal muscle. Pharmacol This publication was supported by the National Institute of Arthritis and Rev. 1965;17(3):265–320. Musculoskeletal and Skin Diseases of the National Institutes of Health under 21. Dulhunty AF. Excitation-contraction coupling from the 1950s into the new Award Number R37-AR055099 (to M. F. S.). The content is solely the responsi- millennium. Clin Exp Pharmacol Physiol. 2006;33(9):763–72. bility of the authors and does not necessarily represent the official views of 22. Hodgkin AL, Horowicz P. 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A calcium channel mutant mouse model of hypokalemic periodic paralysis. J Clin Invest. 2012;122(12):4580–91. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Skeletal Muscle Springer Journals

Voltage sensing mechanism in skeletal muscle excitation-contraction coupling: coming of age or midlife crisis?

Skeletal Muscle , Volume 8 (1) – Jul 19, 2018

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Life Sciences; Cell Biology; Developmental Biology; Biochemistry, general; Systems Biology; Biotechnology
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Abstract

The process by which muscle fiber electrical depolarization is linked to activation of muscle contraction is known as excitation-contraction coupling (ECC). Our understanding of ECC has increased enormously since the early scientific descriptions of the phenomenon of electrical activation of muscle contraction by Galvani that date back to the end of the eighteenth century. Major advances in electrical and optical measurements, including muscle fiber voltage clamp to reveal membrane electrical properties, in conjunction with the development of electron microscopy to unveil structural details provided an elegant view of ECC in skeletal muscle during the last century. This surge of knowledge on structural and biophysical aspects of the skeletal muscle was followed by breakthroughs in biochemistry and molecular biology, which allowed for the isolation, purification, and DNA sequencing of the muscle fiber membrane calcium channel/transverse tubule (TT) membrane voltage sensor (Cav1.1) for ECC and 2+ of the muscle ryanodine receptor/sarcoplasmic reticulum Ca release channel (RyR1), two essential players of ECC in skeletal muscle. In regard to the process of voltage sensing for controlling calcium release, numerous studies support the concept that the TT Cav1.1 channel is the voltage sensor for ECC, as well as also being 2+ aCa channel in the TT membrane. In this review, we present early and recent findings that support and define the role of Cav1.1 as a voltage sensor for ECC. Keywords: Skeletal muscle, Excitation-contraction coupling, Charge movement, Voltage sensors, DHPR/Cav1.1, 2+ L-type voltage-gated calcium channel, Ca release, RyR1 Background transverse tubular (TT) system [6, 7]. The AP In skeletal muscle, electrical impulses carried by the axons depolarization activates skeletal muscle voltage-gated of motoneurons travel to the nerve endings at the muscle calcium channels (Cav1.1; also known as dihydropyridine endplate (the muscle synapse), where these electrical sig- receptors, DHPR) [8]. The Cav1.1 channels serve as the nals are converted into chemical signals that produce de- voltage sensing machinery for the process of TT polarizing postsynaptic potentials at the neuromuscular depolarization-induced calcium release from the sarco- junction sarcolemma of the muscle fiber [1, 2]. In all but a plasmic reticulum [9] via intracellular sarcoplasmic few “tonic” muscle fibers, these postsynaptic endplate po- reticulum (SR) calcium release channels, the type 1 ryano- tentials elicit a further depolarization of the muscle fiber, dine receptors (RyR1) [10]. This process that begins with carried out by skeletal muscle voltage-gated sodium chan- the muscle AP propagation and results in muscle contrac- nels, initiating and propagating the muscle action poten- tion is known as excitation-contraction coupling (ECC). tial [3–5]. The muscle action potential (AP) travels both This term was coined by Sandow in the 1950s [11]to in- longitudinally away from the fiber endplate along the clude these main events critical for muscle activation, well muscle fiber surface sarcolemma and radially into the fiber before the molecular identities or even the existence of via invaginations of the sarcolemma that form the the molecular players (Cav1.1 and RyR1) was identified or established. Since then we have accumulated an incredible amount of information concerning the structural aspects * Correspondence: mschneider@som.umaryland.edu Department of Biochemistry and Molecular Biology, University of Maryland and molecular and functional details of the ECC process. School of Medicine, 108 N. Greene Street, Baltimore, MD 21201, USA © The Author(s). 2018 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 2 of 20 The knowledge that the muscle contraction was con- discovery and early functional studies of the ECC voltage trolled by electrical signals was already established by sensor and its role and properties, largely carried out the pioneering work of Galvani, Volta, and Walsh [12, during the last quarter of the twentieth century. We 13]. Subsequent studies and discoveries by Nobili, then consider more recent molecular, structural, and Matteucci, Du Bois-Reymond, and Ringer, to mention mechanistic studies, as well as possible future directions. just a few of the pillars, formed the foundations for A detailed review of the cloning of the Cav1.1 and RyR, modern understanding of bioexcitability of muscle and and identification of their skeletal muscle isoforms as 2+ other excitable tissues [14]. The remarkable work by the ECC voltage sensor and skeletal muscle SR Ca re- Hodgkin, Huxley, and Katz [15–19] established the play- lease channel, respectively, is beyond the present scope ing field for the subsequent wave of functional studies and can be found elsewhere [38–42]. dealing with excitability in general, and with the TT volt- age sensor for EEC in particular, as we consider here. Intramembrane charge movement and ECC Voltage sensor charge movements were predicted by Introduction to excitation-contraction coupling Hodgkin and Huxley By the 1950s and 1960s, the processes that initiated and In their classic work on the membrane potential- accompanied skeletal muscle contraction had been stud- dependent ionic conductances underlying the nerve ied from several angles [20, 21]. Muscle biologists and axon action potential, Hodgkin and Huxley [15] pre- physiologists were working collectively trying to de- dicted that any voltage-sensitive process, such as + + cipher the details of the machinery that controls the voltage-dependent Na or K conductance, should be process of muscle contraction using state-of-the-art controlled by mobile charges that are trapped within the techniques from that period. These pioneers made im- membrane but can be displaced in response to changes portant fundamental contributions to understanding in electrical potential energy due to changes in trans- ECC, including the following. (1) Hodgkin and Horowicz membrane voltage. They further predicted that such [22, 23] proposed that the event that normally induces intramembrane charges should give rise to tiny charge muscle contraction is a change in membrane potential displacement currents in response to changes in trans- rather than the longitudinal spread of current along the membrane voltage (Vm), as the putative charges trapped fiber; they also showed that the development of tension within the membrane redistribute within the membrane was dependent on membrane potential and was de- in response to the change in electrical potential. How- scribed by a steep sigmoidal curve of tension as a func- ever, charge displacement currents were not detected by tion of membrane potential. (2) The experiments of Hodgkin and Huxley [15]. In fact, it took over two de- Huxley and Taylor [7] showing activation at the Z disk cades to prove the charge movement hypothesis of in frog muscle fibers and of Huxley and Straub showing Hodgkin and Huxley [15, 37]. In addition, at the time of local activation at the A-I band junction in lizard muscle Hodgkin and Huxley, and continuing well through the [24], together with the localization of the TT system at time of the early experimental studies characterizing the the Z disk in frog muscle [25] and at the A and I band functional properties of voltage sensor charge move- junction in lizard muscle [26], indicated that the trans- ments, the molecular identity of the voltage sensors was verse tubules (TT) of the skeletal muscle fibers form the not even known. Consequently, it was unknown at the network which conducts the surface depolarization radi- time whether the putative and subsequently measured ally into a muscle