1Institute of Molecular Physiology and Genetics, Slovak Academy of Sciences, 833 34 Bratislava, Slovakia; and 2Department of Physiology, Texas Tech University Health Sciences Center, Lubbock, Texas 79430
Submitted 30 June 2003 ; accepted in final form 29 September 2003
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ABSTRACT |
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excitation-contraction coupling
Cardiac calcium channels inactivate by a calcium-dependent mechanism (40, 33), which is mediated by binding of Ca2+ ions to the two COOH-terminal binding sites on the calmodulin molecule prebound at the cytoplasmic side of the DHPR (41, 50). Therefore, in parallel with activation of calcium release, calcium current (ICa) becomes rapidly inactivated by the released calcium (1, 4, 16, 45, 46, 48, 58). Thus the calcium release-dependent inactivation (RDI) might be considered a convenient probe of calcium release because it senses the released Ca2+ ions directly in the dyadic junction (1, 14, 42, 45, 46). However, the difficulty in isolating RDI from the other two coexisting inactivation mechanisms of ICa inactivation, i.e., from the voltage-dependent and calcium current-dependent inactivation (17, 27, 32, 33, 40), has limited the use of RDI for quantitative assessment of calcium release mechanisms. Sham (46), using caffeine as an agent to deplete SR of calcium, visualized the calcium release-sensitive component of the calcium current by comparing calcium currents occurring in the presence of all three inactivation mechanisms and after RDI had been selectively removed by caffeine addition. By means of these intricate experiments, Sham (46) was able to show that calcium RDI results from local calcium release. For quantitative description of calcium release activation, however, this or analogous pharmacological approach would have serious limitations due to nonspecific effects of drugs and the complexity of experiments.
We have developed an alternative approach based on the known efficiency of tail calcium currents to induce Ca2+ release (5, 8, 10, 12, 14). We have varied the properties of the calcium current evoked by brief voltage prepulses and related them to the extent of RDI measured by the subsequent test pulse. This approach allowed us, for the first time, to induce RDI in the absence of other components of ICa inactivation, as well as to use it as a local, dyadic probe of calcium release activation. The relationship between Ca2+ influx and the ensuing RDI was analyzed on the basis of the law of mass action to show that the potency of a given calcium influx to induce calcium release, and hence RDI, was dependent on previous calcium influx. Increased probability of RyR activation by prior elevations of local Ca2+ was confirmed at the level of single RyRs in planar lipid bilayers by using brief, photolytically generated Ca2+ elevations under ionic conditions similar to those of the intracellular environment. Altogether, these findings suggest that the probability of calcium release activation by a single DHPR opening can be modulated by the recent history of calcium influx within the dyad.
Part of this work has been published in abstract form (Ref. 34).
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MATERIALS AND METHODS |
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Calcium current measurements. The L-type ICa were recorded from single myocytes by using the standard whole cell patch-clamp technique (28). The external solution contained (in mM) 135 NaCl, 5.4 CsCl, 10 HEPES, 5 MgCl2, 0.33 NaH2PO4, and 1 CaCl2, pH 7.3. Patch pipettes (12 M) were filled with the internal solution containing (in mM) 135 CsCH3SO3, 10 CsCl, 10 HEPES, 1 EGTA, 3 MgSO4, and 3 ATPNa2, pH 7.3. When necessary, sodium channels were inhibited with 20 µM tetrodotoxin (TTX) in the external solution and by a holding potential of 50 mV. In experiments involving hyperpolarizing stimuli, the external solution with Na+ ions replaced by Cs+ ions was used to minimize sodium channel currents on subsequent depolarization. Potassium currents were suppressed by replacement of K+ with Cs+ in all solutions. Calcium channels were kept in a phosphorylated state by adding 50 µM cAMP to the internal solution and 10 µM isobutylmethylxanthine (IBMX, a membrane-permeant phosphodiesterase inhibitor) to the external solution. Calcium currents could not be further stimulated by addition of 0.1 µM isoprenaline and were completely blocked by external Cd2+ or DHP antagonists. IBMX, ATP, and cAMP were from Sigma (St. Louis, MO). TTX was from Alomone Labs (Jerusalem, Israel), and ryanodine was from Calbiochem (La Jolla, CA). All other chemicals were of analytical grade. Whole cell currents measured with the Axopatch 200B (Axon Instruments, Union City, CA) patch-clamp amplifier were low-pass filtered at 25 kHz and digitized at 510 kHz by a Labmaster analog-to-digital board (Scientific Solutions) using pCLAMP (ver. 5.5.1, Axon Instruments) implemented on an IBM-AT type computer. Series resistance was electronically compensated by 5085%. Capacitance charging current was canceled electronically. Where indicated, online subtraction procedure was used additionally to cancel the remaining linear currents. Experiments were carried out at room temperature.
Inactivation of calcium current was probed using a constant test pulse (70 ms, 0 mV) preceded by very brief (5 ms) prepulses varying in either prepulse potential or tail potential (i.e., the potential following the prepulse). Individual protocols are described in detail in RESULTS.
