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Originally published In Press as doi:10.1074/jbc.M511971200 on December 22, 2005

J. Biol. Chem., Vol. 281, Issue 8, 4691-4698, February 24, 2006
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Endogenous and Exogenous Ca2+ Buffers Differentially Modulate Ca2+-dependent Inactivation of CaV2.1 Ca2+ Channels*

Lisa Kreiner and Amy Lee1

From the Department of Pharmacology and Center for Neurodegenerative Disease, Emory University School of Medicine, Atlanta, Georgia 30322

Received for publication, November 7, 2005 , and in revised form, December 21, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Voltage-gated Ca2+ channels undergo a negative feedback regulation by Ca2+ ions, Ca2+-dependent inactivation, which is important for restricting Ca2+ signals in nerve and muscle. Although the molecular details underlying Ca2+-dependent inactivation have been characterized, little is known about how this process might be modulated in excitable cells. Based on previous findings that Ca2+-dependent inactivation of Cav2.1 (P/Q-type) Ca2+ channels is suppressed by strong cytoplasmic Ca2+ buffering, we investigated how factors that regulate cellular Ca2+ levels affect inactivation of Cav2.1 Ca2+ currents in transfected 293T cells. We found that inactivation of Cav2.1 Ca2+ currents increased exponentially with current amplitude with low intracellular concentrations of the slow buffer EGTA (0.5 mM), but not with high concentrations of the fast Ca2+ buffer BAPTA (10 mM). However, when the concentration of BAPTA was reduced to 0.5 mM, inactivation of Ca2+ currents was significantly greater than with an equivalent concentration of EGTA, indicating the importance of buffer kinetics in modulating Ca2+-dependent inactivation of Cav2.1. Cotransfection of Cav2.1 with the EF-hand Ca2+-binding proteins, parvalbumin and calbindin, significantly altered the relationship between Ca2+ current amplitude and inactivation in ways that were unexpected from behavior as passive Ca2+ buffers. We conclude that Ca2+-dependent inactivation of Cav2.1 depends on a subplasmalemmal Ca2+ microdomain that is affected by the amplitude of the Ca2+ current and differentially modulated by distinct Ca2+ buffers.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cav2.1 (P/Q-type) voltage-gated Ca2+ channels mediate Ca2+ signals that regulate neuronal excitability, synapse formation, and neurotransmitter release (1–5). Fidelity of Ca2+ signaling by Cav2.1 requires fine control of voltage-gated Ca2+ entry, in part by the Ca2+ ions that permeate the channel. Self-regulation of Cav2.1 channels by Ca2+ is manifest as an initial increase (facilitation) and gradual decrease (inactivation) in Ca2+ current amplitude during high frequency stimuli (6–8). Ca2+-dependent facilitation (CDF)2 and inactivation (CDI) of Cav2.1 channels depend on calmodulin binding to the pore-forming {alpha}1-subunit of Cav2.1 (7, 9) and can cause activity-dependent changes in synaptic efficacy (6, 10, 11).

A fundamental distinction between CDF and CDI is their sensitivity to cytoplasmic Ca2+ buffering. The blockade of CDI, but not CDF, by high concentrations of the Ca2+ chelators EGTA and BAPTA (8, 12) suggests that the extent to which Cav2.1 channels undergo CDI may largely be influenced by factors that regulate intracellular Ca2+ concentrations. Such factors include parvalbumin (PV) and calbindin (CB), which are EF-hand Ca2+-binding proteins that alter the amplitude and time course of Ca2+ signals in some nerve and muscle cells (13–17). Unlike calmodulin, which directly interacts with and confers Ca2+-dependent regulation to numerous effectors (18), PV and CB were generally thought to act as passive Ca2+ buffers, which help protect cells from Ca2+ overloads. However, by modifying the spatial and temporal aspects of intracellular Ca2+ elevations, PV and CB can influence Ca2+ signals that modulate the activity of inositol 1,4,5-trisphosphate receptors (19, 20). Similarly, PV and CB might physiologically regulate Ca2+-dependent modulation of Cav2.1, as PV and CB are concentrated in subsets of neurons, such as cerebellar Purkinje neurons, where Cav2.1 channels are also highly expressed (21–24). Previous studies implicate a role for PV and CB in modulating CDI of L-type voltage-gated Ca2+ channels in neurons (25, 26), although whether these proteins also affect Cav2.1 (P/Q-type) channels is not known.

In this study, we compared the effects of Ca2+ buffers (EGTA and BAPTA) and Ca2+-buffering proteins (PV and CB) on CDI of Cav2.1 in transfected 293T cells. Our analyses indicate that inactivation of Cav2.1 Ca2+ currents varies significantly with current amplitude and is more sensitive to Ca2+ buffering by EGTA than BAPTA. PV and CB do not simply replicate the effects of EGTA and BAPTA, but differentially altered the current dependence of CDI. These findings reveal the importance of cellular Ca2+-buffering mechanisms in the negative feedback regulation of Cav2.1 channels by Ca2+, which may further diversify the properties of these channels in different neuronal cell types (27).


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
cDNA Expression Constructs—Cav2.1 subunits used in electrophysiological experiments were {alpha}12.1 (rbA isoform), beta2a, and {alpha}2{delta} (28–30). cDNAs corresponding to rat parvalbumin and calbindin were isolated by PCR amplification with specific primers from a rat brain cDNA library. Parvalbumin was subcloned into the HindIII/BamHI sites of pcDNA3.1+, and calbindin was subcloned into the BamHI/XhoI sites of pcDNA3.1-topo. The PVCDEF mutant containing amino acid substitutions D51A, E62V, D90A, and E101V was based on that described by Pauls et al. (31) and generated by multiple rounds of QuikChange mutagenesis and subcloning into pcDNA3.1+. The identity of all cDNA constructs was confirmed by sequencing prior to use in electrophysiological experiments.

Cell Culture and Transfection—293T cells were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum at 37 °C under 5% CO2. Cells plated in 35-mm tissue culture dishes were grown to 65–80% confluency and transfected with Gene-PORTER transfection reagent (Gene Therapy Systems Inc., San Diego, CA) according to the manufacturer's protocol with a 1:1 molar ratio of cDNAs for Ca2+ channel subunits (total of 5 µg) and 0.7 µg of a CD8 expression plasmid for identification of transfected cells. Parvalbumin and calbindin cDNAs were transfected at a 1:1 molar ratio with Ca2+ channel subunits.