fiber to initiate contraction. (3) Inves- intramembrane charge movements were generated by tigations started by Ringer [27] and continued by (1) positive charges held near the inside of the mem- Heilbrunn [28], Kamada and Kinoshita [29], and others in- brane at the inside-negative resting potential, moving 2+ troduced the role of Ca as key regulator of striated outward during depolarization (Fig. 1a, right) and muscle activation. Further details of the complex action of returning inward after repolarization (Fig. 1a, left); (2) 2+ Ca on muscle contractile activation were eventually pro- negative charges positioned near the outside of the vided by Weber [30, 31] and Ebashi [32]; reviewed in more membrane at rest, moving inward during membrane detail by Endo [33]. (4) Robertson [26], Andersson-Cedegren depolarization and returning outward during repolariza- [34], Francini-Armstrong and Porter [35], and Peachey tion (Fig. 1a, left); or even (3) dipolar charges rotating in [36], using electron microscopy, described that the ultra- the membrane as the positive end moves outward and structure of transverse tubules (TT) and that the terminal the negative charge moves inward during depolarization cisternae of the SR are in close proximity to the TTs. and reverses this movement during repolarization This year (2018) is the 45th anniversary of the demon- (Fig. 1b). Subsequently, with the establishment of the stration of ECC voltage sensor charge movement [37]. In molecular identity, amino acid sequence, and predicted keeping with the theme of “coming of age/midlife crisis” or experimentally determined molecular structure of the of the ECC voltage sensor, here we will first review the ECC voltage sensor (Cav1.1) [38, 43], as well as those of Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 3 of 20 Fig. 1 Hypothetical mechanisms for a mobile charged intramembrane voltage sensor. a Voltage-dependent intramembrane charge movement could be generated by positive charges moving outward during depolarization, negative charges moving inward, or dipoles rotating. In each, the charge would return to its starting distribution on membrane repolarization, so the charge moved outward during membrane depolarization must equal the charge moving back on repolarization. b Cartoon illustration of current concept of intramembrane voltage sensors as positively charged amino acid residues on transmembrane alpha helices. As in a, the charge moved outward during depolarization (here on an alpha helix) will equal the charge returning on repolarization other voltage-sensitive channels, it is now accepted that were expected to occur over the Vm range where muscle positively charged transmembrane alpha helical “S4” seg- fiber contraction was activated, roughly between about − ments in membrane-spanning domains (Fig. 1b; consid- 50 and + 20 mV [23]. Thus, the putative muscle voltage ered in detail below) are the electrically charged sensor charge displacement current was predicted to be molecular components that serve as voltage sensors for an “extra” non-linear component of the total membrane both ECC and channel gating of plasma membrane and current. Formally, this extra current was “capacitative” in + 2+ + TT Na ,Ca , and K channels [44]. nature since whatever charge moved outward during depolarization was trapped within the membrane and was Voltage sensor charge movements were first detected in obliged to move back to its starting intramembrane loca- skeletal muscle fibers tion when the membrane was repolarized (Fig. 1). To ex- The first successful intentional measurement of the volt- tract the non-linear component from the total measured age sensor charge displacement currents predicted by membrane capacitative current for each “test” pulse (P) Hodgkin and Huxley was carried out on skeletal muscle applied from the holding potential (Fig. 2a,left), the same fibers. Assuming that the charge displacement currents depolarizing pulse (P) was superimposed on a negative would be small, Schneider and Chandler [37] voltage prepulse (ΔV pre) of larger absolute amplitude than clamped frog skeletal muscle fibers in the presence of the test pulse (Fig. 2a,right) [37]. In this way the blockers for each of the major ionic conductances (TTX same amplitude pulse (P) was now applied over a Vm + + + + for Na conductance, Rb replacing K for K conduct- range that was entirely negative to the initial holding ance, methanesulphonate for Cl− conductance). Under potential. This pulse served as the “control” pulse and these conditions, ionic currents were essentially absent. is assumed to contain only the linear capacitative However, the linear capacitative current, needed to current (Fig. 2b,right). charge the linear capacitance of the muscle fiber lipid bi- The currents for both the test and control pulses were layer membrane when the fiber membrane potential was recorded digitally during the experiment using a digital changed, still remained and obscured the putative volt- signal averaging device, which preceded the introduction age sensor charge movement. of laboratory computers and the use of PCs. Using the A strategy was needed for removing the linear capacita- digitized records of total current, the current recorded tive current in order to “unmask” the current carried by for the “control” depolarization (in the range negative to voltage sensor charge movement. The expected voltage the holding potential; Fig. 2b, right) was digitally sensor charge displacement current was anticipated to sat- subtracted from the current for the “test” depolarization urate at highly positive or highly negative membrane po- (over a voltage range positive to the holding potential; tentials, as all mobile charges were maximally displaced Fig. 2b, left) to give the “non-linear” membrane current during large depolarizations or hyperpolarizations (Fig. 1). (Fig. 2c). Assuming any remaining non-linear ionic Furthermore, the voltage sensor charge displacements current to be constant (i.e., time-independent) during Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 4 of 20 cd Fig. 2 Protocol for original recording of intramembrane charge movements. a Pulse protocol used to extract current carried by intramembrane charge movements. The same pulse (P) was applied either from the holding potential (test, left) or superimposed on a hyperpolarizing prepulse (Δ Vpre; control, right). The pulse over the control voltage range is assumed to cause no intramembrane charge movement. b The total current recordings for the pulses in a.Tracesin a and b were recorded as photos of oscilloscope display. c Difference between membrane current in the test pulse minus the current in the control pulse, obtained by digital subtraction of digital recordings of test and control currents using an analog to digital converter and a digital recording system. d Schematic diagram of the three microelectrode voltage clamp systems used in this experiment. All records obtained at the tendon termination of a muscle fiber in a frog sartorius muscle stretched to reduce contraction and bathed in solution to block or remove essentially all membrane conductance. From ref. [37], with modification the pulse, the time-dependent component of the amplitude. In contrast, the time course of the charge non-linear current (test−control) was taken to be the movement current after the pulse (i.e., during fiber repo- non-linear “charge displacement current” or “charge larization to the initial holding potential) did not notice- movement current” due to the voltage sensor movement. ably change in kinetics as the pulse depolarization was This current is the current in excess of the steady increased (Fig. 3a). As discussed further below, from the non-linear current during the pulse (Fig. 2c; upper original report, the charge movement kinetics and voltage dashed line during the pulse), and the current due to the dependence were generally in the range that would be ap- return of the voltage sensors was taken to be the current propriate for muscle contractile activation, so it was not below the initial and final zero current (Fig. 2c; dashed unreasonable to identify these charge movements with the line after the pulse) when the voltage sensors return to intramembrane movement of ECC voltage sensors in the their starting distribution after the pulse. These initial TT membrane [37]. With subsequent sophistication of ex- recordings of voltage sensor currents were made at the perimental and recording procedures, records with better end of a single muscle fiber in an isolated frog sartorius signal to noise and corresponding resolution of kinetic de- muscle using the three microelectrode voltage clamp tails were obtained (a) from frog individual muscle fibers system developed by Adrian et al. (Fig. 2d;[45, 46]). in a single Vaseline gap voltage