1Department of Anesthesia Research, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts 02115; 2Department of Molecular Biosciences, School of Veterinary Medicine, University of California, Davis, California 95616; 3Laboratory of Cellular Physiology, CeSI Center for Research on Ageing, Università degli Studi G. d'Annunzio, 66013 Chieti, Italy; and 4Department of Life Science, Kwangju Institute of Science and Technology, Kwangju 500-712, Korea
Submitted 1 May 2003 ; accepted in final form 11 September 2003
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ABSTRACT |
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ryanodine receptor type 1; dihydropyridine receptor; excitation-contraction coupling; negative module
Expression of the cDNA encoding RyR1 in dyspedic 1B5 myotubes (RyR1/) restores skeletal type EC coupling and rescues Ca2+ current through the DHPR (27). The latter is a result of the recovered retrograde signaling from the RyR to DHPR in dyspedic muscle. Neither orthograde nor retrograde signaling is restored by expression of cDNAs encoding RyR2 or RyR3 in dyspedic myotubes (8, 29, 33, 38). In an attempt to define which regions of RyR1 are responsible for orthograde and retrograde signaling with the DHPR, we have found that both orthograde and retrograde signaling are strongly restored by the R10 region (1,6352,636 aa). The smaller R16 region (1,8372,154 aa, contained within the R10 region) mediates only weak orthograde signaling. In the case of the adjacent R9 region (2,6593,720 aa), orthograde signaling is weekly restored but retrograde signaling is strongly restored (28, 36, 38).
In this study, allosteric interactions between the DHPR and RyR1 were further examined by expressing cDNAs encoding wild-type RyR1, RyR2, and four RyR1-RyR2 chimeras [R4, R9, and R10 based on Nakai et al. (28), and R16 based on Proenza et al. (36)] in 1B5 myotubes by using helper-free Herpes Simplex virus 1 (HSV-1) amplicon virions (49). This study revealed that at rest, before activation, the DHPR exerts a negative long-range allosterism on RyR1, reducing its sensitivity to direct agonists such as caffeine. This repression can be lifted pharmacologically with DHPR blockers at concentrations that place the DHPR into the preactivated state. Using RyR1-RyR2 chimeras, we were not able to discover the location of the negative module but demonstrated that the DHPR engages long-range allosteric interactions with the R9 region of RyR1 (2,6593,720 aa), manifested as decreased caffeine sensitivity when the DHPR was placed in a preactivated state. These results provide the first example of how long-range allosterism between the DHPR and RyR1 moderates the activity of RyR1 and Ca2+ release in response to direct agonists such as caffeine.
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MATERIALS AND METHODS |
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Cell culture. The methods used to create the 1B5 cell line and derive the wild-type cell line from neonatal mice are described in detail elsewhere (6, 24). 1B5 cells were cultured on collagen-coated 10-cm tissue culture plates, and primary myoblasts were cultured on 10-cm plates coated with ECL (Upstate Biotechnology, Lake Placid, NY) at 37°C in a 5% CO2 incubator in low-glucose DMEM (growth medium) containing 20% fetal bovine serum, 100 U/ml penicillin, 100 µg/ml streptomycin, and an additional 2 mM L-glutamine (plus 20 nM basic fibroblast growth factor in case of wild-type cell line). After 48 h, the 1B5 cells were replated in 1) 96-well plates with ultrathin clear bottoms (Corning, Costar, NY) coated with Matrigel (BD Biosciences, Bradford, MA) for Ca2+ imaging experiments or 2) 10-cm tissue culture plates coated with Matrigel for crude SR membrane preparations. After
48 h, primary myoblasts were replated in 96-well plates with ultrathin clear bottoms coated with Matrigel for Ca2+ imaging experiments. When cells reached confluence of
70%, the growth medium was replaced with differentiation medium (containing 5% heat-inactivated horse serum instead of 20% fetal bovine serum in growth medium) and placed into an 18% CO2 incubator to induce differentiation. 1B5 cells (1B5 myotubes) were subjected to viral infection 56 days later.