Western Blots—293T cells, plated and transfected as for electrophysiological experiments, were homogenized in ice-cold lysis buffer (25 mM Tris, pH. 7.4), 137 mM NaCl, 2.7 mM KCl, 0.1% phenylmethylsulfonyl fluoride, and 1% Triton X-100) and stored at –20 °C until use. Cell lysates (50 µg) were electrophoresed on denaturing 4–20% Tris-glycine gels (Invitrogen) and transferred to a nitrocellulose membrane that was blocked in 3% milk/TBS and incubated with antibodies against PV (1:1000, Chemicon International, Temecala, CA), or CB (1:1000, Chemicon International). Chemiluminescent detection was achieved with horseradish peroxidase-conjugated secondary antibodies (1:2000, Amersham Biosciences) and ECL reagents (Amersham Biosciences).

Electrophysiological Recordings—At least 48 h after transfection, 293T cells were incubated with CD8 antibody-coated microspheres (Dynal, Oslo, Norway) for identification of transfected cells. Ca2+ or Ba2+ currents were recorded in whole cell patch clamp recordings with a HEKA EPC-9 patch-clamp amplifier driven by PULSE software (HEKA Electronics, Lambrecht/Pfalz, Germany). Leak and capacitive transients were subtracted using a P/-4 protocol. Extracellular recording solutions contained (in mM): 150 Tris, 1 MgCl2, and 10 CaCl2 or 10 BaCl2. Intracellular recording solutions contained (in mM): 130 N-methyl-D-glucamine, 60 HEPES, 1 MgCl2, 2 Mg-ATP, and EGTA (0.5 mM) or BAPTA (0.5 mM or 10 mM). The pH of extracellular and intracellular recording solutions was adjusted to 7.3 with methanesulfonic acid. Because of shifts in the activation curve of –10 and +10 mV when extracellular Ba2+ or intracellular BAPTA were used, respectively, voltage protocols were adjusted to compensate for this difference as noted.

Data Analysis—All data were analyzed using custom written procedures in IGOR Pro software (Wavemetrics, Portland, OR). Averaged data represent the mean ± S.E. Statistical differences in averaged inactivation (Ires/Ipk) between groups was determined by Student's t test. I-V curves were fit with the function: g(V – E)/{1 + exp[(V – V1/2)/k] + b} where g is the maximum conductance, V is the test potential, E is the apparent reversal potential, V1/2 is the potential of half-activation, k is the slope factor, and b is the baseline. Linear and nonlinear regression and statistical analyses were done with Sigma Plot (SPSS, Inc., Chicago, IL). Significant deviations of percent inactivation data from regression models were determined by Runs test. Data describing current dependence of ICa inactivation were fit with a nonlinear regression equation, y = axb where y is % inactivation, x is the current amplitude, and a and b are constants. F-tests were used for comparisons of nonlinear regression curves, with statistical significance considered as p < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Current Dependence of CDI and Sensitivity to EGTA—Ca2+ microdomains near the pore of individual Ca2+ channels may reach micromolar concentrations, but are too short-lived to be significantly buffered by high concentrations of slow Ca2+ buffers such as EGTA or low concentrations of fast Ca2+ buffers such as BAPTA (32). For this reason, the blockade of CDI of Cav2.1 by high intracellular concentrations of EGTA and BAPTA (10 mM) implies a requirement for a "global" Ca2+ signal that is supported by multiple open channels (8, 9, 12). A simple prediction of this model is that CDI should increase to some extent with the amplitude of the whole cell Ca2+ current (ICa). In support of this prediction, human splice variants of {alpha}12.1 with poor expression levels in transfected cells, as reflected by low amplitude ICa, exhibited less CDI than channel variants with higher mean current amplitudes (12).

In the present study, we also observed a similar relationship between current amplitude and CDI for channels containing the rat brain (rbA) {alpha}12.1 variant (Fig. 1). In transfected 293T cells, inactivation of currents carried by Ca2+ (ICa) or Ba2+ (IBa) was measured as Ires/Ipk, which was the amplitude of the current at the end of a 2-s pulse normalized to the peak current amplitude. With minimal Ca2+ buffering of the intracellular recording solution (0.5 mM EGTA), we found that inactivation was significantly greater (smaller Ires/Ipk) for large ICa (>0.4 nA) than for small ICa (<0.4 nA) (p < 0.01; Fig. 1, A and B) and that the increase in ICa inactivation with current amplitude was significantly nonlinear (p < 0.05) (Fig. 1C). Because Ba2+ does not support calmodulin-dependent conformational changes that underlie CDI of voltage-gated Ca2+ channels (7, 33–36), Cav2.1 currents carried by Ba2+ ions did not vary significantly with current amplitude (Fig. 1B). High concentrations (10 mM) of BAPTA in the intracellular recording solution, which block CDI of Cav2.1 (7, 8), also prevented the current-dependent increase in inactivation (Fig. 1B), such that the relationship between current amplitude and inactivation for ICa+ 10 mM BAPTA and for IBa did not significantly deviate from a straight line (p = 0.83 for ICa+10 BAPTA and p = 0.53 for IBa; Fig. 1C). The positive slope for the current dependence of IBa inactivation (0.02 ± 0.01) could have resulted from Ba2+-dependent effects on inactivation that have been described for L-type Ca2+ channels (37). It is not clear why the corresponding relationship for ICa with 10 mM BAPTA exhibited a negative slope (–0.01 ± 0.01), although stimulatory effects of BAPTA on the amplitude of Cav2.1 currents, have been reported in previous studies (38, 39). The difference between inactivation for ICa and IBa, which reflects the magnitude of CDI, was significantly greater for larger currents (~20%, p < 0.05, Fig. 1, A and B), primarily as a consequence of stronger, current-dependent inactivation of ICa. Our analyses demonstrate that Ca2+ influx and its intracellular accumulation cause nonlinear increases in Cav2.1 inactivation and that current-dependent variations in CDI should be considered when evaluating factors that regulate this process.

