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Arachidonic acid (AA) inhibits the activity of several different voltage-gated Ca2+ channels by an unknown mechanism at an unknown site. The Ca2+ channel pore-forming subunit (CaVα1) is a candidate for the site of AA inhibition because T-type Ca2+ channels, which do not require accessory subunits for expression, are inhibited by AA. Here, we report the unanticipated role of accessory CaVβ subunits on the inhibition of CaV1.3b L-type (L-) current by AA. Whole cell Ba2+ currents were measured from recombinant channels expressed in human embryonic kidney 293 cells at a test potential of −10 mV from a holding potential of −90 mV. A one-minute exposure to 10 µM AA inhibited currents with β1b, β3, or β4 58, 51, or 44%, respectively, but with β2a only 31%. At a more depolarized holding potential of −60 mV, currents were inhibited to a lesser degree. These data are best explained by a simple model where AA stabilizes CaV1.3b in a deep closed-channel conformation, resulting in current inhibition. Consistent with this hypothesis, inhibition by AA occurred in the absence of test pulses, indicating that channels do not need to open to become inhibited. AA had no effect on the voltage dependence of holding potential–dependent inactivation or on recovery from inactivation regardless of CaVβ subunit. Unexpectedly, kinetic analysis revealed evidence for two populations of L-channels that exhibit willing and reluctant gating previously described for CaV2 channels. AA preferentially inhibited reluctant gating channels, revealing the accelerated kinetics of willing channels. Additionally, we discovered that the palmitoyl groups of β2a interfere with inhibition by AA. Our novel findings that the CaVβ subunit alters kinetic changes and magnitude of inhibition by AA suggest that CaVβ expression may regulate how AA modulates Ca2+-dependent processes that rely on L-channels, such as gene expression, enzyme activation, secretion, and membrane excitability.
In the nervous system, voltage-gated L-type (L-) Ca2+ channels are composed of several proteins: the pore-forming CaVα1 subunit, through which Ca2+ ions pass, and accessory CaVβ and α2δ subunits (Catterall, 2000). Neurons in the brain express two isoforms of the L-channel CaVα1 subunit: CaV1.2 and CaV1.3 (Hell et al., 1993). The CaV1.3 isoform plays a role in gene expression (Gao et al., 2006; Zhang et al., 2006), exocytosis (Brandt et al., 2005), and membrane excitability (Brandt et al., 2003; Olson et al., 2005), depending on the cell type and localization.
L-channel activity is inhibited by signal transduction pathways downstream of neurotransmitters, including certain types of dopamine (Wikstrom et al., 1999; Banihashemi and Albert, 2002; Olson et al., 2005), glutamate (Chavis et al., 1994), serotonin (Cardenas et al., 1997; Day et al., 2002), and acetylcholine receptors (Pemberton and Jones, 1997; Bannister et al., 2002; Liu et al., 2006). Activation of these G protein–coupled receptors (GPCRs) also releases arachidonic acid (AA; C20:4) (Axelrod et al., 1988; Lazarewicz et al., 1992; Yehuda et al., 1998; Tang et al., 2006). Our laboratory has documented that endogenous AA release is necessary for muscarinic M1 receptor (M1) inhibition of L-current in superior cervical ganglion (SCG) neurons (Liu et al., 2006). Moreover, exogenously applied AA inhibits L-current in SCG neurons similarly to M1R agonists (Liu et al., 2006). The CaV1.3b L-channel isoform has been detected and cloned from SCG neurons (Lin et al., 1996), suggesting that endogenous AA modulates CaV1.3b.
The mechanism by which AA acts downstream of GPCR activation to inhibit L-current remains incompletely characterized. Single-channel recordings from SCG indicate that AA decreases the open probability of L-channels by increasing the dwell time in a closed state with no effect on unitary channel conductance (Liu and Rittenhouse, 2000). Similar findings of AA affecting closed states have been reported for the T-type (T-) Ca2+ channel, CaV3.1 (Talavera et al., 2004). A second family member, CaV3.2, is also inhibited by AA but via a leftward shift in holding potential–dependent inactivation (Zhang et al., 2000). Additionally, both T-channel studies reported increases in the rate of fast inactivation after AA, whereas our study on whole cell SCG L-current revealed no such changes (Liu et al., 2001). One obvious difference between T- and L-channels is that T-channels lack the recognition sequence in the I-II linker for binding CaVβ subunits (Arias et al., 2005), whereas CaVβ binding to L-channels fine-tunes their kinetics and voltage dependence of activation and inactivation (Singer et al., 1991; Hering et al., 2000; Kobrinsky et al., 2004). Whether certain CaVβ subunits block kinetic changes elicited by AA or whether CaV1.3 lacks a homologous site that confers the kinetic changes is unknown.