Bilayer experiments. Heavy SR microsomes were isolated from canine left ventricles by differential centrifugation as described previously (60). Single SR Ca2+ release channels were reconstituted by fusing heavy SR microsomes into planar lipid bilayers as described previously (24, 61). Single-channel currents through RyRs were recorded in 400 mM CsCH3SO3, 10 mM Cs-HEPES, and 1 mM glutathione, pH 7.4. The cytoplasmic (cis) solution contained additionally 3 mM MgATP, and the luminal (trans) solution contained 1 mM CaCl2 (Orion). Single-channel currents were measured at +40 mV using Axopatch 200A (Axon Instruments), filtered at 5 kHz, digitized at 25 kHz, and acquired using Digidata 1200A and pCLAMP (ver. 8.0, Axon Instruments). All chemicals were from Sigma if not stated otherwise. Fast changes of the Ca2+ concentration in the microenvironment of the reconstituted channel were performed by flash photolysis of caged calcium as described previously (24, 26, 61), except that the Ca2+ cage compound NP-EGTA (Molecular Probes, Eugene, OR), which is highly selective for Ca2+ over Mg2+ (19), was used instead of DM-nitrophen. After channel incorporation, NP-EGTA (a final concentration of 3 mM) containing the appropriate amount of CaCl2 (Orion) to attain a free Ca2+ of 0.30 µM was added to the cytoplasmic (cis) side of the channel. Intense (2 mJ) and brief (9 ns) ultraviolet laser flashes produced by a pulsed, frequency-tripled Nd:YAG laser (Spectra-Physics, Mountain View, CA) were applied through a fused silica fiber optics (600-µm diameter) positioned in front of the bilayer (100-µm diameter) to illuminate the whole volume between the fiber optics and the bilayer. The concentration of steady-state free Ca2+ in the cis chamber was determined with a Ca2+-selective minielectrode (26). The local Ca2+ changes near the bilayer were calibrated by transforming the bilayer aperture into a Ca2+ electrode, using Ca2+ ionophore resin (ETH 129; Fluka, Switzerland) as described previously (26, 61). The amplitude and time course of free Ca2+ signals in response to the photolyzing laser pulses were computed in Mathematica (ver. 4.2, Wolfram Research, Champaign, IL) from the values of steady-state Ca2+ before and after the flash and from the concentration of total NP-EGTA (61), using published kinetic constants (18, 20). The calcium affinity of ATP was corrected for ionic strength and pH using the MaxChelator program (7, ver. 2.40; http://www.stanford.edu/~cpatton).
Data analysis. Experimental records were analyzed using Origin (ver. 7.0, OriginLab) on a PC computer. The data are reported as means ± SE.
The fraction of the test pulse ICa inactivated by the prepulse, Fpi, was estimated as
![]() | (1) |
The fraction of ICa inactivated during the test pulse, Fti, was estimated as
![]() | (2) |
Dependence of the Ca2+ influx on the prepulse voltage. The amount of Ca2+ ions, mCa, entering via Ca2+ channels was calculated as QCa/(2F), where F is the Faraday constant. QCa was determined by integrating the inward ICa and separated into the prepulse and tail components. The amount of Ca2+ ions injected into the cell during the prepulse (mCa,p) was estimated using integration of the ICa during the depolarization. The amount of Ca2+ ions injected during the tail current (mCa,t) was estimated by integrating the tail calcium currents relative to the value observed 5 ms after hyperpolarization so that Na+/Ca2+ exchange current did not contribute substantially to the measured mCa,t.
The experimentally determined voltage dependence of mCa,p was fitted by the functions
![]() | (3a) |
![]() | (3b) |
The voltage dependence of mCa,t was fitted by the function
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Contribution of prepulse and tail currents to inactivation of ICa. Both calcium charge injected during the brief prepulse (mCa,p) and during the tail (mCa,t) may trigger calcium release and, hence, inactivation of the test ICa, perhaps with different potency because of the differences in kinetics and single-channel current amplitude of calcium channels at different voltages. To estimate the relative contribution of the prepulse and the tail current to inactivation of the test pulse peak ICa, we have used short prepulses of variable amplitude.
ICa inactivation induced by SR calcium release should depend on the extent of activation of RyRs, which in turn increases with both the amplitude and the duration of the activating calcium stimulus (61). Under such conditions, the extent of RyR activation can be considered as dose dependent on the time integral of the Ca2+ stimulus (3, 35), in our case, on the ICa integral. Therefore, we have assumed that the fraction (Fpi) of ICa inactivated by Ca2+ release is dose dependent on the effective value of the Ca2+ influx sensed by the RyRs (), which is a fraction of the measured calcium influx
![]() | (5) |
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The fraction of the test pulse ICa inactivated by the prepulse (Fpi) then can be expressed by the function
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Because the parameters kp, kt, and in Eq. 7 are mutually dependent and because none of them can be independently estimated, we introduced the apparent half-effective dose
and the relative potency of prepulse calcium influx
= kp/kt, so that the dependence of Fpi on the amount of Ca2+ ions injected into the cell during the prepulse and during the tail current can be described by the formula
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The relative potency of the calcium influx during the prepulse was estimated by fitting the fraction of inactivated calcium current by Eq. 8. A value of = 1 means that calcium ions injected by the prepulse and by the tail current have equal potency to induce inactivation of ICa, and values of
> 1 and
< 1 mean that calcium influx is more effective when occurring during the prepulse and during the tail, respectively. The apparent half-effective dose for tail calcium influx is therefore equal to M50, and that for prepulse calcium influx is equal to M50/
.
For the purpose of visually comparing the measured data with the theoretical predictions, the values of mCa,p and mCa,t in the voltage range of 50 to +60 mV were calculated from the parameters obtained from the best fits of mCa,p and mCa,t by Eq. 3 and Eq. 4, respectively, and substituted into Eq. 8 to yield the theoretical values of Fpi. The theoretical dependence of Fpi on mCa,p only, or on mCa,t only, was calculated using Eq. 8 by setting Ca2+ influx by the other pathway to zero.
Contribution of tail-current calcium channel reopenings to inactivation of ICa. Calcium ions triggering Ca2+ release enter the cell in quanta determined by the duration and amplitude of the single-channel openings. The single-channel amplitude of the calcium current, iCa, depends linearly on the membrane potential (23, 55). Duration of calcium channel openings has exponential distribution with mean open time independent of membrane potential (31, 43). Within the short time interval during which the tail calcium current flows, the channel may reopen only few times before it finally deactivates. Because of the exponential distribution of the mean open time, not every opening during the tail may be long enough to provide sufficient amount of Ca2+ ions to activate calcium release. However, if the activated channel reopens, its potency to induce calcium release might be substantially increased because of the pre-elevated free-Ca2+ concentration in the junctional space. We have attempted to quantify the contribution of calcium channel reopenings to induction of ICa inactivation by analyzing the fraction of inactivated calcium current in experiments, in which a constant prepulse was followed by a variable tail potential.