Viral infection. The cDNAs encoding for wild-type RyR1, RyR2, and four RyR1-RyR2 chimeras (see Fig. 1) were packaged into HSV-1 amplicon virions, using the helper virus-free packaging system. The methods have been described in detail elsewhere (9, 49). Five to six days after differentiation was begun, 1B5 myotubes were infected with 1 ml of differentiation medium containing HSV-1 virions at 4 x 105 infectious units/ml (a multiplicity of infection of 4). This mixture was removed 2 h later and replaced with differentiation medium. The 1B5 myotubes were imaged or disrupted to prepare crude SR membrane preparations 36 h after viral infection.
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Ca2+ imaging. The 1B5 myotubes expressing either one of the wild-type RyR isoforms or one of the chimeras or the primary myotubes cultured on the 96-well plates were loaded with 5 µM fluo 4-AM (Molecular Probes, Eugene, OR) in an imaging buffer at 37°C for 30 min. The imaging buffer consisted of 125 mM NaCl, 5 mM KCl, 2 mM KH2PO4, 2 mM CaCl2, 25 mM HEPES, 6 mM glucose, 1.2 mM MgSO4, and 0.05% BSA (fraction V) at pH 7.4. Each well of the 96-well plates was then washed three times with 100 µl of the imaging buffer. The myotubes were transferred to an inverted stage microscope equipped with an Olympus Uaop/340 x40 oil-immersion objective (NA 1.35) (Nikon Diaphot300; Melville, NY). The microscope was modified to incorporate a pair of separate three-dimensional (3-D) micromanipulators on either side of the vertical post holding the condenser. The myotubes were imaged by using a PTI delta-RAM (Photon Technology International, Lawrenceville, NJ) as the light source with a 12-bit digital intensified charge-coupled device camera (Stanford Photonics, Stanford, CA). Fluo 4 in the myotubes was excited at 494 nm, and fluorescence emission was measured at 516 nm. Data were displayed and analyzed using QED Imaging software (v1.3; Pittsburgh, PA). For sensitivity, caffeine was dissolved in the imaging buffer and applications were performed via a perfusion system with a 16-channel perfusion pipette (AutoMate Scientific, Berkley, CA). The 3-D micromanipulators were used to place a pair of capillaries precisely into each well as 1) an inlet that allows delivery of the desired medium or 2) an outlet that allows suction through vacuum of the exceeding media, keeping the level of media in the well constant. The perfusion inlet was positioned about 1 mm above the imaged myotubes to allow a very efficient and rapid stimulus of the imaged area. When La3+ (0.1 mM) and Cd2+ (0.5 mM) or nifedipine (50 µM) were used to inhibit ion permeability through DHPR, they were added to the washing solution on myotubes for 2 min before caffeine challenges and were included in all caffeine and washing solutions after that time. For measurement of SR Ca2+ content of single myotubes, 10 µM cyclopiazonic acid (CPA) was manually applied. The SR Ca2+ contents were calculated in the following way: the maximal fluorescence after 10 µM CPA challenge (Fmax) minus the average baseline fluorescence for the 10 s immediately preceding the challenge (Frest) was then divided by Frest. Individual caffeine sensitivity curves were fitted via the Hill equation by using Origin 4.1.
Measurements of resting Ca2+ levels and membrane potentials. Double-barreled Ca2+-selective microelectrodes were prepared from piggyback borosilicate glass capillaries [PB15OF-4; outer/inner diameter (OD/ID) 1.5/0.8 mm; WPI, Salem, NJ]. In short, before being pulled, the capillaries were washed with HCl and distilled water and then dried at 150°C for 3 h. They were then pulled to a short taper with a total outside tip diameter of 0.6 µm, using a flaming brown micropipette puller (P-87; WPI). The 1.5-mm-OD tube was sealed by exposing the tip to dimethyl dichlorosilane, and then 24 h later the tip was backfilled with the calcium ionophore ETH 129 (Fluka, St. Louis, MO). The remainder of the barrel was backfilled with a solution of pCa 7 48 h afterward. Ca2+-selective microelectrodes were calibrated in a solution of known Ca2+ concentrations (pCa 38) at 24°C as described previously (19), but with the addition of 1 mM Mg2+ to each solution to mimic intracellular ionic conditions. They were calibrated individually, and only those that gave a Nernstian response between pCa 3 and 7 (29.5 mV per pCa unit at 24°C) were used experimentally. Individual Ca2+-selective microelectrodes were used only for a maximum of 1012 determinations of intracellular free Ca2+ concentration. The 0.8-mm-OD barrel was backfilled with 3 M KCl (tip resistance 1015 M) just before the determination was carried out. To prevent artifacts due to changes in the sensitivity of the Ca2+-selective microelectrode, microelectrodes were calibrated before and after the intracellular Ca2+ determinations.