Therefore, to examine the extent of AA's actions on L-channel activity, we tested whether coexpression of CaV1.3b with different CaVβ subunits accounts for the lack of kinetic changes observed by AA inhibition of whole cell L-current in SCG neurons. We show that AA inhibits CaV1.3b currents expressed in human embryonic kidney (HEK) 293 cells by stabilizing channels in a closed state. Inhibition occurs regardless of which CaVβ subunit is coexpressed; however, the magnitude of inhibition produced and whether kinetic changes occur after AA depend on the CaVβ subunit. Physiologically, the tissue-specific expression of different CaVβ subunits is predicted to affect the response of L-channels after the stimulation of certain GPCRs or exposure to high levels of free AA, such as during ischemia.
The HEK cell line, stably transfected with the M1 muscarinic receptor (HEK M1; provided by E. Liman, University of Southern California, Los Angeles, CA, and originally transfected by Peralta et al., 1988) was propagated at 37°C with 5% CO2 in Dulbecco's MEM (DMEM)/F12 supplemented with 10% FBS, 1% G418, 0.1% gentamicin, and 1% HT supplement (Invitrogen). Cells were passaged once they became 80% confluent.
HEK M1 cells were placed in 12-well plates at ~60–80% confluence and transfected with a 1:2:1 molar ratio of CaV1.3b, CaVβ, and α2δ-1 subunits (Xu and Lipscombe, 2001) using Lipofectamine PLUS (Invitrogen) according to the manufacturer's instructions. Constructs for CaV1.3b (+exon11, Δexon32, and +exon42a; GenBank accession no. AF370009), CaVβ3 (no. M88751), and α2δ-1 (no. AF286488) were provided by D. Lipscombe (Brown University, Providence, RI). Constructs for CaVβ1b (GenBank accession no. X61394), CaVβ2a (no. M80545), and CaVβ4 (no. L02315) were provided by E. Perez-Reyes (University of Virginia, Charlottesville, VA). CaVβ2a C3,4S was constructed by M. Hosey's lab (Chien et al., 1996) and supplied by A. Fox (University of Chicago, Chicago, IL). The CaVβ2e (AF423192) construct was provided by H.M. Colecraft (Columbia University, New York, NY), and the CaVβ2aC3,4S-CD8 construct was provided by P. Charnet (Centre National de la Recherche Scientifique, Paris, France). For all transfections, 0.5 µg DNA was used per well of a 12-well plate. Green fluorescent protein cDNA was <10% of the total cDNA used. Cells were washed with DMEM, and the DNA mixture was added dropwise to each well, gently rocked, and then incubated for 1 h at 37°C in a 5% CO2 incubator. Supplemented media, without antibiotics, was returned to the cells to bring the volume up to 1 ml (normal growth medium volume). After 2 h, cells were washed with full media. Cells were washed a final time 2 h later, and 10 mM MgSO4 was added. Cells were transferred 24–72 h after transfection using 2 mM EDTA in 1× PBS to poly-l-lysine–coated coverslips; recording began 1 h after transfer.
Electrodes were pulled from borosilicate glass capillary tubes (Drummond Scientific Company). Each electrode tip was fire polished to a diameter of ~1 µm to give the pipette a resistance of 2–4 MΩ. Whole cell currents were recorded at room temperature (20–24°C) with an Axon 200B patch clamp amplifier (MDS Analytical Technologies). Currents were filtered at 5 or 20 kHz and digitized at five times the filter cut-off frequency of the four-pole Bessel filter. Data were acquired using Signal software (Cambridge Electronic Design) and stored for later analysis on a personal computer. For time course experiments, a holding potential of −60 or −90 mV was used, and currents were measured at a test potential of −10 mV for 40 ms at 5-s intervals. For holding potential–dependent inactivation experiments, a holding potential of −60 mV was used and the “conditioning” potential varied from −100 to +50 mV for 2.2 s, immediately followed by a test potential of −10 mV for 40 ms.