The number of openings depends on the tail potential, Vt. At very negative voltages, only those channels that were open at the end of the prepulse will contribute to the tail current. Therefore, at very negative tail potentials, the amount of calcium ions entering the cell during the tail is equivalent to the amount entered during the first openings, mCa,1, which is
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The total amount of ions, mCa,t, is thus decomposed into the contribution of mCa,1 and mCa,r
![]() | (11) |
To analyze whether the first and subsequent openings during the tail differ in their ability to inactivate ICa, the quantities mCa,1 and nr had to be extracted from mCa,t. Because to is voltage independent and iCa(Vtp) is linear (23, 55), the voltage dependence of mCa,1(Vtp) (see Eq. 9) is also linear and can be expressed as
![]() | (12) |
![]() | (13) |
![]() | (14) |
For the purpose of visually comparing the measured data with the theoretical predictions, the values of mCa,1 and mCa,r obtained using the parameters k, nmax, V, and S from the fit of the experimental data from Eqs. 1113 were substituted into Eq. 14 to yield the theoretical values of Fpi. The theoretical dependence of Fpi on mCa,1 only, or on mCa,r only, was calculated using Eq. 14 by setting Ca2+ influx by the other pathway to zero.
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RESULTS |
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The slow inactivation that dominated ICa decay in cells that were depleted of releasable calcium suggests that the effects of the mechanisms responsible for slow inactivation could be negligible if very brief prepulses were used. Indeed, as illustrated in Fig. 1, B and D, a brief prepulse to a positive potential induced pronounced suppression of the test ICa only in cells showing the fast, calcium release-dependent component of inactivation. On average, Fpi induced by a prepulse to 0 mV was 0.48 ± 0.05 in 6 cells with RDI and 0.06 ± 0.02 in 4 cells without RDI. Such a dramatic difference in the effect of the short prepulse could not have been due to a difference in peak calcium current density between these two groups, which was not found to be statistically significant (P > 0.05).
These experiments revealed that brief prepulses represent a potential tool for selective induction of the calcium release-dependent ICa inactivation without the confounding contributions of the calcium current- and the voltage-dependent mechanisms inherent to inactivation evoked by prolonged prepulses. We next explored the use of brief prepulses, using the calcium release-dependent inactivation as a probe of calcium release activation.
Dependence of ICa inactivation on the potential of the brief prepulse. To investigate the fast ICa inactivation induced by brief depolarizations, we applied 5-ms prepulses from a holding potential of 50 mV to variable potentials (20 to +60 mV). After a 10-ms interval at 50 mV, the prepulses were followed by a 70-ms test pulse to 0 mV. A typical set of currents in response to this stimulation protocol is shown in Fig. 2A. Increasing the amplitude of the prepulse led to a progressive depression of both the peak amplitude and the inactivation rate of the test calcium current. It is notable that although the test ICa after a prepulse to +60 mV was suppressed only to 40% of its original peak amplitude, its decay completely lacked the fast inactivation component. In the absence of prepulse, however, calcium currents with peak amplitude of about 40% of maximum peak ICa underwent fast and pronounced inactivation (see traces at 20 and +30 mV in Fig. 1C). This means that the calcium current remaining after partial inactivation has a dramatically decreased ability to trigger calcium release-dependent inactivation, even if it retains relatively large amplitude. In the case of the calcium current-dependent inactivation, the rate of fast inactivation was not decreased by 50% preinactivation (52).
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We compared the fraction, Fpi, of the peak test ICa inactivated by the prepulse (Eq. 1) with the fraction, Fti, of the respective ICa that inactivated during the test pulse (Eq. 2). As shown in Fig. 2B, the larger was the fraction of the peak test calcium current inactivated by the prepulse; the smaller was the fraction of the test calcium current inactivated during the test pulse, resulting in a close negative correlation between the respective inactivation fractions for all prepulse potentials (R = 0.88, P < 0.0001). In other words, the fraction of channels not subjected to calcium release-dependent inactivation by the prepulse underwent RDI during the test pulse. Such a relationship suggests that calcium channels undergoing inactivation by the prepulse and during the test pulse are recruited from the same pool of channels, namely, the pool of channels that face calcium ions released from the SR. This observation implies that prepulse-induced RDI affected only calcium channels in those dyads in which calcium release was activated by the prepulse.
Prepulse- vs. tail-induced ICa inactivation. The prepulse as a trigger of calcium release can be thought of as a composite stimulus that can be divided into two parts, the calcium influx during the prepulse current and the Ca2+ influx during the tail current. We attempted to compare the potency of these two components in triggering calcium release and, thus, RDI. The amount of calcium flowing into the cell during the prepulse (mCa,p) and that during the tail current (mCa,t) was estimated by integrating the respective segments of the calcium currents. The amount of calcium ions injected into the cell during the tail current (Fig. 3A, ) increased with the amplitude of the prepulse sigmoidally, whereas mCa,p had bell-shaped voltage dependence (Fig. 3A,
). The voltage dependences of mCa,p and mCa,t were approximated by Eqs. 3 and 4, respectively. The resulting parameters of the best fits are given in Table 1, and the theoretical curves are overlaid with the data in Fig. 3A. It can be seen that mCa,t saturated at prepulses above +20 mV, in accordance with the voltage dependence of ICa activation (43).