Single myotubes were impaled with the Ca2+-selective microelectrodes while being observed through an inverted microscope (Axiovert 10; Thornwood, NJ) fitted with a x40 eyepiece and a x40 dry objective. The potential from the 3 M KCl barrel (Vm) and the potential from the Ca2+ barrel (VCae) were recorded via an amplifier (FD-223; WPI). The subtraction of Vm from VCae gives a differential signal (VCa) that represents the resting myoplasmic Ca2+ concentration. The potentials Vm and VCa were filtered at 200 Hz (LPF-30; WPI) to improve the signal-to-noise ratio. The recorded potentials were stored in a computer for future analysis.
Crude SR membrane preparation. Crude SR membrane preparations from the 1B5 myotubes expressing the RyR isoforms and the chimeras were prepared 36 h after viral infection. The 1B5 myotubes were collected by adding a harvest buffer (137 mM NaCl, 3 mM KCl, 8 mM Na2HPO4, 1.5 mM KH2PO4, and 0.6 mM EDTA at pH 7.2) from 10 to 15 plates (10 cm) and were centrifuged for 10 min at 250 g at 4°C. From this, samples were kept at 4°C during all following steps. The pellet was suspended with a buffer consisting of 250 mM sucrose, 10 mM HEPES, 1 mM EDTA, pH 7.4, and protease inhibitors (10 µg/ml leupeptin, 0.7 µg/ml pepstatin A, 5 µg/ml aprotinin, and 0.1 mM PMSF) and was then homogenized using a Polytron cell disrupter (Brinkmann Instruments, Westbury, NY). The homogenates were centrifuged for 20 min at 1,500 g, and the supernatants were recentrifuged for 1 h at 100,000 g. The pellet was finally suspended in a storage buffer consisting of 250 mM sucrose and 20 mM HEPES at pH 7.4, frozen in liquid N2, and stored at 80°C.
[3H]ryanodine binding assay. Crude SR membrane preparations (0.1 µg/µl) were incubated with 5 nM [3H]ryanodine (58 Ci/mmol; NEN) in a binding buffer (250 mM KCl, 20 mM HEPES, and 100 µM CaCl2 at pH 7.4) at 37°C for 3 h. Nonspecific binding was assessed by the presence of 1,000-fold unlabeled ryanodine and 1 mM EGTA. Separation of the bound from the free ligand was performed by rapid filtration through Whatman GF/B glass fiber filters using a Brandel cell harvester (Gaithersburg, MD). The filters were washed with three volumes of 5 ml of ice-cold buffer containing 20 mM Tris·HCl, 250 mM KCl, 15 mM NaCl, and 50 µM CaCl2 at pH 7.1 and placed into vials with 5 ml of scintillation cocktail (ReadySafe; Beckman Instruments, Fullerton, CA). The remaining [3H]ryanodine on the filters was quantified by a liquid scintillation spectrometry. Free Ca2+ concentrations in each binding assay were adjusted by adding EGTA or CaCl2 based on the calculations from the Bound and Determined program. Individual sensitivity curves were fitted with the Hill equation by using Origin 4.1.
Statistical analysis. Results are given as means ± SE, with the number of experiments reported. Significance of the differences was analyzed by the paired or unpaired t-test (GraphPad InStat, v. 2.04). Differences were considered to be significant when P < 0.05.
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RESULTS |
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Different sensitivities of wtRyRs and four chimeras to caffeine. Caffeine sensitivities of the individual 1B5 myotubes expressing wtRyRs and four chimeras were measured by imaging fluo 4 fluorescence (Fig. 2). 1B5 myotubes were sequentially challenged with increasing caffeine concentrations (0.140 mM). Each challenge lasted 30 s, with a rest interval between doses of 30 s. Figure 2A shows representative traces of the caffeine dose-response relationship from individual 1B5 myotubes. Individual dose-response curves were fitted to the Hill equation (Fig. 2B). EC50 values for wtRyR1 and wtRyR2 were 2.80 ± 0.12 mM (nH = 2.41 ± 0.15) and 0.84 ± 0.11 mM (nH = 2.07 ± 0.53), respectively, revealing that RyR2 is 3.3 times more sensitive to caffeine than RyR1 when tested in the context of 1B5 myotubes. EC50 values of the four chimeras tested were higher than that for wtRyR2 but less than that for wtRyR1, having a rank order of wtRyR2 < R9 < R4 < R10 = R16 < wtRyR1 (Table 1).