The pipette solution consisted of (in mM): 125 Cs-aspartate, 10 HEPES, 0.1 BAPTA, 5 MgCl2, 4 ATP, and 0.4 GTP brought to pH 7.50 with CsOH. High resistance seals were established in Mg2+ Tyrode's consisting of (in mM): 5 MgCl2, 145 NaCl, 5.4 KCl, and 10 HEPES brought to pH 7.50 with NaOH. Once a seal was established and the membrane ruptured, the Tyrode's solution was exchanged for an external bath solution consisting of (in mM): 125 NMG-aspartate, 20 Ba2+, and 10 HEPES brought to pH 7.50 with CsOH. AA (all-cis 5,8,11,14-eicosatetraynoic acid; NuCheck Prep), oleic acid (NuCheck Prep), and ETYA (BioMol) were dissolved in 100% EtOH and stored under nitrogen as stock solutions at −70°C. Working dilutions were made fresh daily by diluting stock solutions at least 1:1,000 in external bath solution. A final concentration of 10 µM AA was used unless otherwise noted. Palmitic acid (PA) was prepared daily as a 50-mM stock solution in EtOH and diluted to a final concentration of 10 µM in external bath solution. BSA (fraction V, heat shock, fatty acid ultra-free; Roche) was diluted 1 mg/ml directly in bath solution. All chemicals were purchased from Sigma-Aldrich unless otherwise noted.
Linear leak and capacitive currents were subtracted and maximal inward current amplitudes were measured after the onset of the test pulse (peak current). Percent inhibition was calculated as
where ICTL is the average of five peak inward current values before the application of test material, and IINHIB is the average of five peak inward current values 1 min after the application of test material (unless otherwise noted). Difference current was measured as
Time to peak (TTP) was measured using a trough-seeking function in Signal within the test pulse duration. Current remaining was measured from an average of five individual sweeps per recording using the equation:
where IR is the current remaining at the end of a 40-ms (r40) or 2.2-s (r2200) test pulse, IEND is the value at the end of the test pulse, and IPEAK is the maximum inward current measured during the test pulse. Conductance was calculated from a modified Ohm's Law equation:
where IPEAK is the peak current at each test potential, Vm is the test potential, and Vrev is the reversal potential. Relative conductance (G/Gmax)-voltage (Vm) curves were plotted and fit using the Boltzmann equation:
where Gmax is the maximal conductance, Gmin is the minimal conductance, Vm is the test potential, Vm1/2 is the voltage at half-maximal conductance, and k is the slope factor. Holding potential–dependent inactivation was plotted as normalized current during the test pulse against the conditioning pulse voltage. The inactivation curve was fit using the Boltzmann equation:
where Vm is the conditioning pulse and Vinact1/2 is the voltage where half the maximal number of channels has inactivated. The rate of inactivation was measured by fitting the current traces to a biexponential function yielding time constants for a fast (τfast) and slow (τslow) component:
where Afast and Aslow are the current amplitudes with the respective time constants, and A0 is any remaining current. The time courses for recoveries from fast and slow inactivation were fit to the same biexponential function yielding time constants for τfast and τslow components.
Data are presented as mean values ± SEM. Data were analyzed statistically using either a one-way ANOVA followed by a Tukey multiple comparison post-hoc test or a two-tailed paired or unpaired Student's t test. Statistical significance was set at P ≤ 0.05. Analysis programs included Signal (Cambridge Electronic Design), Excel (Microsoft), and Origin (OriginLab).
Fig. S1 shows that 10 μM oleic acid had no significant effect on β2a- and β3-containing channels. Fig. S2 shows that no significant correlations among control current amplitude, control r40, and percent inhibition by 10 μM AA were found for CaV1.3 currents coexpressed with any of the β-subunits. Correlations were calculated for recordings using a holding potential of −90 or −60 mV. Fig. S3 shows that rate of activation is faster in the presence of AA when fitting current traces to the equation: . CaV1.3 currents, regardless of the CaVβ subunit coexpressed, were best fitted to one exponential. Figs. S1–S3 are available at http://www.jgp.org/cgi/content/full/jgp.200810047/DC1.