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The fraction, Fpi, of the test pulse peak ICa amplitude inactivated by the prepulse (Fig. 3B, ) increased sigmoidally with the prepulse potential. It is obvious that the voltage dependence of mCa,t but not of mCa,p is similar to the voltage dependence of Fpi. Therefore, it can be speculated that the factual trigger of the calcium release-dependent inactivation of the test ICa might be constrained to the tail calcium current of the prepulse. Plotting the fraction of the test ICa inactivated by prepulses to different potentials against the amount of Ca2+ charge injected during the prepulse and tail currents (Fig. 3, C and D) revealed that although the relationship between Fpi and mCa,p was quite complex (Fig. 3C), Fpi increased sigmoidally with increasing mCa,t (Fig. 3D). This analysis suggests again that the tail current following the prepulse might be responsible for inactivation of the test ICa. As in the absence of the calcium release-dependent inactivation the tail current did not induce significant inactivation of ICa (Fig. 1B), it appears that release of Ca2+ evoked by the tail current was the immediate cause of ICa inactivation. However, in the voltage range between 30 and +20 mV, the values of mCa,p and mCa,t are of comparable size (Fig. 3A). Therefore, we tested the hypothesis that mCa,p contributes to prepulse-induced inactivation, albeit with a different potency (Eq. 6). As outlined in MATERIALS AND METHODS, we assumed that the extent of calcium release, and consequently the fraction Fpi of ICa inactivated by Ca2+ release, are dose-dependently proportional to the effective calcium influx sensed by the RyRs and that the calcium influxes mCa,p and mCa,t contribute differently to the effective calcium influx. The potency of mCa,p to induce a given Fpi is then a constant fraction of the potency of mCa,t, i.e., the relative potency of mCa,p with respect to mCa,t can be expressed by the factor
(Eq. 8). Fitting Eq. 8 to the values of Fpi provided the optimal value of
substantially less than 1 (Table 2). It means that the potency of mCa,p is substantially less than that of mCa,t. Figure 3, BD, show the theoretical relationships between Fpi, mCa,p, mCa,t, and membrane potential, calculated from the voltage dependencies of mCa,p and mCa,t (Eqs. 3 and 4, Table 1) and from the dependencies of Fpi on mCa,p and mCa,t (Eq. 8, Table 2) overlaid on the experimental data measured at different prepulse potentials. It can be seen that the theoretical curves reproduce well the observed relationships.
Although it is not possible to expose the cells to the prepulse and tail Ca2+ influx in separation from each other, the theoretical effects of these two modes of calcium influx on Fpi can be calculated from their potencies (see dotted and dashed curves in Fig. 3, BD). The difference between the effect of the prepulse and the tail Ca2+ influx separated from each other is then even more pronounced than the difference in the potencies because in contrast to mCa,t, the value of mCa,p is below the apparent half-effective dose at all potentials.
Dependence of ICa inactivation on the tail potential. The above findings might be interpreted as if it were the increased amplitude of the single-channel current during the tail that increased the potency of calcium influx to evoke inactivation of ICa. However, variation in the tail potential invokes variation in the tail current kinetics as well. It follows from the recent understanding of calcium channel function that both the magnitude of the single-channel current and the rate of the current deactivation increase with hyperpolarizing membrane potential. Then, at less hyperpolarized tail voltages, the single-channel current amplitude is lower due to reduced driving force on calcium ions, but the current deactivates more slowly, due to increased probability of channel reopenings, than at more hyperpolarized voltages. From the point of release-dependent inactivation, we may speculate that if the amplitude of the single-channel openings is more important for triggering calcium release, RDI should increase with the tail hyperpolarization. Conversely, if reopenings are more important, then RDI should decrease with the tail hyperpolarization.
In this series of experiments, the extent of calcium channel activation at the end of the prepulse was kept constant by using constant prepulse duration, and Ca2+ influx during the prepulse was minimized by depolarization to the reversal potential. A very short prepulse (3 ms, +60 mV) was followed by a 10-ms interval at a variable tail potential (Vt = 120 to 40 mV), then by a 20-ms interpulse interval at the holding potential of 50 mV, and finally by the test pulse (70 ms, 0 mV). To avoid activation of sodium currents by repolarization steps from the very negative tail potentials to the holding potential and subsequent activation of the Na+/Ca2+ exchange, we used Na+-free extracellular solutions. A typical experiment is presented in Fig. 4A. Suppression of the peak test ICa and of the fast inactivation of calcium current during the test pulse increased with increasing (less negative) tail potentials (Fig. 4B) in parallel with increasing amount of calcium injected into the cell by the tail current (Fig. 4C, half-open symbols). As in the previous series of experiments, Fpi had again a tendency to saturate at a level lower than unity (Fig. 4D). Incomplete suppression of the peak test ICa suggests that there was a population of calcium channels that were not subjected to inactivation by Ca2+ released from the SR. Similar to experiments with a variable prepulse amplitude, there was again a close negative correlation between Fpi and Fti (data not shown), suggesting once more that the prepulse inactivates the same pool of channels as the test pulse.
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The theoretically expected effects of the pure Ca2+ influx during the first openings and the reopenings, if they could occur independent of each other, on Fpi were estimated from their potencies and from Eq. 14 (see dotted and dashed curves in Fig. 4, B, E, and F, and the corresponding legend). The difference in their effect arises from the fact that mCa,1, in contrast to mCa,r, is below its apparent half-effective influx (M50/) at all studied potentials. The first opening and the reopenings contributed equally to the calcium influx at 60 mV, but due to their different potencies, the contribution of the two calcium influx modes to Fpi would be equal at 90 mV. Therefore, at repolarization to the holding potential of 50 mV, the RDI of ICa induced by the tail current following the prepulse was brought about mainly by the reopenings during the tail current.
In both types of experiments presented above (Figs. 3 and 4), Ca2+ influx was more effective when occurring later. That is, the tail current was more effective than the prepulse current, and the reopenings were more effective than the first openings. The higher potency of Ca2+ influx occurring shortly after previous Ca2+ influx suggests that the potency of Ca2+ influx to induce calcium release depends on the recent history of Ca2+ influx rather than on the overall Ca influx.