clamp system which allowed fiber movement without movement artifacts in Critical steps: voltage sensor currents and charge the membrane current records during contractile activa- movements in skeletal muscle tion [47, 48], (b) from frog fibers studied in a double Vas- The original voltage sensor charge movement currents de- eline gap when stretched to eliminate mechanical tected during and after pulses to a range of membrane po- movement and the corresponding movement artifacts tentials using the pulse protocol in Fig. 2a are shown in during activation [49, 50], and (c) from whole cell voltage Fig. 3a [37]. The voltage sensor current amplitude both clamped mammalian short skeletal muscle fibers adhering during and after the pulses increased with increasing to a glass coverslip [51–54]. depolarization (Fig. 3a). The time course of the charge As embodied in the cartoons in Fig. 1, any hypothet- movement currents during the pulse became increasingly ical positive voltage sensor charges that moved outward rapid as the depolarizing pulses were increased in within the membrane during fiber depolarization (or any Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 5 of 20 ab Fig. 3 Initial characterization of intramembrane charge movement currents. a Non-linear difference currents between test pulses to the indicated membrane potentials and corresponding control pulses covering a membrane potential range negative to the holding potential. Same protocol as in Fig. 1. Pulse duration was decreased for the two largest depolarizations displayed. b Equality of charge moved during the “on” and “off” of different amplitude test pulse depolarizations. c Voltage dependence of non-linear charge moved for test pulses to the indicated membrane potentials. All records and graphs from ref. [37], with modification hypothetical negative voltage sensor charges that moved Non-linear charge movement (Q) had a sigmoidal de- inward) are expected to return to their initial intramem- pendence on test membrane potential, according to a brane location when the fiber was repolarized to its ini- two-state Boltzmann function: tial membrane potential. Thus, the amount of voltage sensor charge moved outward during depolarization was Q ¼ Q max=½ 1 þ expðÞ ðÞ −V þ Vh =k anticipated to equal the amount of voltage sensor charge moved inward during the repolarization. This “on/off” where Qmax is the maximum charge (per unit of linear equality of amounts of charge moving outward during capacitance), Vh is the mid-point, and k a measure of depolarization and inward during repolarization was the steepness (Fig. 3c, continuous line through symbols). expected signature of voltage sensor charge displace- While this procedure allows for an approximation of the ment currents (Fig. 3b). From the first report of mea- voltage dependence of the charge movement in ECC, it surements of the ECC voltage sensor currents in muscle may not be adequate to estimate total charge (i.e., total fibers, it was established that the on/off equality criter- number of elementary charges), especially if the charge ion was in fact fulfilled both for the charge movement moves in multiple sequential steps [55]. currents generated by test pulses to various membrane potentials (Fig. 3b) as well as by pulses of various dura- Voltage sensor charge moved predicts pulse durations tions to the same membrane potential [37]. needed to give detectable contraction An immediate question that arose after the first detec- Voltage dependence of charge movement, and its first tion of charge movement currents was whether the volt- interpretation age sensor currents detected in muscle fibers were in The measured amount of charge moved increased in a fact the control system for depolarization-induced con- sigmoidal manner as a function of increasing membrane tractile activation. Two early studies addressed this ques- depolarization from the resting holding potential and tion, using different pulse protocols to show that voltage approached saturation for the largest depolarizations sensor charge movement measurements can be used to used (Fig. 3c). The voltage dependence of charge moved closely predict the initiation of muscle contraction. First, was interpreted using a model in which a single uniform it was previously well-established that during prolonged population of intramembrane charges were each as- (10s of sec) fiber depolarization, fibers first contracted sumed to occupy one of two possible membrane and then became mechanically relaxed [23]. During locations that differed in energy by a fraction of the full similar prolonged voltage clamp depolarizations, muscle electrical potential energy across the membrane. voltage sensor charge displacement properties were also Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 6 of 20 modified [56]. Comparing the time course of recovery of decreased the time needed to reach contractile threshold charge movement after repolarization of fully depolar- during the test pulse (Fig. 4b). The shortening of the test ized fibers with the time for recovery of just-detectable pulse duration for just-detectable contraction could be contraction during repolarization of a depolarized fiber, predicted from the charge movement recordings as the it was found that charge recovery could predict the re- time to move the prepulse charge at the test pulse covery of contractile ability, implying a close relationship voltage. These studies demonstrated a close correlation between charge movement and contractile activation between the voltage sensor charge movement and [57]. Second, during voltage clamp depolarization of just-detectable contractile activation of muscle fibers. fully polarized fibers, the pulse duration required to pro- The charge required to attain a just-detectable contrac- duce a microscopically just-detectable contraction at dif- tion is here termed “pre-activating” charge since it must ferent depolarizations moved a constant amount of be moved in a step or sequence of steps prior to the voltage sensor charge [47, 48]. In this experiment, step(s) that actually activate contraction, but it does not non-linear capacitive currents (IQ(t)), charge movement itself activate contraction (discussed further below). (Q(t)), and the occurrence of just-detectable contraction were all monitored in the same single muscle fiber Voltage sensors control other membrane potential- (Fig. 4a). Contraction was elicited by test pulses to − 45, dependent processes − 35, and − 25 mV, but not by the smallest test pulse to As predicted by Hodgkin and Huxley [15], any − 55 mV (Fig. 4a). Furthermore, using a test pulse alone Vm-sensitive process was expected to involve a voltage (to − 32 mV) or together with two different amplitude sensor charge movement. Indeed, shortly after the initial prepulses (Fig. 4b, bottom), which alone did not produce measurements of intramembrane charge movement in detectable contraction, it was found that the prepulses skeletal muscle fibers, analogous charge displacement Fig. 4 Depolarizing pulses that produce a just-detectable muscle fiber movement displace a set (“threshold”) amount of charge, which can be termed “pre-activating” charge since it must move in order to attain detectable fiber activation. a Charge movement records during muscle fiber depolarization to indicated voltages. The dashed vertical lines indicate the pulse duration needed to give a microscopically just-detectable fiber movement for shorter pulses to the same voltage. No contraction was detected in at − 55 mV. b Pulse to − 32 mV applied alone or together with indicated prepulses, which move only pre-activating charge, since no contraction was detected during the prepulses alone. Charges moved for just-detectable fiber movement (height of black dots) in the test pulses were the same with or without prepulses at the pulse durations for just-detectable fiber contraction (dashed vertical lines). The prepulses decrease the pulse duration required to reach detectable fiber contraction during the test pulse, and this decrease was equal to the time to move the prepulse (pre-activating) charge at the test pulse voltage. Reproduced, with modification from ref. [47] Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 7 of 20 currents were monitored in squid axons [58]. Based on release from the SR (red arrow in Fig. 5; discussed in their properties, the intramembrane charge movements detail below). It is crucial to note that even though iso- 2+ detected in axons were identified as “gating currents” for lated RyR1/Ca release channels of skeletal muscle can + 2+ the axon Na channels. Following the initial work of be activated by elevated Ca [71, 72], it is well estab- 2+ 2+ Schneider and Chandler in muscle and of Armstrong lished that Ca entry via the Cav1.1 Ca channel 2+ and Bezanilla in