To determine if AA inhibits CaV1.3b activity, peak current amplitudes were measured from −90 mV to a test pulse of −10 mV every 5 s before and after the application of 10 µM AA to cells transiently transfected with CaV1.3b, β3, α2δ-1, and green fluorescent protein. AA produced a 51 ± 8% inhibition of current after 1 min (n = 7; P < 0.001). We then tested whether inhibition occurred in a concentration-dependent manner and found that the magnitude of inhibition increased with increasing AA concentration (Fig. 1, A and B). To stay well below the critical micelle concentration, concentrations >10 µM AA were not tested. In measuring current inhibition, we noticed an apparent acceleration in activation kinetics that was most pronounced with 10 µM AA (Fig. 1 B) and was apparent when current traces were normalized to the end of the 40-ms test pulse (Fig. 1 C). The increased rate of activation (Fig. 1 D, top), measured as TTP, maximally decreased after 1 min, whereas current inhibition (Fig. 1 D, bottom) developed over the span of 3 min. The application of 1 mg/ml BSA, which binds free fatty acid, reversed both the kinetic changes and current inhibition, indicating that channels were specifically modulated as opposed to nonspecific rundown of channel activity over time.
The change in TTP was unanticipated because native L-current in SCG neurons exhibits no apparent kinetic change when inhibited by AA (Liu et al., 2001). Additionally, whole cell L-current in SCG neurons is non-inactivating as opposed to the relatively rapid inactivating current observed with unmodulated transfected CaV1.3b, β3 currents (Fig. 1 B, control [CTL]). Non-inactivating current kinetics are characteristic of channels associated with β2a, which is uniquely palmitoylated, tethering it to the membrane and conferring different inactivation properties (Chien et al., 1996). Therefore, we tested whether the AA-induced TTP change also occurs with β2a. Fig. 1 E (bottom) shows that 10 µM AA significantly inhibited current from cells transfected with CaV1.3b, α2δ, and β2a by 31 ± 7% after 1 min (n = 8; P < 0.001). However, no change in TTP was observed (Fig. 1 E, top). After 3 min, current was inhibited by 56 ± 15% (n = 5; P < 0.001), suggesting that currents from β2a-containing channels can be inhibited to a similar extent as β3-containing channels after 1 min. The application of BSA recovered inhibited current. These differences suggest that different CaVβ subunits uniquely tune CaV1.3b modulation by AA.
Four genes in the CaVβ family (β1, β2, β3, and β4) and several splice variants have been identified to date (Birnbaumer et al., 1998; Takahashi et al., 2004). To test the effects of other CaVβ family members on the modulation of CaV1.3b by AA, we compared representative current traces for CaV1.3b currents coexpressed with β1b, β2a, β3, or β4 in the absence (CTL) or presence of 10 µM AA (Fig. 2 A). After a 1-min exposure to AA, the magnitude of inhibition was 58 ± 6% for β1b (n = 7; P < 0.001) and 44 ± 4% for β4 (n = 6; P < 0.001) (Fig. 3 A). Thus, AA inhibits CaV1.3b regardless of the CaVβ subunit coexpressed, but the magnitude of inhibition is less for β2a and is not accompanied by kinetic changes. The finding that β2a-containing channels exhibit a profile of modulation identical to our studies of native L-current (Liu et al., 2001) suggests that CaV1.3 in SCG neurons primarily couples to β2a.
To visualize differences in kinetics, individual traces were normalized to the end of the 40-ms test pulse (Fig. 2 B). When the inhibited current was subtracted from CTL current, the difference current revealed a relatively non-inactivating shape for currents with each of the CaVβ subunits (Fig. 2 C). The loss of non-inactivating current after AA is similar to whole cell difference currents recorded in SCG neurons and is consistent with AA-stabilizing channels in closed or inactivated states, thus removing inhibited channels from contributing to the whole cell current. This result suggests that the current actually inhibited by AA inactivates very little, whereas the residual current displays more inactivation for β1b-, β3-, or β4-containing channels.
Single-channel analysis of L-current inhibition by AA revealed an increase in null sweeps and a decrease in open probability (Liu and Rittenhouse, 2000), suggesting that AA increases the number of channels dwelling in a non-conducting conformation: either closed or inactivated. Additionally, first latency, the time for a single channel to transition from closed to open in response to a test pulse, increases from ~30 to 90 ms (Liu and Rittenhouse, 2000), indicating that when an L-channel is affected by AA, the time to transition from closed to open increases. These results support a model where AA stabilizes a closed conformation rather than an inactivated conformation because inactivated channels will not respond to voltage until sufficient time has transpired for channels to recover.