Activation of RyRs by paired brief calcium elevations. To directly examine whether the variable potency of a given amount of Ca2+ ions to activate calcium release is apparent at the level of single RyR channels, we have fused cardiac SR microsomes into lipid bilayers and recorded the activity of single RyRs with Cs+ as the charge carrier. Cytosolic concentrations of free Mg2+ and total ATP in cardiac cells are in the range of 0.51.2 and 35 mM, respectively, and the free luminal Ca2+ concentration is close to 1 mM (6). Therefore, to imitate the conditions present in the cell, the cytoplasmic solution contained 3 mM total ATP (free Mg2+, 0.6 mM) and the luminal solution contained 1 mM Ca2+. To mimic two consecutive openings of a calcium channel, we applied two consecutive ultraviolet laser pulses at a 100-ms interval to generate rapid Ca2+ changes by photolyzing the calcium cage compound NP-EGTA. The amount of Ca2+ liberated by the two flashes was identical. Subtle differences between the shape of the first and the second stimulus were present because of the differences in the saturation of the Ca2+ buffer, NP-EGTA, at the time of the stimulus. The first flash induced a rapid and brief (full duration at half amplitude, FDHA = 0.14 ms) Ca2+ elevation from
0.3 µM to
65 µM, followed by a sustained Ca2+ elevation to
0.4 µM; the second flash produced a slightly longer Ca2+ signal (FDHA = 0.19 ms) that again reached a peak amplitude of
65 µM (Fig. 5A). The probability of RyR openings by the first Ca2+ stimulus was low (P = 0.03; Fig. 5C), whereas the second Ca2+ stimulus triggered many more openings (P = 0.1; Fig. 5, B and C). Based on results obtained in four individual experiments, the probability of RyR activation during the second stimulus was four times larger than that during the first Ca2+ stimulus (0.16 ± 0.03 vs. 0.04 ± 0.01, P < 0.05). The changes in probability of RyR activation (300%) were much more pronounced that the changes in basal Ca2+ concentration (
40%) and in the duration of the stimulus (
30%). These results support the notion that the probability of RyR activation by the same amount of liberated Ca2+ steeply increases with the Ca2+ preceding the calcium stimulus and/or with the duration of the stimulus.
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DISCUSSION |
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It is well known that the fast inactivation of ICa depends on the release of Ca2+ ions from the sarcoplasmic reticulum (1, 4, 45, 46, 48, 58). However, quantitative determinations of the extent of RDI have been limited due to lack of straightforward experimental approaches. RDI was isolated pharmacologically by comparing currents in the presence and absence of SR Ca2+ release (employing caffeine to deplete the SR of calcium) under conditions that supported also current- and voltage-dependent inactivation of ICa (46). However, besides depleting the SR of calcium, caffeine has several additional effects on the calcium current (58). In addition, the influence of RDI on the kinetics of current- and voltage-dependent inactivation mechanisms, which is manifested as a prominent increase in the rate of the slow calcium current-, and voltage-dependent inactivation observed in the absence of RDI (Fig. 1, see also Fig. 2 in Ref. 46) might present an additional limitation of the pharmacological approach. Employing brief prepulses as stimuli inducing RDI without contribution of other inactivation mechanisms (Fig. 1) enabled us to determine the extent of RDI directly.
Relationships between RDI and calcium release activation. Our results and analysis indicate (Figs. 1 and 2) that the calcium release-dependent inactivation of ICa results from exposure of only a fraction of calcium channels to Ca2+ ions released from the nearby SR rather than from exposure of all the calcium channels to globally increased cytoplasmic calcium. This is inferred from the finding that the fraction of the test pulse calcium current that underwent fast inactivation in the absence of a prepulse could be inactivated by a brief prepulse (Fig. 2B), a stimulus specific for RDI. RDI affected 75% of the peak ICa amplitude (Figs. 2, 3, 4), suggesting that ICa activated only about 75% of release units under our conditions. Our estimates of excitation-contraction (E-C) coupling efficacy made by measuring RDI are consistent with those based on fluorescence imaging of local calcium signals (i.e., calcium "sparks" and "spikes"). Thus using rapid two-dimensional confocal imaging and fluo 3, Cleemann et al. (14) found that depolarization to 0 mV leads to activation of about 70% of release units in rat ventricular myocytes. By measuring spatially resolved fluorescence spikes with Oregon Green BAPTA-5N, Song et al. (51) estimated that 6090% of all available dyads become activated during E-C coupling under fully phosphorylated conditions in the same voltage range. In addition, the dependencies of Fpi (this work) and calcium release (8) on the tail potential are very similar. Thus the principles of local calcium signaling that apply to regulation of SR Ca release (12, 25, 30, 38, 53, 56) also manifest themselves in features of RDI.
The direct proportionality between the fraction of activated calcium release units and the extent of RDI, ensuing from the local control of RDI, enabled us to derive the relationships among calcium influx, activation of calcium release, and induction of RDI (Eqs. 8 and 14). Varying single-channel properties during the prepulse and tail current portions of the calcium release-inducing stimulus enabled us to show for the first time that different modes of calcium influx contributed differently to the effective calcium trigger signal. In the two-pulse experiments with 50 mV tail potential, changing the amplitude of the prepulse varied the relative proportion of calcium ions entering the cell during the prepulse and during the tail current. Analysis of the relationship between ICa inactivation and Ca2+ influx (Fig. 3), based on the principle of mass action (Eq. 8), showed that the same amount of calcium influx was much more effective in triggering calcium release when entering during the tail (mCa,t) than when entering during the prepulse (mCa,p; compare the dashed and dotted curves in Fig. 3B). In other words, the fraction of mCa,t that contributed to the effective calcium influx was larger than the fraction of mCa,p. In agreement with this analysis, maximal suppression of the test ICa by RDI was observed at prepulses to the reversal potential at which all Ca2+ ions entered the cell during the tail current, and not around prepulse voltages at which the total Ca2+ influx was maximal. Similar observations are usually interpreted as showing that the probability of triggering calcium release increases with the single-channel amplitude of the calcium current (12, 44, 51). It should be noted, however, that the dependence of the extent of RDI on the prepulse tail potential appeared to be at variance with such conclusion. When the DHPR channels were preactivated in the absence of calcium influx (ICa near the reversal potential), and their deactivation rate and single-channel amplitude were varied by changing the tail potential, the potency of calcium influx to activate calcium release (i.e., the extent of RDI observed for the same amount of calcium influx) appeared to increase with decreasing single-channel amplitude. This apparent paradox was resolved by realizing that tail calcium influx comprises of two phenomena with different voltage dependences: the single-channel current amplitude and the number of channel reopenings. Because of the much higher potency of calcium channel reopenings to activate calcium release, they contributed more substantially to the effective Ca2+ influx than did the first openings at all tail potentials positive to 90 mV (compare the dashed and dotted curves in Fig. 4B). These observations strongly indicate that the probability that a solitary DHPR opening will activate its neighboring RyRs is less than one even at the most negative voltages (120 mV).