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In short, La3+ and Cd2+ increased the caffeine sensitivity of 1B5 myotubes expressing wtRyR1 but not wtRyR2, suggesting the possibility of long-range allosteric influences between the DHPR and RyR1 that normally engage a negative regulatory module within the Ca2+ release channel. Replacement of the corresponding regions of RyR2 with either the R4, R9, R10, or R16 regions of RyR1 has been shown to restore skeletal-type EC coupling (38), but interestingly, in the present study they did not reveal the locus of the negative module in wtRyR1 and failed to restore the inhibitory influence of the DHPR on caffeine-induced Ca2+ release in situ. In fact, rather than showing an increased sensitivity when the DHPR was placed into a preactivated state, there was a diminution of caffeine sensitivity of 1B5 myotubes expressing R4 and R9 under these conditions.
Effects of nifedipine on caffeine sensitivities of wtRyRs and four chimeras. The influence of La3+ and Cd2+ on the pattern of negative modulation of caffeine-induced Ca2+ release in 1B5 myotubes expressing wtRyRs and four chimeras extended our study to the organic DHPR blocker nifedipine (Fig. 4). Nifedipine is known as a specific inhibitor of DHPR (31) with no direct effect on RyR (50). The same experimental procedure utilized in the experiments shown in Fig. 2 was performed in the presence of external solutions containing 50 µM nifedipine. Figure 4A shows representative traces of the caffeine dose-response relationship from individual 1B5 myotubes in the presence of nifedipine. Figure 4B shows the number of responding 1B5 myotubes at increasing caffeine concentrations. Overall changes in the caffeine sensitivities seen when DHPR function was blocked by nifedipine were similar to those seen when the 1B5 myotubes were exposed to La3+ and Cd2+. The EC50 value for RyR1 was shifted sixfold to the left (from 2.80 ± 0.12 to 0.47 ± 0.02 mM) (Table 1). Similar to the results obtained with La3+ and Cd2+, there were no changes in the RyR2, R10, and R16, whereas the caffeine sensitivities of R4 and R9 were shifted to the right (from 1.27 ± 0.05 to 2.21 ± 0.11 mM and from 1.15 ± 0.03 to 2.28 ± 0.05 mM, respectively) (Table 1).
Effect of La3+ and Cd2+ or nifedipine on caffeine sensitivities of wild-type myotubes. To confirm increased caffeine sensitivity of 1B5 myotubes expressing wtRyR1 in the presence of La3+ and Cd2+ or nifedipine, cultured wild-type myotubes originating from the skeletal muscle of neonatal mice were incubated at 2 mM caffeine in either the absence or presence of La3+ and Cd2+, and the responses were normalized to the total number of wild-type myotubes (Fig. 5). Wild-type myotubes responded with a 2.8-fold enhanced frequency to 2 mM caffeine in the presence of La3+ and Cd2+ (from 32.83 ± 4.76 to 90.71 ± 3.57%; P < 0.05) (Fig. 5B), which is in accordance with the shift in caffeine sensitivity seen in 1B5 myotubes expressing wtRyR1 in the presence of La3+ and Cd2+. Interestingly, in the presence of La3+ and Cd2+, but not nifedipine, cells did not return to baseline after the 30-s washout period, making it impossible to obtain comparable caffeine dose-response curves using the standard protocol. Thus we were not able to calculate caffeine EC50 in wild-type myotubes in the presence of La3+ and Cd2+. It was possible to calculate caffeine EC50 in wild-type myotubes in the presence of 50 µM nifedipine. Figure 5C shows representative traces of the caffeine dose-response relationship from individual wild-type myotubes in the absence or presence of nifedipine. Figure 5D shows the number of responding wild-type myotubes at each caffeine concentration. Similar to the results obtained with La3+ and Cd2+ or nifedipine in 1B5 myotubes expressing wtRyR1, the caffeine EC50 for wild-type myotubes was shifted 2.6-fold to the left (from 3.29 ± 0.20 to 1.27 ± 0.35 mM; P < 0.05) and the Hill coefficient was significantly reduced (from 2.39 ± 0.24 to 1.23 ± 0.42; P < 0.05).