axons, charge movement of voltage sen- current is not required for activation of RyR1 Ca re- sors has been used extensively to study channel gating lease during muscle fiber depolarization [73], where kinetics and putative voltage-dependent molecular rear- depolarization beyond the reversal potential for L-type 2+ 2+ rangements in a variety of voltage-sensitive channels Ca current [74]orin zeroCa external with EGTA 2+ [59–62] and even in membrane potential-dependent en- [75], which eliminates inward Ca current, does not zymes, pumps, and receptors [63–65]. Over the years, alter muscle activation. Indeed, “skeletal” type of ECC 2+ various pulse protocols (“P/n”,+/− P) have been devised is defined as being Ca influx-independent ([76]; see to extract the non-linear capacitative current (the further discussion below). “charge movement current” or “gating current”) from the total capacitative current [44, 66, 67], in addition to Monitoring and characterizing TT membrane 2+ the P test–P control protocol (Fig. 2a, b) developed for depolarization-induced SR Ca release the initial measurements of muscle voltage sensor An important experimental distinction exists between 2+ charge movement [37]. the two Ca channels regulated by the TT Cav1.1 volt- 2+ age sensor. L-type Ca current can be monitored dir- 2+ A multi-tasking Ca channel: the ECC voltage sensor ectly using the same voltage clamp circuit as used for 2+ controls two distinct Ca channels in two different monitoring voltage sensor charge movement [69, 70]. In 2+ membranes contrast, SR Ca release occurs across the SR mem- Activation of the TT voltage sensor within the Cav1.1 brane, which is not part of the electrical circuit for 2+ molecule controls two different TT voltage-sensitive Ca current flow between the cytoplasm and bathing solu- channels [68]. First, Cav1.1 voltage sensor movement tion that is monitored by the voltage clamp circuit. Con- 2+ 2+ leads to opening of the ion conducting Ca channel sequently, SR Ca release cannot be monitored by the within the Cav1.1/ECC voltage sensor molecule itself voltage clamp system. A second experimental measuring 2+ [69, 70]. This allows Ca influx across the TT mem- system and analysis procedure is needed to calculate SR 2+ brane and into the cytoplasm (blue curved arrow in Ca release. 2+ 2+ Fig. 5), which is manifested as L-type inward Ca The first step in determining SR Ca release is to 2+ current across the voltage clamped TT system. Second, monitor the free myoplasmic Ca concentration during the Cav1.1 voltage sensor movement promotes opening a voltage clamp depolarization [77, 78] (Fig. 6a, b), or 2+ 2+ of the SR RyR1/Ca release channel [8, 9], allowing Ca during an action potential or train of action potentials 2+ 2+ Fig. 5 Cav1.1 (pale blue) serves as voltage sensor for two different Ca channels: its own intramolecular Ca channel in the TT membrane 2+ (current illustrated in blue) and the RyR1 Ca release channel (tan) in the SR membrane (current illustrated in red). Cartoon representation of 2+ 2+ the simplest gating mechanism. RyR1 Ca channel is directly controlled by molecular coupling of Cav1.1 to RyR1. Note that Ca influx via 2+ 2+ the TT Cav1.1 Ca channel is not needed for activation of the RyR1 SR Ca release channel Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 8 of 20 ab 2+ 2+ Fig. 6 Rate of Ca release from the SR during muscle fiber depolarization calculated from the myoplasmic Ca transients measured experimentally 2+ 2+ in individual muscle fibers. a Measured Ca transients (top) and corresponding calculated time course of rate of Ca release from the SR (middle) for 2+ voltage clamp depolarizations to indicated membrane potentials (bottom). The rate of Ca release reaches an early peak and then 2+ declines appreciably during continued depolarization. Reproduced, with modification from ref. [78]. b Time course of recovery of Ca release following an initial inactivating pulse, followed at various times by a repeat application of the same pulse. After a lag of about 100 ms, the early peak begins to recover and is fully recovered by 600 ms. However, at 600 ms recovery, the release wave form is still smaller than in the initial pulse and recovers much more slowly, indicating recovery from a second process, which was attributed to 2+ 2+ 2+ recovery from SR Ca depletion. Reproduced, with modification from ref. [83]. c Ca transients (top) and rate of SR Ca release 2+ (bottom) calculated from the measured Ca time courses for a single action potential or for a train of action potentials. Release in the second and later action potentials is considerably reduced compared to the release in the first action potential. Reproduced, with modification from ref. [79] (Fig. 6c)[79] using a calcium-sensitive indicator dye and amplitude and duration, with a variable time interval be- appropriate optical apparatus [77, 80–82]. However, the tween the two pulses (Fig. 6b). For test pulses applied 2+ measured myoplasmic free Ca transient represents shortly after the conditioning pulse, the time course of 2+ 2+ only a small fraction of the total Ca released during the Ca release completely lacked the early peak 2+ the fiber depolarization. A much larger fraction of the (Fig. 6b). Importantly, the inactivation of Ca release 2+ 2+ released Ca is bound to endogenous myoplasmic Ca during a 20–50-ms pulse does not appear to be due to 2+ binding sites (troponin C, parvalbumin, SR Ca pump) modification of the voltage sensor since charge move- 2+ or transported back to the SR. Taking the Ca binding ment is not modified during these pulses, as judged by properties of these binding sites and transport into con- the criteria Qon = Qoff, and Q kinetics are not modified 2+ 2+ sideration, the Ca release flux (rate of Ca release) after an inactivating prepulse [84]. can be calculated [78]. An important first result of such 2+ calculations was the conclusion that Ca release is not Pre-activating and activating components of charge 2+ maintained during a step depolarization or during a movement for SR Ca release train of action potentials, but instead declines during a In theory, “pre-activating” (also termed “sub-threshold” 20–50-ms step depolarization (Fig. 6a, middle records) or “threshold”) charge movement would be generated by [78] or during a 100-Hz train of actin potentials (Fig. 6c, charge-generating molecular transitions that precede the 2+ lower records) [79]. A slower phase of decline of release actual SR Ca channel opening event in the signaling during longer duration voltage clamp depolarizations pathway from charge movement to RyR1 activation. In 2+ also was observed and was attributed to Ca depletion contrast, the “activating” charge would coincide with 2+ from the SR [83, 84], but the faster developing decline of and determine the actual opening of the SR Ca release 2+ Ca release during a voltage clamp pulse or train of channel. As described above, the pulse duration needed 2+ APs appears to reflect inactivation of SR Ca release. to produce a microscopically just-detectable fiber move- Another important feature of the inactivation is its re- ment for various voltages was found to be the pulse dur- covery, as demonstrated using a double-pulse protocol ation that produced the same constant (= “threshold”) [83]. Here a first conditioning pulse of fixed amplitude amount of charge moved at each voltage, including dur- and duration is followed by a second pulse of the same ing the stepped-on pulse pattern (Fig. 7, inset voltage Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 9 of 20 protocol) [50]. This observation implicates the threshold preceded by the same prepulse, which moved a substan- (or pre-activating) charge as being a precursor that tial amount of the pre-activating charge, but did not ac- 2+ moves before the charge component that actually causes tivate detectible Ca release (Fig. 7, right inset). Using 2+ Ca release and the subsequent contractile activation. these data, it was possible to systematically separate the 2+ 2+ Using Ca -sensitive dyes and the Ca release calcula- charge movement that was predominantly precursor 2+ tion described in the preceding section, together with (“pre-activating”) for producing SR Ca release (and the stepped-on pulse protocol (e.g., Fig. 7, lower left was moved in the prepulse) from charge that was pre- 2+ inset), it became possible to relate the activating compo- dominantly activating for Ca release (and was moved 2+ 2+ nent of charge movement to the rate of SR Ca release, during the test pulse) (Fig. 7, inset right). Ca transients the process directly downstream of voltage activated were simultaneously measured for the same pulses, and 2+ charge movement, and thus directly controlled by the the Ca release time course was calculated for each test voltage sensor [50]. pulse (Fig. 7, inset top left). Over a wide range of test pulse amplitudes and durations (Fig. 7, voltage protocol), 2+ 2+ Peak SR Ca release due to a pulse is proportional to the the peak rate of Ca release during the test pulse was amount of activating charge moved by the pulse found to increase linearly with the charge that was 2+ To relate Ca release to charge movement, charge moved by the test pulse (Fig. 7). The linear relationship movement was determined for a range of test pulse am- had a small positive charge value for the extrapolation to plitudes and durations, with each test pulse immediately zero peak rate of release (x intercept), indicating a small amount of pre-activating charge that was not moved during the subthreshold prepulse, but was instead moved during each test pulse, presumably the initial charge moved during the test pulse. These results dem- onstrated a close relationship between the extent of acti- 2+ vation of SR Ca release by a pulse and the amount of activating charge that moved during the same pulse [50]. Minimal model for voltage sensor control of a coupled 2+ RyR1 Ca release channel Figure 5 presents a cartoon of the functional states of 2+ the voltage sensor/L-type Ca channel in the TT mem- 2+ brane (top) and the RyR1 Ca release channel in the SR membrane (bottom) for a hypothetical minimal (two-s- tate) model [85] for regulation of RyR1 by its directly coupled TT voltage sensor(s). In this highly simplified gating scheme, fiber depolarization (top, left to right) 2+ Fig. 7 The peak rate of Ca release evoked by various depolarizing causes the mobile voltage sensor charges within the TT test pulses is linearly related to the amount of “activation” membrane voltage sensor protein (Cav1.1), which are intramembrane charge moved by the same pulse. Each test constrained to remain within the TT membrane, to re- pulse was immediately preceded by a depolarizing prepulse (see spond by generating intra-membrane movement (de- pulse schematic in lower left inset), which by itself did not 2+ tected by charge movement measurements [86]). In the activate Ca release. Many of the test pulses were too short to establish the ionic current baseline for calculating charge moved minimal scheme of Fig. 5, the voltage sensor charge during the test pulse, so Qon could not be measured for short movement obligatorily induces the opening of the RyR1 test pulse durations due to uncertainty regarding the level of 2+ Ca channel directly coupled to the voltage sensor(s) ionic current remaining during the test pulse. Charge moved by 2+ (bottom; left to right), resulting in a release of Ca from the test pulse was consequently determined as Qoff–Q pre. The the SR, as detected by monitoring the total increase in test pulse charge represents an upper estimate of the activating 2+ charge moved during the test pulse; the x intercept on the cytoplasmic and transported Ca . In this minimal graph is interpreted as representing the amount of pre-activating model, there are only two states of the voltage sensor (i.e., precursor) charge still present in the test pulse (i.e., pre-activating and its directly coupled RyR1 unit: voltage sensor rest- charge not moved by the prepulse). The lower right inset gives the ing/RyR1 closed (left) or voltage sensor active/RyR1 charge movement current records obtained using various duration test open (right). In addition, in this minimal scheme, the pulses to 0 mV, each immediately following the prepulse to − 50 mV. 2+ 2+ 2+ The upper left inset gives the Ca release calculated from the Ca L-type Ca channel in the TT membrane is also open transient recorded simultaneously with the charge movement records when the voltage sensor is active [8]. shown in the lower right inset for each of the same test pulses. The minimal model (Fig. 5) already raises several basic Reproduced, with modification from ref. [50] issues regarding the TT voltage-dependent gating of Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 10 of 20 RyR1. First, intramembrane charges (transmembrane charge movement, it is necessary to consider inactivation 2+ positively charged S4 alpha helices in the voltage sensor, of the SR Ca release channel, which occurs during the Cav1.1) move outward during TT depolarization fiber depolarization (Fig. 6)[83, 87]. The inactivation 2+ (i.e., away from the RyR). However, the resulting mo- process obscures the time course of Ca release channel 2+ lecular rearrangements in the voltage sensor accompany- activation, reducing Ca release to a fraction of the ing or following charge movement could bring other peak value reached earlier during the pulse (Fig. 6a, b) domains of the Cav1.1 voltage sensor (alpha 1) sub- [87]. One approach eliminated the effects of inactivation unit, or beta subunit either toward or away from the during a pulse by pre-inactivating the fibers (Figs. 6b RyR1, so at present we do not know whether the and 8a). Here a large “inactivating” prepulse, sufficient 2+ voltage-sensitive step constitutes the removal of an to produce maximal inactivation of RyR1 Ca release, inhibition to RyR1 opening, or the application of a was immediately followed by a brief repolarization (to positive factor for voltage-dependent RyR1 activation. return all charge to the resting state and to turn off the 2+ This basic issue awaits high-resolution molecular non-inactivating Ca release) and then by a test structure-function studies of the Cav1.1/RyR1 inter- depolarization during which activation of both the 2+ action in various functional states. Second, the gating “non-inactivating” component of Ca release and the scheme in Fig. 5 includes a major simplification. If charge movement (which does not inactivate in a each of the two ellipses shown in Fig. 5 represents few hundred ms time window: see above) were mon- one of the 4 Cav1.1 α1 subunits coupled to a single itored (Fig. 8a, b)[83, 88]. Using this approach it RyR1 molecule, then each Cav1.1 (i.e., each ellipse) was found that both the voltage dependence (Fig. 8c) 2+ should have four S4 segment charged transmembrane and time course (Fig. 8b) of the non-inactivating Ca α helices, one in each of the four transmembrane do- release very closely agreed with the voltage depend- mains of each Cav1.1, rather than the single charged ence and time course of (Q/Qmax) [88], where helix shown for each Cav1.1 in the simplified cartoon Qmax is the maximum charge, moved during a large in Fig. 5. depolarization This remarkable finding is consistent A clear shortcoming of the minimal scheme in Fig. 5 with a reaction scheme in which the voltage depend- is that all of the voltage sensor charge movement is dir- ence and kinetics of the non-inactivating component 2+ ectly involved in the closed to open transition of the of SR Ca releaserateiscontrolledbyfouridentical RyR1 to which it is coupled (i.e., all of the charge moved and independent voltage sensors [88]. Each RyR1 is “activating” charge movement, as defined above). This channel would be controlled by four identical voltage property clearly does not agree with the experimental sensors and is open when and only when all four characterization of the control system in muscle fibers. voltage sensors are in the active conformation, giving An appreciable fraction of the total charge movement rise to release being proportional to the fourth that is recorded from a fiber is pre-activating charge, power of Q/Qmax. which moves during depolarization, but prior to the If one “voltage sensor” is one Cav1.1 molecule, the charge for the actual activation step in the control voltage sensors could be independent (i.e., in different 2+ mechanism for Ca release (above) [50], yet is an es- molecules), and the RyR1 could require all four sential prerequisite for depolarization-activated SR Cav1.1s to be active in order for RyR1 opening. In- 2+ Ca release. In the following sections, we will exam- deed, early freeze fracture studies using electron mi- ine two functional studies that consider more compli- croscopy (EM) revealed that Cav1.1 forms four cated models for voltage sensor/RyR1 interaction(s), ordered clusters (tetrads) and the space between tet- 2+ as well as other changes occurring in the individual rads suggested an overall 1:2 ratio of tetrads to Ca components, leading to the need for increased com- release channels [89], corresponding to the observa- plexity and refinement of the minimal gating scheme tion that only half of the RyR1 homotetramers in the in Fig. 5. triad junction were coupled to Cav1.1 tetrads. How- ever, each Cav1.1 molecule contains four positively 2+ Model for RyR1 Ca release activation requiring charged S4 transmembrane helices, as described simultaneous activation of four identical but independent below. One possibility is that in the Cav1.1-RyR1 voltage sensors complex in a functioning muscle fiber, interaction be- In a preceding section and Fig. 7, we considered the em- tween the Cav1.1 and RyR1 gives rise to the condition 2+ pirical (linear) relationship between the peak rate of Ca that only one of the four S4 helices in each Cav1.1 is release produced by a given pulse and the activating physically able to move, resulting in a single mobile charge moved in the same pulse [50]. In order to next charged group per Cav1.1, with four independent 2+ relate the time course of Ca release activation during voltage sensors (one in each Cav1.1 of the tetrad) per 2+ TT depolarization to the time course of voltage sensor coupled RyR1 homo tetramer Ca release channel. Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 11 of 20 ab Fig. 8 Empirical evidence for a model for activation of each RyR1 in which four identical and independent voltage sensors must all be in the 2+ 2+ active configuration for the SR Ca release channel to open after pre-inactivation of the RyR1 channel. a Ca transients (top) and calculated rate 2+ of SR Ca release (middle) for the pulse protocol at the bottom. Pulses to various test pulse amplitudes were each preceded by the same inactivating prepulse, followed by brief return to the holding potential to reset the voltage sensors and close the RyR1 channel and then the 2+ 4 2+ various depolarizations (bottom). b Wave form of Ca release during the test pulses closely follows Q . c Voltage dependence of Ca released follows the fourth power of charge moved (dashed curve). From ref. [88], with modification Allosteric mechanisms for RyR channel control by TT justify the added transitions of the allosteric model, add- Cav1.1 voltage sensor itional experimental data are required and were intro- 2+ In the preceding model, the Ca release channel opens duced [92]. Also note that each “voltage sensor” when and only when a certain enabling configuration of considered here, as in the four independent voltage sen- the voltage sensors was achieved. An allosteric model sor models above, is an entire Cav1.1, containing a [90, 91] for gating of each RyR1 homo-tetramer by four charged transmembrane “S4” helix in each of its four independent DHPRs provides an alternative approach transmembrane domains. (Fig. 9a, b). In the allosteric system, each RyR1 channel can open when any number of its coupled voltage sen- Molecular components and mechanisms in ECC sors is active [92]. However, RyR1 opening becomes in- Cav1.1: the TT voltage sensor creasingly likely as more voltage sensors are active Membrane depolarization of the TT system, during an (Fig. 9b). In this model, each RyR1 channel has two AP or voltage clamp step depolarization, is detected by states, closed (C) and open (O). Lateral transitions rep- Cav1.1 channels, the TT voltage sensor. Cav1.1 channels resent opening or closing transitions of the RyR1. Each were initially identified using electrophysiological ap- voltage sensor has two states, inactive or active (− or +, proaches (charge movement and ionic currents) [37, 56, respectively in cartoon). Vertical transitions represent 69, 70, 93]. Cav1.1 channels are principally expressed in changes in the activation (+) or deactivation (−) status of the membrane of the TT system of adult skeletal muscle each of the four voltage sensors, as indicated by the four fibers and are members of a diverse family of 2+ circles with + or − representing the four voltage sensors voltage-dependent Ca channels. Molecular details of 2+ controlling each RyR1 [92]. It should be noted that re- the voltage-gated Ca channels from skeletal muscle moving the open states O through O , which are open were first identified by binding, purification, and recon- 0 3 and have less than four active voltage sensors, and the stitution [94]. Using molecular biology techniques, their transitions to and from each of these states removes the amino acid sequences were determined by cDNA clon- possibility of opening without movement of all four volt- ing and sequencing [43]. Contrasting with their promin- age sensors, and thereby reduces the allosteric model to ent functional status, TT voltage sensor (Cav1.1 the four independent voltage sensor model. Thus, the channels) was somewhat apart from the saga of allosteric model includes the four voltage sensor models, structure-function studies for other types of ion chan- 2+ which already fit the data for non-inactivating release nels [95]. The skeletal muscle Ca channel complex is a very closely [88], as a subset of possibilities. In order to hetero-tetramer, comprised of a main pore-forming α1 Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 12 of 20 Fig. 9 Allosteric model for gating of each coupled RyR1 by four different voltage sensors. a Cartoon of a single RyR1 homo-tetramer controlled by four independent voltage sensors, with all four voltage sensors in the inactive configuration (left) or with one voltage sensor in the activated 2+ configuration (right). b Kinetic reaction scheme for the RyR1 Ca channel gating process. Channel opening becomes more likely as more voltage sensors become active, which is the basis for the allosteric effect. However, the channel can open with any number of voltage sensors active. The subscript i on C or O is the number of voltage sensors active. Forward rate constant k for Ci→ Oi increases with i; the backward rate constant k for Oi→ Ci decreases with i. Note that f < 1. Elimination of states O to O from this scheme reduces it to the model proposed in Fig. 8. From ref. 0 3 [92], with modification 2+ subunit, complexed with β, α2δ, and γ subunits [38, 39, of Ca channels and their structural examination [95], 96] (Fig. 10a) and other ligands (i.e., STAC3) [97]. The which is changing with the molecular interpretation of Cav1.1 α1 subunit consists of a single polypeptide chain, their function. Recent cryo-EM studies at a resolution of with four highly homologs but non-identical intramem- 3.6 Å [100] revealed more details about the molecular brane domains (I–IV), each containing six transmem- architecture of the Cav1.1 channel of skeletal muscle with brane (TM) alpha helical segments (S1–S6), shown in its complete set of auxiliary subunits (Fig. 10a). The cen- cartoon representation in Fig. 10b [43, 96], as well as tral α1-subunit of CaV1.1 has a core structure and is asso- amino and carboxyl terminals. While the organization of ciated with an extracellular α2δ-subunit, an intracellular the TM domains of the Cav1.1 α1 subunit has a strong β-subunit, and a 4-TM γ-subunit (Fig. 10a). pseudo fourfold symmetry in the plane of the TT mem- brane, the intracellular structure Cav1.1, including the 2+ RyR1, the SR Ca release channel in skeletal muscle single β subunit is highly asymmetrical, which could 2+ 2+ RyR1-dependent SR Ca release via the Ca release have important implications for Cav1.1-RyR1 coupling. channel RyR1 initiates muscle contraction. The RyR1 is Segments S1–S4 of each transmembrane domain of the a colossal protein of approximately 2.3 MDa assembly of α1 subunit form a voltage-sensing domain (VSD) [44, four identical subunits [101, 102]. Each subunit contains 98], whereas segments S5 and S6 from all four intra- 2+ an intramembrane region, located within the C-terminal membrane domains contribute to the Ca -conductive region and representing ca. 20% of the total protein, plus pore (Fig. 10b and ribbon diagram in Fig. 11a [98]). The a cytoplasmic region that represents 80% of the total fourth TM helical segment (S4) contains a number of protein, and is known as the foot region (Fig. 11b, blue positively charged amino acids (Arg and Lys), separated structure; note that the spatial scale is about five times by two hydrophobic residues (Fig. 10b)[43, 98]. During compressed to Fig. 11a, so Cav1.1 appears much smaller changes in TT membrane potential, the S4 segments are in Fig. 11b than a); [103–107]. The cytoplasmic region believed to rearrange, moving outward across the plane of the RyR1 channel (280 Å × 280 Å × 120 Å) is continu- of the TT membrane, in response to membrane ous with the transmembrane region (120 Å × 120 Å × depolarization, establishing the determinants for voltage 60 Å; Fig. 11b)[41]. The RyR1 SR transmembrane re- sensitivity [9, 99]. Historically, there was a considerable 2+ gion forms the Ca release channel [106–109]. time gap between recognition of the biological importance Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 13 of 20 ab Fig. 10 Architecture and membrane topology of the Cav1.1. a Side view of a cryo-EM reconstruction of the Cav1.1 and its subunits at 3.6 Å (Protein Data Bank (PDB) 5GJW) [100]. The Cav1.1 α1 subunit is highlighted in blue. Auxiliary β-, α2δ-, and γ-subunits are colored in green, red, and yellow, respectively. This model was created in Chimera [174]. The asterisk symbol indicates the missing