To evaluate the modulatory potential of Ca2+ buffers on CDI, we compared the effects of BAPTA and EGTA at concentrations (0.5 mM) that are permissive for CDI. Whereas EGTA and BAPTA bind Ca2+ with nearly equal affinity, Ca2+ on- and off-rates for BAPTA are at least 100 times faster than for EGTA (40). Although BAPTA binds Ca2+ faster than EGTA, it will also retain it for shorter periods of time, such that CDI may be more evident with BAPTA than with EGTA. Consistent with this prediction, Ires/Ipk was significantly smaller with BAPTA (0.5 mM) than with the same concentration of EGTA (~54% for ICa<0.4 nA and ~60% for ICa>0.4 nA, p < 0.01; Fig. 2A). For ICa>0.4 nA, the rate of inactivation with BAPTA (0.5 mM) was significantly faster than with EGTA (~36%, Table 1). The effect of BAPTA was evident as an upward shift in the relationship between ICa inactivation and current amplitude, which was significantly different from that with EGTA (p < 0.001, Fig. 2A). Greater inactivation with BAPTA than with EGTA was particularly apparent during repetitive depolarizations (Fig. 2B). With this voltage protocol and low EGTA (0.5 mM), ICa underwent initial facilitation, which is also calmodulin-dependent (8), followed by inactivation. However, with BAPTA (0.5 mM), only strong inactivation of ICa was observed, such that ICa was reduced ~23% more by the end of the train compared with ICa recorded with EGTA (Fig. 2B). These results show that the ability of ICa to generate Ca2+ signals that cause CDI of Cav2.1 is greater in the presence of fast Ca2+ buffers like BAPTA than with slow Ca2+ buffers like EGTA.


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  		TABLE 1
Effect of Ca2+ buffers on kinetics of inactivation of Cav2.1 Ca2+ current

ICa was evoked by a 2-s test pulse from –80 mV to +10 mV (EGTA) or +20 mV (BAPTA), and inactivation time constants ({tau}) were estimated by fitting the current trace to the equation: I(t) = Io + I[exp(–t/{tau})], where t = time, I is current amplitude, and Io is residual current amplitude at equilibrium. Values are means ± S.E.; n is the number of cells tested.

 


Figure 1
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  		FIGURE 1.
CDI depends on the amplitude of Cav2.1 Ca2+ currents. A, normalized current traces showing Cav2.1 currents evoked by 2-s pulses from –80 mV to +10 mV (ICa+0.5 mM EGTA), 0 mV (IBa), or +20 mV (ICa+10 mM BAPTA) in 293T cells transfected with Cav2.1. Top traces represent ICa (black, solid) and IBa (gray, dashed) with 0.5 mM EGTA in the intracellular recording solution. Bottom traces show ICa recorded with either 0.5 mM EGTA (black, solid) or 10 mM BAPTA (gray, dashed). B, for currents evoked as in A, Ires/Ipk was determined as the current amplitude at the end of the pulse normalized to the peak current amplitude. ICa – IBa was determined by subtracting Ires/Ipk of IBa from that for I. * Ca, p < 0.05 compared with IBa,**, p < 0.05 compared with I < 0.4 nA. C, relationship between peak current amplitude and inactivation. For data obtained in A and B, % inactivation, which was calculated as (1 – Ires/Ipk) x 100, was plotted against peak current amplitude for ICa (•, {blacksquare}) or IBa ({circ}) recorded with 0.5 mM intracellular EGTA (•, {circ}) or 10 mM BAPTA ({blacksquare}). Each point represents a different cell. Smooth line represents fit from nonlinear (ICa + 0.5 EGTA) or linear (IBa, ICa + 10 BAPTA) regression.

 


Figure 2
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  		FIGURE 2.
Inactivation of Cav2.1 Ca2+ currents is greater with BAPTA than with EGTA. A, inactivation during sustained depolarization. ICa was evoked and Ires/Ipk determined as in Fig. 1, but intracellular solution contained EGTA (black bars) or BAPTA (gray bars) at 0.5 mM. Current traces show normalized ICa from cells recorded with 0.5 mM EGTA (black, dashed) or BAPTA (gray, solid). *, p < 0.01. Lower panel shows relationship between % inactivation and current amplitude for individual cells. % inactivation and nonlinear curve fitting was the same as in Fig. 1C. Dashed line indicates curve fit of data in Fig. 1C obtained for ICa with 0.5 mM EGTA. B, inactivation during repetitive stimuli. ICa was evoked by 5-ms pulses from –80 mV to +10 mV (0.5 EGTA, •) or +20 mV (0.5 BAPTA, {circ}) at a frequency of 100 Hz. Fractional current represents test current amplitude normalized to that for the first pulse in the train. Each point represents mean ± S.E. for ICa>0.4 nA. Every second point is plotted against time during the train. Traces above show representative currents evoked by the first and last pulses. Dotted line indicates initial current amplitude.

 
Effects of PV on CDI of Cav2.1—To determine if Ca2+-buffering proteins might similarly regulate inactivation of Cav2.1 Ca2+ currents, we investigated the effect of coexpressing Cav2.1 with parvalbumin (PV, Fig. 3). We chose PV since its Ca2+ binding properties and effects on Ca2+ signals are well characterized (21). The EF-hands of PV can bind Ca2+ with high affinity and Mg2+ with lower affinity (31, 41). Under resting conditions, the concentration of Mg2+ in cells is generally far greater than that for Ca2+ (42), so that the rate of Ca2+ binding is slow because of a requirement for Mg2+ to first unbind (43). As a consequence, PV is considered a slow Ca2+ buffer in cells, with similar Ca2+ binding kinetics as EGTA (21).