The difference in efficiency between single and multiple openings can be understood on the grounds of our previous findings (59, 61) that significant levels of RyR activation may be reached only with sufficiently prolonged exposure of RyRs to elevated Ca2+. The effective exposure time for any given concentration of calcium is determined by the kinetics of RyR activation. In addition, preceding influx of Ca2+ that failed to activate release may result in Ca2+ accumulation in the junctional space. The increased level of Ca2+ may potentiate the effects of subsequent influx of Ca2+ by partial occupation of the mobile and immobile calcium buffers (39) that reduces calcium-buffering capacity of the junctional space. In effect, less Ca2+ from the subsequent Ca2+ influx is consumed by calcium buffers, and a larger amount of free Ca2+ is left available for activation of RyRs. Both these nonlinear processes are effective only locally and briefly due to the fast chemistry of Ca2+ ions and the quantal character of calcium influx from the point source(s). Pre-elevated junctional Ca2+ may also have direct potentiating effect on RyR activation by increasing the fraction of Ca2+-bound RyR states at the expense of Mg2+-bound states (36, 57). Local changes of dyadic Ca2+ on a longer time scale were speculated to contribute to the increase of the frequency of spontaneous calcium sparks upon inhibition of the forward mode sodium-calcium exchange (22) and to potentiation of calcium current-induced calcium release by the calcium influx by the reverse-mode Na+/Ca2+ exchange (37).
In agreement with these notions, we have directly demonstrated in single-channel experiments that the probability of activation of RyRs by a given Ca2+ influx increases with repeated stimulus application. The observed increase in open probability could have been due to changes in the Ca2+ concentration preceding the stimulus and/or due to prolongation of the stimulus, both due to the partial saturation of the calcium buffer. Because of the long time constant of diffusion in the 0.1-mm3 volume exposed to flash photolysis in our experimental setting (10 s; Ref. 26), the changes in buffer saturation fully persisted during the 100-ms interval between the two stimuli. The volume of the dyadic gap is only 0.001 µm3 and, therefore, the changes in buffer saturation due to local calcium influx are expected to persist only on the millisecond time scale (49).
Implications for excitation-contraction coupling. The voltage dependence of Fpi on the calcium influx through an isolated DHPR opening enables direct estimation of the fidelity of coupling for a given number of DHPR channels in the dyad and known DHPR open probability (Po,DHPR). Previous estimates, in addition to experimental measurements, required also specifying the amplitude of the DHPR single-channel current and DHPR open time, as well as of the amplitude and duration of the calcium influx during a single spark (62), parameters that can be inferred only indirectly. Assuming 20 DHPRs per calcium release unit (6, 21) and a maximum Po,DHPR of 5% (43), our data (Table 4) suggest a probability of Ca2+ release activation by a solitary DHPR opening (coupling fidelity; Refs. 51, 55, 62) of 35% at 120 mV, 15% at 40 mV, 6% at 0 mV, and <3% at +20 mV. These values are in good accord with the previous estimates of coupling fidelity (15 and 2% at 40 and 0 mV; Ref. 62). Extrapolation to calcium currents in the presence of the calcium agonist FPL 64176 by assuming a 20-fold increase of mCa per opening (43, 51) and Po,DHPR = 0.8 (51) gives an estimate of coupling fidelity of 0.65 at 0 mV in agreement with direct measurements of coupling fidelity in the presence of FPL 64176 (55). The good correspondence between our estimates for two widely different conditions and the corresponding estimates by two different methods (51, 55) suggests that the relationship between Ca2+ influx and calcium release activation for solitary DHPR openings (dotted line in Fig. 4E) might be quite general.
In summary, we have shown that tail calcium currents following repolarization from short depolarizing pulses to positive membrane potentials are better triggers of calcium release than calcium currents carrying a similar charge during the depolarizing pulses themselves. Moreover, the potency of calcium current to trigger calcium release is higher at less negative repolarization potentials. This higher potency might result from a combination of the potentiating effects of futile channel openings that failed to activate RyR channels but increased the basal Ca2+ level during depolarization and of the high temporal synchronization of the calcium influx through preactivated calcium channels on repolarization.
The physiological consequences of these findings may become comprehensible when brief stimuli in our experiments are related to the shape of the action potential: the rising phase and the peak of the action potential resembles the brief depolarizing prepulse, whereas the early repolarization resembles the end of the prepulse. Our results then suggest that the efficiency of calcium influx to trigger calcium release should achieve its maximum during the early repolarization phase of the action potential. In this way, our findings explain why the peak of calcium release occurs within a few milliseconds after the peak of the action potential (9, 29, 42).
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ACKNOWLEDGMENTS |
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GRANTS
The research of A. Zahradníková was supported in part by a Howard Hughes Medical Institute International Scholar's Award. This work was supported by VEGA Grant 2/1082/21 (to A. Zahradníková) and Fogarty International Research Collaboration Award 1 R03-TW-05543-01 (to S. Györke).
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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.
* A. Zahradníková and Z. Kubalová contributed equally to this work.
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REFERENCES |
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---|
2. Adachi-Akahane S, Cleemann L, and Morad M. BAY K 8644 modifies Ca2+ cross signaling between DHP and ryanodine receptors in rat ventricular myocytes. Am J Physiol Heart Circ Physiol 276: H1178H1189, 1999.