II–III loop sequence in the cryo-EM structure. b Membrane topology of the Cav1.1 α1 subunits. The α1 subunit is composed by an interconnect array of four homologous (but not identical) domains, each domain consisting of six transmembrane domains, S1–S6. S1–S4 from each domain form a voltage sensor domain, whereas S5 and S6 from all four domains form the pore domain. Intracellular loops connect the domains; the loops II–III and I–II are important for ECC, as indicated ab Fig. 11 Structural details of the Cav1.1 α1 subunit and the RyR1. a Shown are cryo-EM reconstructions of the Cav1.1 a subunit, side view (top) and upper view (bottom) (PDB 5GJW) [100]. The auxiliary subunits, the cytoplasmic tails, and loops are not shown. Each domain is color coded, positively charged residues are indicated in red, negatively charged residues are shown in blue. The dashed ellipse indicates the location of S1–S4 from domain I and the black circle shows the corresponding pore domain (S5–S6) from domain I. Note that each voltage sensor domain (S1–S4) is not in close proximity to its corresponding pore domain (S5–S6). b Side (top) and upper (bottom) views of cryo-EM reconstruction of the RyR1 (PDB, 5TAL) [175] and four superimposed Cav1.1 α1 subunits (PDB 5GJW) [100], forming a tetrad. This model was created in Chimera [174] using the cryo-EM maps with relative location of the Cav1.1 subunits as suggested by Samsó [118] Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 14 of 20 2+ RyR1s are arranged in a regular array within the ter- reversal potential, where Ca influx is greatly sup- minal cisternae of the junctional SR [89, 110, 111]. Simi- pressed or eliminated [74]. Second, the frequency of oc- 2+ 2+ larly, Cav1.1 channels are clustered in groups of four (or currence of Ca sparks, elementary intracellular Ca tetrads) in the TT membrane that is adjacent to the signals [129, 130], in frog skeletal muscle fibers can be 2+ junctional SR [89, 112], with a Cav1.1 tetrad facing every used as a measure of activation of “microscopic” Ca other RyR1 in the junctional SR RyR1 array (Fig. 11b). release events, which combine at high frequencies dur- For symmetry and simplicity, each Cav1.1 molecule ing fiber depolarization to produce the “macroscopic” 2+ composing a tetrad is believed to be oriented in the Ca transient observed during depolarization [131, same coordinated position relative to the subunits of its 132]. These events are extremely infrequent in resting fi- apposed RyR1homo tetramer (see Fig. 11b)[89, 112]. bers, but increase tremendously in frequency during Since these interfaces take place at alternate RyR1s, half depolarization, so much so that spark frequency can of the RyR1s are “uncoupled” with Cav1.1s, and half are only be monitored experimentally during large depolar- coupled to Cav1.1s [113]. The location of RyR1 (coupled izations by using depolarized fibers, and restoring only a and uncoupled) determines the organization of the small fraction of the release units by brief repriming re- Cav1.1 channels in the juxtaposed TT membrane, creat- polarizations [130, 133–135]. These types of experiments ing a “checkerboard” array of coupled and uncoupled indicate that in functioning muscle fibers, the RyR1 SR 2+ RyR1s that produces the Cav1.1 lattice organization Ca release channels are essentially fully off when the [114]. Depolarization-induced activation of RyR1 is be- voltage sensors are in the resting condition, but turn on lieved to be mediated via direct or indirect interactions strongly and rapidly during the AP or voltage clamp with TT voltage sensors [8, 115–117]. However, despite depolarization that activates the voltage sensors [132, extensive biophysical and ultrastructural studies, the 135, 136]. The simplest hypothesis is that under normal molecular basis for TT voltage sensor function and the conditions in mature functioning muscle fibers, RyR1s chemical mechanisms that support TT voltage regulated coupled to TT voltage sensors are locked in the off con- 2+ RyR1 SR Ca release, as well as the orientation of the figuration due to an inhibitory influence of the voltage Cav1.1 tetrads relative to the RyR1 have remained un- sensor in its resting configuration [137]. Movement of clear. In Fig. 11b, the location of the tetrads is based on the voltage sensor into the active configuration during reference [118], whereas the relative orientation is depolarization removes this “lock” on RyR1 opening, and arbitrary. the RyR1 channels open and, more slowly, inactivate The RyR1 components, including SR luminal seg- (Fig. 6a)[78]. When the voltage sensor returns to the ments, transmembrane domains, and large cytosolic do- resting configuration at the end of the depolarization, mains, and their interaction with the Cav1.1 channels, the RyR1 is relocked. In this scenario, all the other li- SR luminal proteins, and accessory proteins, metabolites, gands that modulate RyR1 channel activity in isolated and ions, as well as post-translational modifications, membrane or protein preps may or may not also simi- allow the RyR1s to be fine-tuned by numerous mecha- larly modulate the RyR1 in a functioning fiber, but the nisms [71, 107, 109, 119–128]. However, the TT voltage TT voltage sensor serves as master regulator determin- sensor (CaV1.1) is believed to be the “ligand” that ing whether or not the channel can open at all. uniquely enables RyR1 opening in functioning skeletal muscle fibers. Dysgenic muscle and restoration of skeletal vs. cardiac The TT voltage sensor is the master regulator of RyR1 ECC Despite the large number of modulatory interactions Advances in biochemistry, molecular biology, and that influence RyR1 activation, it is difficult to pharmacology allowed the identification of the molecular overemphasize the importance of the TT voltage sensor, components that are essential for ECC. One crucial dis- and/or some component(s) directly coupled to it, in the covery was the characterization of a naturally occurring physiological regulation of RyR1 in skeletal muscle. In- “knock out” of the Cav1.1 α1 subunit (“dysgenic” mouse; deed, the voltage sensor can be considered as the master [138]). This model demonstrated that myotubes derived 2+ ligand for controlling SR Ca release in normal mature from the dysgenic mice lacked ECC and intramembrane muscle fibers. First, no other component, except possibly charge movement; the expression of α1s subunit (skel- 2+ Ca influx into the TT/SR gap, would be anticipated to etal muscle isoform) in these cells restored “skeletal” 2+ change drastically during the few millisecond action po- type of ECC, which is independent of Ca influx [139]. 2+ tential which raises cytoplasmic Ca sufficiently to The expression of the cardiac isoform (α1c subunit) of 2+ cause a twitch contraction. However, Ca influx is not the Cav1 channels did not restore skeletal ECC [140]. needed for skeletal muscle activation [73], which is The inability of the cardiac isoform [140] and of other 2+ 2+ maintained for depolarizations well beyond the Ca Ca channel subtypes [141, 142] to rescue the ECC Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 15 of 20 allowed the investigation of essential elements for skel- essential for the skeletal muscle function. Skeletal ECC etal muscle ECC via chimeric channels. was not experimentally evident if Cav1.1/RyR1 were not in the membrane, or forming tetrads. Components of Cav1.1 that are needed for activating RyR1 Discrete S4 voltage sensor (i.e., S4, I–S4, IV) models Chimeric channels made with α1 subunits of Cav1.1 and In voltage-gated sodium channels, a channel structurally Cav1.2 demonstrated that a region in the intracellular and evolutionary similar to Cav1.1 [39], studies using mu- loop between the second and third domains (II–III tagenesis and voltage-clamp fluorometry revealed that the loop), specifically, the region spanning residues 720– four VSDs, each linked to a partial pore-forming region, 764/5, was important for this function (Fig. 10b)[143, may be differentially and allosterically coupled to the pore 144]. Interestingly, in the cryo-EM structure of Cav1.1 opening to various degrees of involvement in the control the structure of the II–III loop is undefined (dashed line of voltage dependence and gating [163]. VSDs I–III acti- in Fig. 10a,[100]), whereas the I–II and III–IV/C-ter- vate in parallel and sufficiently rapidly to modulate Na minal regions are defined and appear in Fig. 10a. While channel opening, whereas VSD IV activates more slowly the identification of the Cav1.1 regions that are critical and initiates fast inactivation [163]. In Cav1.2, a 2+ for ECC has been more active, perhaps due to the voltage-gated Ca channel expressed in cardiac cells, smaller size of the Cav1.1 channel, the identification of site-directed fluorophore labeling, and voltage clamp binding domains in the RyR1 for the II–III loop has fluorometry of individual VSDs showed differential func- been less fruitful. Only a few reports, where deletions of tion for each domain; VSDs II and III exhibited large segments of the RyR1 successfully altered ECC, voltage-dependent and kinetic characteristics compatible concluded that several regions of the RyR1 are involved with channel activation [164]. However, the cardiac iso- in the interaction with Cav1.1 [145–147]. Recent ap- form (α1c subunit) is unable to support RyR1 activation in proaches are revisiting the role of the loop II–III and its myotubes lacking α1s [140, 144]. 