Based on our results with EGTA and BAPTA, we expected that PV, like EGTA, should decrease ICa inactivation. Whereas this was true for ICa<0.4 nA (~32% increase in Ires/Ipk for PV-transfected cells, p < 0.05; Fig. 3A), PV had the opposite effect for ICa>0.4 nA, causing significantly greater inactivation of ICa than in cells transfected with Cav2.1 alone (~50% decrease in Ires/Ipk, p < 0.04; Fig. 3A). Kinetic analyses showed that these effects of PV were caused by a slowing and acceleration of the rate of inactivation for small and large amplitude currents, respectively (Table 1). The dual effects of PV on ICa inactivation resulted in a significant deviation in the relationship between ICa inactivation and current amplitude relative to that for Cav2.1 alone (p < 0.005, Fig. 3B). The peak ICa and shape of the I-V curves for cells transfected with Cav2.1 alone or cotransfected with PV were not different for either small or large ICa (Fig. 3C), excluding the possibility that the dual effects of PV on ICa inactivation were caused by alterations in voltage-dependent activation of Cav2.1, or variability in channel expression between groups. Western blots confirmed that PV was highly expressed in cotransfected cells but not in cells transfected with Cav2.1 alone (Fig. 3D). The effects of PV on the current dependence of ICa inactivation were especially apparent during trains of repetitive stimuli (Fig. 4). In these experiments, inactivation of ICa in cells transfected with Cav2.1 alone varied less with peak ICa amplitude than during sustained test pulses (Fig. 1). Because Ca2+-dependent facilitation as well as inactivation is evident with this voltage protocol, temporal overlap of both forms of Ca2+ regulation may have minimized the current dependence of ICa inactivation in cells transfected with Cav2.1 alone (Fig. 4A). However, in cells cotransfected with PV, the dependence of ICa inactivation on current amplitude was more pronounced (Fig. 4B). Consistent with results obtained with sustained test pulses (Fig. 3, A and B), PV inhibited inactivation of small ICa and had the opposite effect on large ICa. The net effect of PV was to significantly augment differences in inactivation for ICa of different peak amplitudes (Fig. 4B).


Figure 3
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  		FIGURE 3.
PV has opposite effects on inactivation of small and large amplitude ICa. A, Ires/Ipk was determined as in Fig. 1 for ICa in cells transfected with Cav2.1 alone (black, dashed) or cotransfected with PV (gray, solid) for ICa less than or greater than 0.4 nA. Representative normalized current traces are shown above. Intracellular solution contained 0.5 mM EGTA and extracellular solution contained 10 mM Ca2+. *, p < 0.05. B, current dependence of ICa inactivation with PV. Dashed line indicates curve fit of data in Fig. 1C obtained for ICa (+0.5 EGTA) in cells transfected with Cav2.1 alone. Solid line indicates curve fit of data for cells cotransfected with PV. C, current-voltage (I-V) curves for cells transfected with Cav2.1 alone (open symbols) or cotransfected with PV (filled symbols) are shown for ICa<0.4 nA (triangles) or >0.4 nA (circles). D, Western blot detection of PV in cells cotransfected with PV but not in cells transfected with Cav2.1 alone.

 
The Ca2+-buffering properties of PV could account for the suppression of inactivation of small currents, but we wondered if the opposite effect of PV on large currents could have resulted from Ca2+ unbinding from PV. Large ICa through Cav2.1 channels could rapidly saturate the EF-hand Ca2+ binding sites on PV, and subsequent Ca2+ release from PV might then facilitate CDI. At the concentration of free intracellular Mg2+ in our experiments (~ 1 mM), estimated rates of Ca2+ unbinding from PV (~0.9/s, Ref. 44) are consistent with the potential for Ca2+ release from PV to occur within the 2-s depolarizing pulse in our experiments (Fig. 3A). In this context, both the inhibitory and stimulatory effects of PV on ICa inactivation should depend critically on the ability of PV to bind Ca2+. To test this, we generated a PV construct with mutations in the second and third EF-hands (PVCDEF). Because these alterations prevent binding of Ca2+ and Mg2+ (31, 45), PVCDEF should not replicate Ca2+-dependent effects of PV on Cav2.1 inactivation. In these experiments, Ires/Ipk for ICa was not affected by PVCDEF for either small or large amplitude ICa (p = 0.52 and 0.71, respectively), and the relationship between ICa inactivation and current amplitude was not significantly different from that for Cav2.1 channels expressed alone (p = 0.62, Fig. 5A). These results were not caused by limited expression levels of PVCDEF, because Western blots indicated similar amounts of PV and PVCDEF in lysates harvested from the cotransfected cells (not shown). The dual effects of wild-type PV on ICa inactivation were also not observed when CDI was blocked by a high intracellular concentration (10 mM) of BAPTA (p = 0.17 for ICa<0.4 nA, p = 0.45 for ICa>0.4 nA; Fig. 5B). Moreover, there was no effect of PV on the dependence of ICa inactivation on current amplitude in the presence of 10 mM BAPTA (Fig. 5B). These results argued against the possibility that PV influenced Ca2+-independent (i.e. voltage-dependent) mechanisms of Cav2.1 inactivation, because such actions would have been spared by BAPTA. We conclude that the opposing effects of PV on inactivation of large and small ICa are a consequence of Ca2+ binding to PV, which can either stabilize or diffuse Ca2+ pools that underlie CDI, depending on the amplitude of ICa.


Figure 4
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  		FIGURE 4.
PV alters CDI during repetitive stimuli. A and B, left panels, fractional current was determined as in Fig. 2B and plotted against time for cells transfected with Cav2.1 alone (A) or contransfected with PV (B) for ICa of different amplitudes indicated in A. Right panels, fractional current for the last 10 pulses were averaged and compared for the different groups. Numbers of cells are shown in parentheses and p values from one-way analysis of variance are indicated.

 
Effects of CB on CDI—We investigated further the role of Ca2+ buffer kinetics and CDI in cells cotransfected with calbindin-D28k (CB), a protein with physiological Ca2+ binding rates that are faster, by an order of magnitude, than those for PV (25). CB also differs from PV in possessing four functional EF-hands, which have relatively low affinity for Ca2+ compared with those in PV, and do not significantly bind Mg2+ (25). In analyses of ICa evoked by step depolarizations, CB significantly inhibited inactivation of ICa < 0.4 nA (~35% increase in Ires/Ipk compared with Cav2.1 alone, p < 0.02, Fig. 6A) and caused an even greater slowing of inactivation rate than PV (~52% for CB and ~36% for PV, Table 1). For unknown reasons, Cav2.1 Ca2+ currents in cells cotransfected with CB that exceeded 0.8 nA were not stable for electrophysiological recordings. Because CB has been shown to directly interact with and potentially regulate cellular proteins in a Ca2+-dependent manner (46, 47), it is possible that CB had a deleterious effect in cells expressing particularly large numbers of Cav2.1 channels. Therefore, analysis of ICa> 0.4 nA in cells transfected with Cav2.1 alone or cotransfected with CB was restricted to currents with amplitude from 0.4 to 0.8 nA. In this current range, CB did not significantly influence the magnitude or rate of inactivation (p = 0.39, Fig. 6A and Table 1). The suppression of inactivation of low amplitude currents caused a significant downward shift in the relationship between inactivation and ICa amplitude in cells cotransfected with CB compared with that in cells transfected with Cav2.1 alone (p < 0.02, Fig. 6B). The absence of ICa>0.8 nA in cells cotransfected with Cav2.1 and CB precluded determination if, like PV, CB had a stimulatory effect on inactivation of large amplitude currents, as these effects were generally observed in cells cotransfected with PV for ICa>0.8 nA (Fig. 3B). However, the inhibitory effects of both PV and CB on inactivation of small amplitude Cav2.1 Ca2+ currents is consistent with a modulatory role for these Ca2+-buffering proteins in the negative feedback of Cav2.1 by Ca2+.