3. Ashley CC and Moisescu DG. Model for the action of Ca2+ in muscle. Nat New Biol 237: 208211, 1972.[ISI][Medline]
4. Balke CW and Wier WG. Ryanodine does not affect calcium current in guinea pig ventricular myocytes in which Ca2+ is buffered. Circ Res 68: 897902, 1991.[Abstract]
5. Barcenas-Ruiz L and Wier WG. Voltage dependence of intracellular [Ca2+]i transients in guinea pig ventricular myocytes. Circ Res 61: 148154, 1987.[Abstract]
6. Bers DM. Excitation-Contraction Coupling and Cardiac Contractile Force. Boston: Kluwer, 2001.
7. Bers DM, Patton CW, and Nuccitelli R. A practical guide to the preparation of Ca2+ buffers. Methods Cell Biol 40: 329, 1994.[ISI][Medline]
8. Beuckelmann DJ and Wier WG. Mechanism of release of calcium from sarcoplasmic reticulum of guinea-pig cardiac cells. J Physiol 405: 233255, 1988.[Abstract]
9. Bridge JH, Ershler PR, and Cannell MB. Properties of Ca2+ sparks evoked by action potentials in mouse ventricular myocytes. J Physiol 518: 469478, 1999.
10. Cannell MB, Berlin JR, and Lederer WJ. Effect of membrane potential changes on the calcium transient in single rat cardiac muscle cells. Science 238: 14191423, 1987.[ISI][Medline]
11. Cannell MB, Cheng H, and Lederer WJ. Spatial non-uniformities in [Ca2+]i during excitation-contraction coupling in cardiac myocytes. Biophys J 67: 19421956, 1994.[Abstract]
12. Cannell MB, Cheng H, and Lederer WJ. The control of calcium release in heart muscle. Science 268: 10451049, 1995.[ISI][Medline]
13. Cannell MB and Soeller C. Numerical analysis of ryanodine receptor activation by L-type channel activity in the cardiac muscle diad. Biophys J 73: 112122, 1997.[Abstract]
14. Cleemann L and Morad M. Role of Ca2+ channel in cardiac excitation-contraction coupling in the rat: evidence from Ca2+ transients and contraction. J Physiol 432: 283312, 1991.[Abstract]
15. Cleemann L, Wang W, and Morad M. Two dimensional confocal images of organization, density, and gating of focal Ca2+ release sites in rat cardiac myocytes. Proc Natl Acad Sci USA 95: 1098410989, 1998.
16. Delgado C, Artiles A, Gomez AM, and Vassort G. Frequency-dependent increase in cardiac Ca2+ current is due to reduced Ca2+ release by the sarcoplasmic reticulum. J Mol Cell Cardiol 31: 17831793, 1999.[CrossRef][ISI][Medline]
17. Eckert R and Chad JE. Inactivation of Ca2+ channels. Prog Biophys Mol Biol 44: 215267, 1984.[CrossRef][ISI][Medline]
18. Eigen M and Wilkins RG. The kinetics and mechanism of formation of metal complexes. In: Mechanisms of Inorganic Reactions. Adv Chem Ser 49: 5567, 1965.
19. Ellis-Davies GC and Kaplan JH. Nitrophenyl-EGTA, a photolabile chelator that selectively binds Ca2+ with high affinity and releases it rapidly upon photolysis. Proc Natl Acad Sci USA 91: 187191, 1994.[Abstract]
20. Ellis-Davies GC, Kaplan JH, and Barsotti RJ. Laser photolysis of caged calcium: rates of calcium release by nitrophenyl-EGTA and DM-nitrophen. Biophys J 70: 10061016, 1996.[Abstract]
21. Franzini-Armstrong C, Protasi F, and Ramesh V. Shape, size, and distribution of Ca2+ release units and couplons in skeletal and cardiac muscles. Biophys J 77: 15281539, 1999.
22. Goldhaber JI, Lamp ST, Walter DO, Garfinkel A, Fukumoto GH, and Weiss JN. Local regulation of the threshold for calcium sparks in rat ventricular myocytes: role of sodium-calcium exchange. J Physiol 520: 431438, 1999.
23. Guia A, Stern MD, Lakatta EG, Josephson IR. Ion concentration-dependence of rat cardiac unitary L-type calcium channel conductance. Biophys J 80: 27422750, 2001.
24. Györke S and Fill M. Ryanodine receptor adaptation: control mechanism of Ca2+-induced Ca2+ release in heart. Science 260: 807809, 1993.[ISI][Medline]
25. Györke S and Palade P. Role of local Ca2+ domains in activation of Ca2+-induced Ca2+ release in crayfish muscle. Am J Physiol Cell Physiol 264: C1505C1512, 1993.
26. Györke S, Velez P, Suarez-Isla B, and Fill M. Activation of single cardiac and skeletal ryanodine receptor channels by flash photolysis of caged Ca2+. Biophys J 66: 18791886, 1994.[Abstract]
27. Hadley RW and Hume JR. An intrinsic potential-dependent inactivation mechanism associated with calcium channels in guinea-pig myocytes. J Physiol 389: 205222, 1987.[Abstract]
28. Hamill OP, Marty A, Neher E, Sakmann B, and Sigworth FJ. Improved patch-clamp techniques for high resolution current recording from cells and cell-free membrane patches. Pflügers Arch 391: 85100, 1991.
29. Inoue M and Bridge JHB. Ca2+ sparks in rabbit ventricular myocytes evoked by action potentials. Involvement of clusters of L-type Ca2+ channels. Circ Res 92: 532538, 2003.
30. Isenberg G and Han S. Gradation of Ca2+-induced Ca2+ release by voltage-clamp pulse duration in potentiated guinea-pig ventricular myocytes. J Physiol 480: 423438, 1994.[Abstract]
31. Jafri MS, Rice JJ, and Winslow RL. Cardiac Ca2+ dynamics: The roles of ryanodine receptor adaptation and sarcoplasmic reticulum load. Biophys J 74: 11491168.
32. Josephson IR, Sanchez-Chapula J, and Brown AM. A comparison of calcium currents in rat and guinea-pig ventricular cells. Circ Res 54: 144156, 1984.[Abstract]
33. Kass RS and Sanguinetti MC. Inactivation of calcium channel current in the calf cardiac Purkinje fiber. J Gen Physiol 84: 705726, 1984.[Abstract]
34. Kubalová Z, Pavelková J, Zahradník I, and Zahradníková A. Calcium tail current-induced, Ca release-dependent inactivation of ICa in cardiac myocytes (Abstract). Biophys J 82: 1201a, 2002.
35. Lamb GD, Laver DR, and Stephenson DG. Questions about adaptation in ryanodine receptors. J Gen Physiol 116: 883890, 2000.