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SR Ca2+ contents, membrane potentials, and resting Ca2+ levels of single 1B5 myotubes expressing wtRyRs and four RyR1-RyR2 chimeras. To address possible mechanisms for the decreased caffeine sensitivities of R4 and R9 in the presence of La3+ and Cd2+ or nifedipine and the increased caffeine sensitivity of wtRyR1 in the same conditions, we measured SR Ca2+ contents, resting Ca2+ levels ([Ca2+]i), and resting membrane potentials of 1B5 myotubes expressing each construct. SR Ca2+ contents of 1B5 myotubes expressing wild-type RyRs and four chimeras were measured by depletion of Ca2+ from SR using 10 µM CPA, a SR Ca2+-ATPase inhibitor. Figure 6A shows representative traces, and the results are summarized as bar graphs in Fig. 6B. SR Ca2+ content of single 1B5 myotubes expressing RyR1 is higher than that of RyR2 (0.47 ± 0.07 vs. 0.83 ± 0.23, respectively, P < 0.05). Interestingly, R4 and R9 showed significantly lower SR Ca2+ contents compared with wtRyR1 (R4: 0.27 ± 0.03 and R9: 0.19 ± 0.03 vs. RyR1: 0.47 ± 0.07; P < 0.05). Membrane potentials and resting Ca2+ concentrations of 1B5 myotubes expressing each construct were examined by using double-barreled Ca2+-selective microelectrodes in the absence or presence of La3+ and Cd2+ or nifedipine (Table 2). Resting membrane potentials and [Ca2+]i of 1B5 myotubes expressing wtRyR1, wtRyR2, and four chimeras were not significantly different from each other, and the presence of La3+ and Cd2+ or nifedipine did not alter either [Ca2+]i or membrane potentials in any way (Table 2).
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[3H]ryanodine binding by wtRyRs and four chimeras. To confirm previous data for R4 and R9 chimeras with their wild-type RyRs (26, 34, 51, 52) and to extend analysis to chimera R16, we examined the modulation of high-affinity binding of [3H]ryanodine to crude SR membranes preparations by Ca2+. Despite the fact that 1B5 myotubes expressing R4, R10, and R16 restored Ca2+ release channel function in situ (Figs. 2, 3, 4), only R9 was capable of binding [3H]ryanodine with high affinity (data not shown). These results are consistent with those published by Nakai et al. (26) and indicate that some RyR1-RyR2 chimeras assume an unstable conformation, once extracted from their cellular context, that does not permit high-affinity binding of [3H]ryanodine.
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DISCUSSION |
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Why does the DHPR blockade alter the sensitivities of RyR1 R4 and R9 to caffeine? It has been suggested that there is a preactivated state of DHPR that can be induced by both membrane depolarization (7, 20) and binding of DHPR inhibitors such as dihydropyridines (50). Weigl et al. (50) proposed that the preactivated state of DHPR is sufficient to activate the RyR1 in human skeletal muscle cells. In addition, Kitamura et al. (16) reported that nifedipine causes a dose-dependent potentiation of K+ contractures in the presence of normal extracellular Ca2+ without any effect on the membrane potential in single frog skeletal muscle fibers. Therefore, it is possible that binding of La3+ and Cd2+ or nifedipine with the DHPR induces the preactivated state of DHPR. It follows that this conformational transition could influence a long-range allosteric interaction between the DHPR and RyR1 that enhances the sensitivity of wtRyR1 to caffeine. Unlike wtRyR1, wtRyR2 failed to alter its responses to any of the applied caffeine doses in the presence of La3+ and Cd2+ or nifedipine. The most likely explanation for the lack of influence of a DHPR block on RyR2 function is that these channels do not physically couple with the DHPR, as demonstrated by the fact that RyR2 does not support bidirectional signaling. Rather, RyR2 is activated by Ca2+-induced Ca2+ release that is initiated by Ca2+ entry through DHPR (3, 42).