Figure 5
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  		FIGURE 5.
Effects of PV require Ca2+ binding to PV and are specific for CDI. A and B, left panels, Ires/Ipk was determined as in Fig. 1 for ICa less than or greater than 0.4 nA. Traces above show normalized ICa in cells transfected alone (black, dashed), cotransfected with PVCDEF (gray, solid, A), or cotransfected with PV (gray, solid, B). Right, relationship between % inactivation and ICa amplitude for cells transfected with Cav2.1+PVCDEF (circles, A) or Cav2.1+PV (squares, B). Dashed line represents data replotted from Fig. 1C. Smooth line shows fit from nonlinear (A) and linear (B) regression.

 
Besides increasing the Ca2+-buffering capacity of the cell, PV and CB in the transfected 293T cells may have had secondary effects that might influence Cav2.1 properties in our experiments. For example, overexpression of PV and CB could have reduced the levels of endogenous calmodulin required for CDI, which could also explain the inhibition of ICa inactivation in cells cotransfected with Cav2.1 and PV or CB (Figs. 3 and 6). However, Western blots indicated equivalent levels of calmodulin in cells transfected with Cav2.1 alone and in cells cotransfected with PV or CB (not shown). In addition, cotransfection of Cav2.1 with PV and CB did not alter the magnitude of Ca2+-dependent facilitation of ICa, which also depends on calmodulin (Fig. 7, A–C). The enhancement of ICa amplitude during the first 200 ms of a repetitive stimulus protocol was not significantly different in cells transfected with Cav2.1 alone and those cotransfected with PV (p = 0.30) or CB (p = 0.54). The ineffectiveness of PV and CB in these experiments can be explained by a reliance of facilitation on local Ca2+ increases that are not able to be suppressed by even high concentrations of EGTA and BAPTA (8, 12, 48). These results also show that calmodulin was not likely to be a limiting factor contributing to the reduced CDI in cells expressing Cav2.1 with PV or CB. Taken together, our findings indicate that PV and CB modulate Cav2.1 Ca2+ currents by regulating global Ca2+ microdomains that support CDI.


Figure 6
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  		FIGURE 6.
CB inhibits inactivation of small amplitude ICa. A, Ires/Ipk was determined as in Fig. 1 for cells transfected with Cav2.1 alone (black bars) or cotransfected with CB (hatched bars). Current traces show ICa from cells cotransfected with CB (gray, solid) normalized to ICa from cells transfected with Cav2.1 alone (black, dashed) for ICa less than or greater than 0.4 nA. *, p < 0.05. Inset, Western blot shows expression of CB in cells cotransfected with CB and Cav2.1 but not in cells transfected with Cav2.1 alone. B, current dependence of ICa inactivation. Individual data points were plotted and fit (smooth line) as in Fig. 3B. Curve fit of data from cells transfected with Cav2.1 alone was replotted from Fig. 1C (dashed line).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study, the use of endogenous and exogenous Ca2+ buffers revealed new insights into the feedback regulation of Cav2.1 channels by Ca2+. First, we confirmed and extended previous findings (12) that Cav2.1 inactivation increases nonlinearly with the amplitude of ICa, an effect not observed when intracellular Ca2+ is buffered with BAPTA (10 mM) or when Ba2+ is the charge carrier. Second, BAPTA, at submaximal concentrations, strengthens the relationship between current amplitude and CDI compared with EGTA, perhaps because of its faster Ca2+ unbinding kinetics. Third, the Ca2+-buffering proteins PV and CB also alter the Ca2+ current dependence of Cav2.1 inactivation. These findings illustrate how factors affecting Cav2.1 expression and Ca2+ homeostasis may dynamically regulate Cav2.1 properties and Ca2+ signaling in excitable cells.


Figure 7
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  		FIGURE 7.
PV and CB do not influence Ca2+-dependent facilitation of Cav2.1. A, repetitive voltage pulses were applied to cells transfected with Cav2.1 alone ({circ}) or cotransfected with PV (•) as in Fig. 2B. Results are shown for the first 200 ms in cells with ICa < 0.4 nA. B, same as in A, except cells were transfected with Cav2.1 alone ({circ}) or cotransfected with CB (•). C, % facilitation for cells transfected with Cav2.1 alone (white bar) or cotransfected with PV (black bar) or CB (hatched bar) was determined as: [(average fractional current for the first 200 ms) – 1] x 100.

 
Differential Modulation of Cav2.1 Inactivation by Ca2+ Buffers— Based on our results and those published previously, we propose a qualitative model to account for the Ca2+ dependence of Cav2.1 inactivation (Fig. 8). Unlike the relative invariance of the local Ca2+ microdomain associated with individual channel openings, Ca2+ pools that support CDI may be enhanced by larger ICa because of increased overlap of global Ca2+ signals emanating from neighboring channels (Fig. 8A). Small ICa may inactivate less than large ICa because, when not supported by multiple channels, these Ca2+ pools dissipate rapidly due to diffusion of Ca2+ (Figs. 1 and 8A). These Ca2+ gradients surrounding individual channels are collapsed by high concentrations of BAPTA (Fig. 1) or EGTA (8), thus abolishing the dependence of CDI on current amplitude (Figs. 1C and 8A). However, at concentrations of Ca2+ buffers that are permissive for CDI, Ca2+ unbinding from the buffer may paradoxically stabilize the Ca2+ pool supporting CDI (Fig. 8, B and C). Considering first the increased CDI seen with BAPTA compared with EGTA (0.5 mM, Fig. 2), the 100-fold faster association and dissociation rate of Ca2+ from BAPTA allows it to capture but also unload Ca2+ more rapidly than EGTA (40). Therefore, the time that Ca2+ will remain bound before dissociating is considerably less for BAPTA (~6–60 ms), than for EGTA (~700 –2000 ms) (49), such that more rapid Ca2+ unbinding from BAPTA (0.5 mM) may cause greater CDI of both small and large ICa compared with the same concentration of EGTA (Figs. 2 and 8B).