36. Laver DR, Baynes TM, and Dulhunty AF. Magnesium inhibition of ryanodine-receptor calcium channels: evidence for two independent mechanisms. J Membr Biol 156: 213229, 1997.[CrossRef][ISI][Medline]
37. Litwin SE, Li J, and Bridge JH. Na-Ca exchange and the trigger for sarcoplasmic reticulum Ca2+ release: studies in adult rabbit ventricular myocytes. Biophys J 75: 359371, 1998.
38. López-López JR, Shacklock PS, Balke CW, and Wier WG. Local calcium transients triggered by single L-type calcium channel currents in cardiac cells. Science 268: 10421045, 1995.[ISI][Medline]
39. Maeda H, Ellis-Davies GC, Ito K, Miyashita Y, and Kasai H. Supralinear Ca2+ signaling by cooperative and mobile Ca2+ buffering in Purkinje neurons. Neuron 24: 9891002, 1999.[ISI][Medline]
40. Nilius B and Henek M. Pacing dependence of the slow inward current in frog atrial myocardium. Gen Physiol Biophys 1: 307318, 1982.[ISI]
41. Peterson BZ, DeMaria CD, Adelman JP, and Yue DT. Calmodulin is the Ca2+ sensor for Ca2+-dependent inactivation of L-type calcium channels. Neuron 22: 549558, 1999.[ISI][Medline]
42. Puglisi JL, Yuan W, Bassani WM, and Bers DM. Ca2+ influx through Ca2+ channels in rabbit ventricular myocytes during action potential clamp. Circ Res 85: e7e16, 1999.[ISI][Medline]
43. Rose WC, Balke CW, Wier WG, and Marban E. Macroscopic and unitary properties of physiological ion flux through L-type Ca2+ channels in guinea-pig heart cells. J Physiol 456: 277284, 1992.
44. Santana LF, Cheng H, Gómez AM, Cannell MB, and Lederer WJ. Relation between the sarcolemmal Ca2+ current and Ca2+ sparks and local control theories for cardiac excitation-contraction coupling. Circ Res 78: 166171, 1996.
45. Sham JS, Cleemann L, and Morad M. Functional coupling of Ca2+ channels and ryanodine receptors in cardiac myocytes. Proc Natl Acad Sci USA 92: 121125, 1995.[Abstract]
46. Sham JSK. Ca2+ release-induced inactivation of Ca2+ current in rat ventricular myocytes: Evidence for local Ca2+ signaling. J Physiol 500: 285295, 1997.[Abstract]
47. Sham JSK, Song LS, Chen Y, Deng LH, Stern MD, Lakata EG, and Cheng H. Termination of Ca2+ release by a local inactivation of ryanodine receptors in cardiac myocytes. Proc Natl Acad Sci USA 95: 1509615101, 1998.
48. Sipido KR, Callewaert G, and Carmeliet E. Inhibition and rapid recovery of Ca2+ current during Ca2+ release from sarcoplasmic reticulum in guinea pig ventricular myocytes. Circ Res 76: 102109, 1995.
49. Soeller C and Cannell MB. Numerical simulation of local calcium movements during L-type calcium channel gating in the cardiac diad. Biophys J 73: 97111, 1997.[Abstract]
50. Soldatov NM. Ca2+ channel moving tail: link between Ca2+-induced inactivation and Ca2+ signal transduction. Trends Pharmacol Sci 24: 167171, 2003.[CrossRef][ISI][Medline]
51. Song LS, Wang SQ, Xiao RP, Spurgeon H, Lakatta EO, and Cheng H. -Adrenergic stimulation synchronizes intracellular Ca2+ release during excitation-contraction coupling in cardiac myocytes. Circ Res 88: 794801, 2001.
52. Standen NB and Stanfield PR. A binding-site model for calcium channel inactivation that depends on calcium entry. Proc R Soc Lond B Biol Sci 217: 101110, 1982.[ISI][Medline]
53. Stern MD. Theory of excitation-contraction coupling in cardiac muscle. Biophys J 63: 497517, 1992.[Abstract]
54. Viatchenko-Karpinski S and Györke S. Modulation of the Ca2+-induced Ca2+ release cascade by beta-adrenergic stimulation in rat ventricular myocytes. J Physiol 533: 837848, 2001.
55. Wang SQ, Song LS, Lakatta EG, Cheng H. Ca2+ signalling between single L-type Ca2+ channels and ryanodine receptors in heart cells. Nature 410: 592596, 2001.[CrossRef][ISI][Medline]
56. Wier WG, Egan TM, López-López JR, and Balke CW. Local control of excitation-contraction coupling in rat heart cells. J Physiol 474: 463471, 1994.[Abstract]
57. Xu L, Mann G, and Meissner G. Regulation of cardiac Ca2+ release channel (ryanodine receptor) by Ca2+, H+, Mg2+, and adenine nucleotides under normal and simulated ischemic conditions. Circ Res 79: 11001109, 1996.
58. Zahradník I and Palade P. Multiple effects of caffeine on calcium current in rat ventricular myocytes. Pflügers Arch 424: 129136, 1993.[ISI][Medline]
59. Zahradník I, Pavelková J, Györke S, and Zahradníková A:. Tuning of Ca signaling in cardiac E-C coupling by Mg-induced decrease in the rate of RyR Ca binding (Abstract). Biophys J 80: 2675a, 2001.
60. Zahradníková A and Palade P. Procaine effects on single sarcoplasmic reticulum Ca2+ release channels. Biophys J 64: 9911003, 1993.[Abstract]
61. Zahradníková A, Zahradník I, Györke I, and Györke S. Rapid activation of the cardiac ryanodine receptor by submillisecond calcium stimuli. J Gen Physiol 114: 787798, 1999.
62. Zhou YY, Song LS, Lakatta EG, Xiao RP, and Cheng H. Constitutive 2-adrenergic signalling enhances sarcoplasmic reticulum Ca2+ cycling to augment contraction in mouse heart. J Physiol 521: 351361, 1999.