The effects of La3+ and Cd2+ or nifedipine via the DHPR on caffeine sensitivity could also reflect the influence of several other factors, including a change in L-type Ca2+ current magnitude, [Ca2+]i, or membrane potential. Our hypothesis that the change is due to induction of a preactivated DHPR state by La3+ and Cd2+ or nifedipine would suggest a non-conducting conformational change of the DHPR, rather than an alteration of the actively open DHPR. However, without measuring DHPR current, we cannot definitively rule out a reduction in retrograde signaling, which would be revealed as a difference in Ca2+ current as the mechanism for this change in caffeine sensitivity. However, the changes in caffeine sensitivity of either wtRyR1 or the two chimeras induced by DHPR blockade cannot be explained by a simple difference in resting free Ca2+ or membrane potential among the constructs because there is no difference. Also, it is unlikely that the decreased caffeine sensitivities in the presence of La3+ and Cd2+ or nifedipine seen in cells expressing R4 and R9 can be explained simply on the basis of their observed decreased SR Ca2+ contents (Figs. 3, 4, and 6). It has been reported in rabbit and frog skeletal SR that the rate constant of Ca2+ release parallels the luminal Ca2+ concentration (5, 12, 14). At the single-channel level, increasing luminal Ca2+ favored channel activation by increasing the duration of open events of both cardiac and skeletal RyRs (12, 4345, 48). Therefore, even though it is possible that one of regulatory factors of the long-range allosteric interaction between the DHPR and RyR1 that could affect the sensitivity of wtRyR1 to caffeine could be luminal Ca2+, it is unlikely that the decreased caffeine sensitivities of R4 and R9 after DHPR blockade are secondary to their decreased SR Ca2+ contents, because caffeine sensitivities of R4 and R9 were different from that of wtRyR1 under control condition (Fig. 2B and Table 1). Therefore, RyR1 appears to act as a complex integrator of multiple regulatory stimuli including changes in the DHPR and endogenous effectors such as cytoplasmic and luminal Ca2+.
Taking all the results together, the negative allosteric interaction seen after treatment with the two DHPR blockers with the two RyR1-RyR2 chimeras R4 and R9 compared with the positive allosteric interaction seen with wtRyR1 demonstrates that, like the molecular interactions responsible for EC coupling, the mechanisms governing the long-range allosterism between the negative module in RyR1 and the DHPR are not simple but can be affected by other allosteric changes in other regions of RyR1.
Which other SR proteins could be involved with the functional module found in wtRyR1? Based on freeze-fracture replicas (10, 38), DHPRs form tetrad structures appearing to be connected with alternate feet in 1B5 myotubes expressing RyR1, suggesting a functional link between RyR1 and the DHPR. On the other hand, R10 and R16 form incomplete tetrads of DHPRs that exhibit an irregularly spaced pattern. However, R4 and R9 show the same tendency for tetrad formation as RyR1, suggesting that the R9 region has this important functional link with the DHPR and that this link could be one of the regulatory factors for a functional module in the R9 region of RyR1 (R9RyR1).
It has been reported that calmodulin (CaM) inhibits Ca2+-, caffeine-, and AMP-induced Ca2+ release from cardiac and skeletal muscle SR (11, 21, 22, 35). Plank et al. (35) found that the caffeine-induced Ca2+ release from actively loaded SR vesicles is reduced to 51% of the respective control value by 1 µM exogenous CaM. Furthermore, it was reported that a synthetic peptide containing residues 3,614 to 3,643 of the RyR1 was found to be capable of binding with high affinity either apo-CaM or Ca2+/CaM (23, 40), suggesting that CaM could participate as a negative module in R9RyR1.
In addition to orthograde regulation of RyR1, further work aimed at understanding the structural determinants within R9RyR1 that contribute to negative modulation should clarify whether initial activation and inactivation mechanisms of RyR1 and RyR2 during EC coupling differ. However, on the basis of present results with R4 and R9, the chimeric approach may not be a viable strategy given the confounding influence of long-range allosterism. A more direct approach using site-directed mutations or scrambling short sequences within R9RyR1 is likely to be more informative.
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ACKNOWLEDGMENTS |
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GRANTS
This work was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant P01-AR-44650 (to P. D. Allen and I. N. Pessah), Korean Ministry of Education Grant BK21 (to D. H. Kim), and the Postdoctoral Fellowship Program of the Korea Science and Engineering Foundation (to E. H. Lee).
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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