Figure 8
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  		FIGURE 8.
Model for differential regulation of CDI by Ca2+ buffers. Ca2+ microdomains associated with individual Cav2.1 channels are considered as a rapidly accumulating BAPTA-insensitive compartment near the pore (dark gray), which diffuses into a more slowly filling pool away from the pore (light gray), which supports CDI and can be buffered by Ca2+ chelators. A, with 0.5 mM EGTA, large ICa (right) shows more CDI than small ICa (left) because of overlap of Ca2+ microdomains (asterisks). High BAPTA (10 mM) suppresses Ca2+ and CDI for small and large ICa. B, limiting concentrations (0.5 mM) of BAPTA cause more CDI because of faster Ca2+ unbinding than EGTA (0.5 mM) for small and large ICa. C, in the presence of 0.5 mM EGTA, PV, and CB inhibit Ca2+ microdomains when channel density is low (left), but Ca2+ unbinding from PV may increase Ca2+ supporting CDI when channel density is high (right).

 
The dual effects of PV on Cav2.1 inactivation may result not only from its slow Ca2+ binding properties but also more limited mobility of PV relative to EGTA and BAPTA. Because of its larger size, PV has a diffusional range (~43 µm2 s–1) that is about five times less than that for EGTA (~200 µm2 s–1) (44). When Cav2.1 channels are sparsely distributed in the plasma membrane (small ICa), PV may have a net Ca2+ buffering effect in reducing CDI (Fig. 3), as PV may bind and shuttle Ca2+ away from channels prior to releasing it (17). However, with increased channel density (large ICa), Ca2+-saturated PV may unload Ca2+ within microdomains that support CDI of neighboring channels (Fig. 8C), therefore increasing inactivation of large amplitude ICa compared with in cells transfected with Cav2.1 alone (Figs. 3 and 4). Although CB has fast Ca2+ binding kinetics more comparable to BAPTA than EGTA, CB also has considerably lower Ca2+ binding affinity (~1.5 µM) and diffusional range (~20 µm2 s–1) than BAPTA (46, 50). Together, these factors may explain why CB behaved more like a slow Ca2+ buffer in inhibiting CDI only for low amplitude currents (Fig. 6).

For the following reasons, we consider the relatively modest effects of PV and CB on CDI in our experiments to underestimate the potential for these Ca2+-buffering proteins to influence Cav2.1 channels in neurons. First, the impact of PV was likely attenuated by the concentration of free Mg2+ in our intracellular recording solution (~1mM). Because of the mixed Ca2+/Mg2+ affinity of PV EF-hands (31, 51), PV would be occupied mainly by Mg2+ ions prior to evoking ICa in our experiments. Mg2+ unbinding from PV would therefore retard the rate of Ca2+ binding. Given that the estimated concentration of Mg2+ in neurons is ~0.3– 0.6 mM (42, 52), we would predict more rapid Ca2+ binding by PV in neurons than in our transfected cell recordings, which should enhance its current-dependent modulation of CDI (Figs. 3 and 4). Second, PV and CB are found at concentrations in neurons (100 µM-5 mM) (53–56) that may exceed that in our transfected cells. Whereas Western blots of transfected cell lysates indicated strong levels of overexpressed PV and CB (Figs. 3D and 6A), heterogeneous levels of PV and CB between cells could have contributed to intercellular variability and weakened the average impact on CDI. Finally, washout of PV and CB from transfected cells into the recording pipette solution during whole cell recordings may have diminished the intracellular content of Ca2+-buffering proteins. Whole cell patch clamp recordings of dentate granule cells in hippocampal slices suggest considerable dilution of endogenous CB within the first 5 min of obtaining whole cell configuration (57). Although we measured inactivation at similar time points between cells (~3–5 min after establishing whole cell mode) to limit variability, initial washout of PV or CB upon patch rupture could have significantly reduced the overall impact of these proteins on CDI.

Ca2+-buffering Proteins as Regulators of Ca2+ Signaling—The effects of PV in altering feedback regulation of Cav2.1 channels parallels its modulation of inositol 1,4,5-trisphosphate receptors (IP3Rs), which mediate Ca2+ release from intracellular stores (19, 20, 49). Overexpression of PV in Xenopus oocytes was found to stimulate Ca2+-dependent activation of IP3Rs by facilitating Ca2+ diffusion between neighboring channels (19). That BAPTA, but not CB, replicated the effect of PV in this system was interpreted as a sign that the Ca2+ binding affinity and diffusional range of a Ca2+-buffering protein were important determinants of the ability to enhance IP3R activation by Ca2+ (19). The similar effects of PV in stimulating Ca2+ feedback of Cav2.1 channels and IP3Rs may exemplify fundamental mechanisms controlling Ca2+-dependent activation of other signaling pathways. In addition, the dual potential for PV to act as a passive Ca2+ buffer and a facilitator of intracellular and plasma membrane Ca2+ channels may contribute to the paradox that PV can be neuroprotective from pathological Ca2+ overloads in some neurons (58), but can also exacerbate excitotoxic cell death in others (59, 60).

Our findings also provide direct support for previous observations that Ca2+-buffering proteins inhibit CDI of voltage-gated Ca2+ channels in neurons. Infusion of PV and CB in thalamic relay neurons caused an inhibition of CDI of high voltage activated Ca2+ current, although this effect was primarily on L-type channels (26). In addition, loss of CB from surviving granule cells isolated from the hippocampus of patients with mesial temporal lobe epilepsy correlated with increased CDI of voltage-gated Ca2+ currents, which was restored by inclusion of CB in the recording electrode (25). Interestingly, Chaudhuri et al. (24) found that Cav2.1 channels in cerebellar Purkinje neurons show highly variable CDI, which may result partly from cellular variations in Cav2.1 subunits expressed. Our results suggest that developmental or pathological alterations in PV and CB, as well as in the levels of Cav2.1 expression, could add to the heterogeneity of Cav2.1 inactivation in these and other neurons. Such variations in Cav2.1 properties may be important for tailoring Ca2+ influx according to particular physiological contexts.