2+ association with the adaptor protein STAC3 on ECC Similarly, the voltage dependence and timing of Ca [148]. These new results support the notion that the II– entry via Cav1.1, as well as the voltage dependence and 2+ III linker plays a role in ECC. timing of TT voltage-dependent RyR1 Ca release, are ex- The Cav1.1 I–II loop is the site for interaction with the pected to be functions of the α1-subunit of Cav1.1, which β1a subunit (Fig. 10b)[98]. The β1a subunit is important also contains four highly similar but non-identical VSDs, for several aspects of ECC. The β1a subunit is needed for I–IV [43, 96, 98]. Evidence for a differential role of each the functional expression of Cav1.1 α1 subunit [149]and VSD in Cav1.1 channel operation using chimeric studies is crucial for enhancement of Cav1.1 α1 triad expression (interchange of VD SI region, Cav1.1↔Cav1.2) and alter- [150], assembly of Cav1.1 α1in tetrads [151, 152], and native splicing of VSD IV of Cav1.1 suggests that VSDs I elicitation of Cav1.1 α1 charge movement [153]. and IV control the activation kinetics and voltage depend- Skeletal-type ECC is reduced in muscle cells lacking the ence, respectively [165–167]. Because VSD I and VSD IV expression of β1a [150] and is rescued by expression of appear to be linked to the slow activating Cav1.1 ionic β1a [154]. The use of chimeric constructs of β1a [155] current, it was hypothesized that VSDs I and IV do not 2+ with other β subunits, as well as the use of synthetic pep- contribute to the more rapid Cav1.1-dependent SR Ca tides [156], allowed the identification of the C-terminal re- release [168, 169] (see Fig. 12). In support of this hypoth- gion of β1a as an important domain for possible esis, a functional study of a mutation causing malignant interaction with RyR1 during TT voltage-dependent SR hyperthermia susceptibility (R174W) in S4 of VSD I of 2+ Ca release. Cav1.1 revealed that this mutation reduces Cav1.1 ionic 2+ Thus, several sites in the Cav1.1 α1 subunit, in current, but does not affect Cav1.1-dependent SR Ca re- addition to the II–III loop, contribute to the overall lease [170](Fig. 12). Wu and colleagues [171]used a Cav1.1/RyR interaction. These include the loop I–II and mouse model for hypokalemic periodic paralysis with a β-subunit [157, 158], and more indirectly, the III–IV targeted Cav1.1 R528H mutation in S4 of VSD II. Muscle loop [159] and the C-terminal domain of the Cav1.1 α1 fibers from the Cav1.1 R528H homozygous mouse exhib- 2+ [141] (Fig. 10)b. ited impaired depolarization-induced Ca release, sug- Another important advance in the characterization of gesting that VSD II could participate in Cav1.1-dependent 2+ the molecular players of the ECC and their interactions SR Ca release [171](Fig. 12). These results represent was the generation of a RyR1-knockout mouse (the dys- compelling but still indirect evidence of the role of each pedic mouse [160]), which allowed for the expression of VSD in Cav1.1. Currently, the contribution of the individ- various RyR constructs and different Cav1.1 α1/RyR1 ual VSDs to the voltage dependence of Cav1.1 pore open- 2+ combinations [161, 162]. These approaches identified ing and activation of RyR1 Ca release is unknown. Note that the skeletal Cav1.1 α1-subunit and RyR1 are that the Cav1.1 R528H mutation also introduces a “gating Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 16 of 20 Fig. 12 Voltage sensing domains of Cav1.1. From left to right, S4-I, S4-II, S4-III, and S4-IV. The positively charged residues of the S4 segments are labeled in red for each domain. Negatively charged amino acids are shown in blue. The depicted transmembrane helices are from the structure of the cryo-EM of Cav1.1 (PDB 5GJW) [100]. These models were created using Chimera [174]. Refs. for VDSI [165, 170], VDSII [165, 171], VSD III [165, 167], and VSD IV [165–167] pore” or “omega” current, which is normally not present near-atomic level, have provided in depth functional and in the wild-type channel and is responsible for the anom- structural details of the ECC process. However, it is clear alous depolarization seen in hypokalemic periodic paraly- that new approaches are needed to continue to explore sis [171]. Details regarding the gating pore current have the intricacies of ECC. Some of the many remaining un- been reviewed in [172, 173]. answered questions regarding ECC include the follow- ing: Are all four VSDs (I–IV) needed to activate the 2+ Future perspectives RyR1 Ca channel, or is only a subset of charges in- How does the propagated electrical impulse spreading volved? If so, which VSDs are coupled to Cav1.1 pore 2+ along the TT system produce Ca release? The Schneider opening? Which VSDs contribute the voltage sensor ele- and Chandler hypothesis that the excitatory signal passes ment(s) for electromechanical coupling between the 2+ from the TT to the SR membrane by way of charges mov- Cav1.1 and RyR1 Ca release? Which residues are moved ing in the TT membrane connecting with the junctional during Cav1.1 channel activation? Which residues are 2+ 2+ feet of the SR Ca release channel initiated the path to moved for RyR1 Ca release channel activation? How far answer this question. Yet, it is still unknown how this do they move within the membrane electric field? And movement of charge, originating in Cav1.1, transfers a sig- what are the molecular determinants that mediate the 2+ nal across to the RyR1 (feet) to trigger Ca release. This electromechanical coupling between the VSD and RyR1 2+ question is particularly fascinating because of the as yet Ca release? These are some of the interesting questions unknown molecular structure-function relationship be- for current and future investigation. tween these components in two different membrane sys- tems, the TT (Cav1.1 voltage sensors) and the SR (RyR1 2+ Abbreviations Ca release channels). Cryo-EM has revealed amazing 2+ AP: Action potential; Ca 1.1: Voltage-dependent Ca channel skeletal muscle details of the structure of the Cav1.1 (in a closed configur- isoform 1.1; Cryo-EM: Cryogenic electron microscopy; DHPR: Dihydropyridine ation) and of the RyR1 (in closed and ligand-induced open receptor; ECC: Excitation-contraction coupling; EGTA: Ethylene glycol-bis(2- aminoethylether)-N, N, N′,N′-tetraacetic acid; IQ(t): Non-linear capacitive conformations). The next generation of high-resolution currents; k: A measure of steepness in the Boltzmann function; Q(t): Charge cryo-EM, together with electrophysiological assays using movement; Qmax: Maximum charge per unit of linear capacitance; chimeric constructs or site-directed mutagenesis, may Qoff: Charge moved during the repolarization phase of the test pulse; 2+ Qon: Charge moved during the beginning of a test pulse; RyR1: Ca release provide a more comprehensive molecular picture of the channel type 1; SR: Sarcoplasmic reticulum; TT: Transverse tubules; Vh: The interaction between Cav1.1 and RyR1 in their respective mid-point in axis voltage for ½ of Qmax; VSD: Voltage sensor domain membranes. Acknowledgements Conclusions We thank all the contributors of the cited work and express appreciation to Electrophysiological studies and more recently, the solu- the authors in the field of skeletal muscle ECC that, due to space constrains, tion of the structures of the Cav1.1 and the RyR1 at were not referenced in our manuscript. Hernández-Ochoa and Schneider Skeletal Muscle (2018) 8:22 Page 17 of 20 Funding 20. Sandow A. Excitation-contraction coupling in skeletal muscle. Pharmacol This publication was supported by the National Institute of Arthritis and Rev. 1965;17(3):265–320. Musculoskeletal and Skin Diseases of the National Institutes of Health under 21. Dulhunty AF. Excitation-contraction coupling from the 1950s into the new Award Number R37-AR055099 (to M. F. S.). The content is solely the responsi- millennium. Clin Exp Pharmacol Physiol. 2006;33(9):763–72. bility of the authors and does not necessarily represent the official views of 22. Hodgkin AL, Horowicz P. 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Skeletal MuscleSpringer Journals

Published: Jul 19, 2018

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