   FOOTNOTES
 
* This work was supported by Grant NS044922 from the National Institutes of Health (to A. L.), the Whitehall Foundation, and a Predoctoral NRSA Grant F31NS049757 (to L. K.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed: Dept. of Pharmacology, Emory University School of Medicine, 5123 Rollins Research Bldg., 1510 Clifton Rd., Atlanta, GA 30322. Tel.: 404-727-5991; Fax: 404-727-0365; E-mail: alee{at}pharm.emory.edu.

2 The abbreviations used are: CDF, Ca2+-dependent facilitation; CDI, Ca2+-dependent inactivation; PV, parvalbumin; CB, calbindin; BAPTA, 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid; IP, inositol phosphate. Back


   ACKNOWLEDGMENTS
 
We thank Hong Sun for technical assistance, Dr. Frank Gordon for advice on statistical analysis, and Dr. Beat Schwaller for comments on the manuscript.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Cavelier, P., Pouille, F., Desplantez, T., Beekenkamp, H., and Bossu, J. L. (2002) J. Physiol. (Lond.) 540, 57–72[Abstract/Free Full Text]
  2. Miyazaki, T., Hashimoto, K., Shin, H.-S., Kano, M., and Watanabe, M. (2004) J. Neurosci. 24, 1734–1743[Abstract/Free Full Text]
  3. Wheeler, D. B., Randall, A., and Tsien, R. W. (1996) J. Neurosci. 16, 2226–2237[Abstract/Free Full Text]
  4. Dunlap, K., Luebke, J. I., and Turner, T. J. (1995) Trends Neurosci. 18, 89–98[CrossRef][Medline] [Order article via Infotrieve]
  5. Jun, K., Piedras-Renteria, E. S., Smith, S. M., Wheeler, D. B., Lee, S. B., Lee, T. G., Chin, H., Adams, M. E., Scheller, R. H., Tsien, R. W., and Shin, H. S. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 15245–15250[Abstract/Free Full Text]
  6. Forsythe, I. D., Tsujimoto, T., Barnes-Davies, M., Cuttle, M. F., and Takahashi, T. (1998) Neuron 20, 797–807[CrossRef][Medline] [Order article via Infotrieve]
  7. Lee, A., Wong, S. T., Gallagher, D., Li, B., Storm, D. R., Scheuer, T., and Catterall, W. A. (1999) Nature 399, 155–159[CrossRef][Medline] [Order article via Infotrieve]
  8. Lee, A., Scheuer, T., and Catterall, W. A. (2000) J. Neurosci. 20, 6830–6838[Abstract/Free Full Text]
  9. DeMaria, C. D., Soong, T. W., Alseikhan, B. A., Alvania, R. S., and Yue, D. T. (2001) Nature 411, 484–489[CrossRef][Medline] [Order article via Infotrieve]
  10. Cuttle, M. F., Tsujimoto, T., Forsythe, I. D., and Takahashi, T. (1998) J. Physiol. (Lond.) 512, 723–729[Abstract/Free Full Text]
  11. Borst, J. G., and Sakmann, B. (1998) J. Physiol. (Lond.) 513, 149–155[Abstract/Free Full Text]
  12. Soong, T. W., DeMaria, C. D., Alvania, R. S., Zweifel, L. S., Liang, M. C., Mittman, S., Agnew, W. S., and Yue, D. T. (2002) J. Neurosci. 22, 10142–10152[Abstract/Free Full Text]
  13. Schmidt, H., Stiefel, K. M., Racay, P., Schwaller, B., and Eilers, J. (2003) J. Physiol. (Lond.) 551, 13–32[Abstract/Free Full Text]
  14. Li, Z., Decavel, C., and Hatton, G. I. (1995) J. Physiol. (Lond.). 488, 601–608[Medline] [Order article via Infotrieve]
  15. Schwaller, B., Dick, J., Dhoot, G., Carroll, S., Vrbova, G., Nicotera, P., Pette, D., Wyss, A., Bluethmann, H., Hunziker, W., and Celio, M. R. (1999) Am. J. Physiol. 276, C395–C403
  16. Lee, S. H., Rosenmund, C., Schwaller, B., and Neher, E. (2000) J. Physiol. 525, 405–418[Abstract/Free Full Text]
  17. Lee, S. H., Schwaller, B., and Neher, E. (2000) J. Physiol. 525, 419–432[Abstract/Free Full Text]
  18. Chin, D., and Means, A. R. (2000) Trends Cell Biol. 10, 322–328[CrossRef][Medline] [Order article via Infotrieve]
  19. John, L. M., Mosquera-Caro, M., Camacho, P., and Lechleiter, J. D. (2001) J. Physiol. (Lond.) 535, 3–16[Abstract/Free Full Text]
  20. Dargan, S. L., Schwaller, B., and Parker, I. (2004) J. Physiol. (Lond.) 556, 447–461[Abstract/Free Full Text]
  21. Schwaller, B., Meyer, M., and Schiffmann, S. (2002) Cerebellum 1, 241–258[CrossRef][Medline] [Order article via Infotrieve]
  22. Kadowaki, K., McGowan, E., Mock, G., Chandler, S., and Emson, P. C. (1993) Neurosci. Lett. 153, 80–84[CrossRef][Medline] [Order article via Infotrieve]
  23. Mintz, I. M., Adams, M. E., and Bean, B. P. (1992) Neuron 9, 85–95[CrossRef][Medline] [Order article via Infotrieve]
  24. Chaudhuri, D., Alseikhan, B. A., Chang, S. Y., Soong, T. W., and Yue, D. T. (2005) J. Neurosci. 25, 8282–8294[Abstract/Free Full Text]
  25. Nagerl, U. V., Mody, I., Jeub, M., Lie, A. A., Elger, C. E., and Beck, H. (2000) J. Neurosci. 20, 1831–1836[Abstract/Free Full Text]
  26. Meuth, S. G., Kanyshkova, T., Landgraf, P., Pape, H.-C., and Budde, T. (2005) Pflugers Archiv. 450, 111–122[CrossRef][Medline] [Order article via Infotrieve]
  27. Llinas, R., Sugimori, M., Hillman, D. E., and Cherksey, B. (1992) Trends Neurosci. 15, 351–355[CrossRef][Medline] [Order article via Infotrieve]
  28. Stea, A., Tomlinson, W. J., Soong, T. W., Bourinet, E., Dubel, S. J., Vincent, S. R., and Snutch, T. P. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 10576–10580[Abstract/Free Full Text]
  29. Perez-Reyes, E., Castellano, A., Kim, H. S., Bertrand, P., Baggstrom, E., Lacerda, A. E., Wei, X. Y., and Birnbaumer, L. (1992) J. Biol. Chem. 267, 1792–1797[Abstract/Free Full Text]
  30. Starr, T. V., Prystay, W., and Snutch, T. P. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 5621–5625[Abstract/Free Full Text]
  31. Pauls, T. L., Durussel, I., Berchtold, M. W., and Cox, J. A. (1994) Biochemistry 33, 10393–10400[CrossRef][Medline] [Order article via Infotrieve]
  32. Stern, M. D. (1992) Cell Calcium 13, 183–192[CrossRef][Medline] [Order article via Infotrieve]
  33. Brehm, P., and Eckert, R. (1978) Science 202, 1203–1206[Abstract/Free Full Text]
  34. Peterson, B. Z., DeMaria, C. D., Adelman, J. P., and Yue, D. T. (1999) Neuron 22, 549–558[CrossRef][Medline] [Order article via Infotrieve]
  35. Zühlke, R. D., Pitt, G. S., Deisseroth, K., Tsien, R. W., and Reuter, H. (1999) Nature 399, 159–162[CrossRef][Medline] [Order article via Infotrieve]
  36. Qin, N., Olcese, R., Bransby, M., Lin, T., and Birnbaumer, L. (1999) Proc. Natl. Acad. Sci. U .S. A. 96, 2435–2438[Abstract/Free Full Text]
  37. Ferreira, G., Yi, J., Rios, E., and Shirokov, R. (1997) J. Gen. Physiol. 109, 449–461[Abstract/Free Full Text]
  38. Bodding, M., and Penner, R. (1999) Pflugers Arch. 439, 27–38[CrossRef][Medline] [Order article via Infotrieve]
  39. Rousset, M., Cens, T., Gouin-Charnet, A., Scamps, F., and Charnet, P. (2004) J. Biol. Chem. 279, 14619–14630[Abstract/Free Full Text]
  40. Tsien, R. Y. (1980) Biochemistry 19, 2396–2404[CrossRef][Medline] [Order article via Infotrieve]
  41. Haiech, J., Derancourt, J., Pechere, J. F., and Demaille, J. G. (1979) Biochemistry 18, 2752–2758[CrossRef][Medline] [Order article via Infotrieve]
  42. Li-Smerin, Y., Levitan, E. S., and Johnson, J. W. (2001) J. Physiol. 533, 729–743[Abstract/Free Full Text]
  43. Hou, T. T., Johnson, J. D., and Rall, J. A. (1991) J. Physiol. 441, 285–304[Abstract/Free Full Text]
  44. Schmidt, H., Brown, E. B., Schwaller, B., and Eilers, J. (2003) Biophys. J. 84, 2599–2608[Abstract/Free Full Text]
  45. Pauls, T. L., Durussel, I., Cox, J. A., Clark, I. D., Szabo, A. G., Gagne, S. M., Sykes, B. D., and Berchtold, M. W. (1993) J. Biol. Chem. 268, 20897–20903[Abstract/Free Full Text]
  46. Schmidt, H., Schwaller, B., and Eilers, J. (2005) Proc. Natl. Acad. Sci. U. S. A. 102, 5850–5855[Abstract/Free Full Text]
  47. Lutz, W., Frank, E. M., Craig, T. A., Thompson, R., Venters, R. A., Kojetin, D., Cavanagh, J., and Kumar, R. (2003) Biochem. Biophys. Res. Commun. 303, 1186–1192[CrossRef][Medline] [Order article via Infotrieve]
  48. Chaudhuri, D., Chang, S.-Y., DeMaria, C. D., Alvania, R. S., Soong, T. W., and Yue, D. T. (2004) J. Neurosci. 24, 6334–6342[Abstract/Free Full Text]
  49. Dargan, S. L., and Parker, I. (2003) J. Physiol. (Lond.) 553, 775–788[Abstract/Free Full Text]
  50. Nagerl, U. V., Novo, D., Mody, I., and Vergara, J. L. (2000) Biophys. J. 79, 3009–3018[Abstract/Free Full Text]
  51. Gillis, J. M., Thomason, D., Lefevre, J., and Kretsinger, R. H. (1982) J. Muscle Res. Cell Motil. 3, 377–398[CrossRef][Medline] [Order article via Infotrieve]
  52. Stout, A. K., Li-Smerin, Y., Johnson, J. W., and Reynolds, I. J. (1996) J. Physiol. (Lond.) 492, 641–657[Medline] [Order article via Infotrieve]
  53. Plogmann, D., and Celio, M. R. (1993) Brain Res. 600, 273–279[CrossRef][Medline] [Order article via Infotrieve]
  54. Kosaka, T., Kosaka, K., Nakayama, T., Hunziker, W., and Heizmann, C. W. (1993) Exp. Brain Res. 93, 483–491[Medline] [Order article via Infotrieve]
  55. Fierro, L., and Llano, I. (1996) J. Physiol. (Lond.) 496, 617–625[Medline] [Order article via Infotrieve]
  56. Hackney, C. M., Mahendrasingam, S., Penn, A., and Fettiplace, R. (2005) J. Neurosci. 25, 7867–7875[Abstract/Free Full Text]
  57. Muller, A., Kukley, M., Stausberg, P., Beck, H., Muller, W., and Dietrich, D. (2005) J. Neurosci. 25, 558–565[Abstract/Free Full Text]
  58. Van Den Bosch, L., Schwaller, B., Vleminckx, V., Meijers, B., Stork, S., Ruehlicke, T., Van Houtte, E., Klaassen, H., Celio, M. R., Missiaen, L., Robberecht, W., and Berchtold, M. W. (2002) Exp. Neurol. 174, 150–161[CrossRef][Medline] [Order article via Infotrieve]
  59. Maetzler, W., Nitsch, C., Bendfeldt, K., Racay, P., Vollenweider, F., and Schwaller, B. (2004) Exp. Neurol. 186, 78–88[CrossRef][Medline] [Order article via Infotrieve]
  60. Hartley, D. M., Neve, R. L., Bryan, J., Ullrey, D. B., Bak, S. Y., Lang, P., and Geller, A. I. (1996) Brain Res. Mol. Brain Res. 40, 285–296[Medline] [Order article via Infotrieve]

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