Many kinetic models display closed Ca2+ channels in at least four closed conformations representing the movement of each voltage sensor from domains I–IV of the CaVα1 subunit in response to changes in membrane potential to the open conformation (Patil et al., 1998; Serrano et al., 1999). For example, at a holding potential of −90 mV, none of the voltage sensors have moved and channels reside primarily in a deep closed state (Marks and Jones, 1992); at a holding potential of −60 mV, the voltage sensors have moved in response to depolarization and channels reside in closed conformations closer to the open state. Therefore, we tested whether holding cells at −60 mV would alter the magnitude of CaV1.3b inhibition by AA compared with holding cells at −90 mV. AA suppressed CaV1.3b currents with β1b, β3, or β4 by 34 ± 6%, 26 ± 7%, or 26 ± 4%, respectively (Fig. 3 B), but with β2a, only by 12 ± 6% (n = 7–13; P < 0.001 for all CaVβ subunits compared with CTL; P < 0.05 for all CaVβ subunits comparing current inhibition from −90 to −60 mV). From −60 mV, TTP continued to be unaffected by AA application for channels with β2a and decreased for channels with β3 (Fig. 3 C). Overall, current inhibition by AA was greater for all CaVβ subunits recorded with a holding potential of −90 mV than with −60 mV, consistent with AA stabilizing a deep closed state (Liu and Rittenhouse, 2000).
Next, we examined whether channels need to enter the open conformation for AA inhibition of CaV1.3b currents. Here, we used a protocol with short 10-ms test pulses and no depolarization for 1 min in the absence or presence of AA (Fig. 3 D). The protocol was designed to maintain channels in a closed conformation; the shorter test pulse should minimize the amount of inactivation. Additionally, no depolarization for 1 min should bring channels into a resting, closed conformation. The 10-ms test pulse from −60 mV had no effect on the ability of AA to inhibit currents from β3-containing channels. In fact, the shorter 10-ms test pulse showed more inhibition after 1 min (35%; Fig. 3 E) than recordings with 40-ms test pulses (26%; Fig. 3, B and E). In the absence of test pulses, AA inhibited 34% of the current (n = 7; P < 0.001 compared with CTL current before the application of AA). Moreover, BSA reversed inhibition to CTL peak amplitudes (not depicted). Overall, these experiments suggest that CaV1.3b does not need to open to become inhibited by AA.
To address the specificity of AA inhibition and the possible role of AA metabolites, a nonhydrolyzable form of AA, ETYA, was tested for inhibition of CaV1.3b currents. From a holding potential of −60 mV, current was measured 1 min after the application of 30 µM ETYA. Fig. 4 A shows representative individual sweeps for each of the CaVβ subunits before or after ETYA. To visualize differences in kinetics, individual traces were normalized to the end of the 40-ms test pulse (Fig. 4 B). ETYA inhibited currents for β1b-, β2a-, β3-, or β4-containing channels 32 ± 8%, 14 ± 4%, 24 ± 5%, or 22 ± 6%, respectively (n = 5–8; P < 0.001 compared with CTL) (Fig. 4 C). ETYA inhibition did not significantly differ from AA inhibition from a holding potential of −60 mV shown in Fig. 3 B (β1b, P ≥ 0.89; β2a, P ≥ 0.81; β3, P ≥ 0.78; β4, P ≥ 0.59). To test whether the effects of ETYA and AA were additive, 30 µM ETYA was added for 1 min, 30 µM ETYA + 10 µM AA was added for another minute, and current inhibition was measured. Current was further inhibited by ETYA + AA for a total inhibition of 49 ± 7% (n = 4; P < 0.001 compared with CTL; P < 0.05 compared with ETYA alone) for CaV1.3b currents with β3. This magnitude of inhibition is similar to that measured by 10 µM AA alone after application for 2 min (not depicted). These results suggest that inhibition is the result of AA itself and not a metabolite. In contrast to polyunsaturated AA or ETYA, 10 µM oleic acid (C18:1), a monounsaturated fatty acid, had no significant effect on currents from channels containing β2a (n = 4; P ≥ 0.66) or β3 (n = 3; P ≥ 0.58) (Fig. S1), indicating that inhibition requires certain aspects of fatty acid structure.
To thoroughly analyze any changes in the overall closed to open transitions, we examined the effect of AA on peak current over a range of test potentials (−100 mV to +50 mV in 10-mV steps). In I-V plots, all currents, regardless of the CaVβ subunit expressed with CaV1.3b, activated at −50 mV, peaked at −10 mV, and reversed at +50 mV, similar to previous reports of CaV1.3 activating at low voltages (Koschak et al., 2001; Xu and Lipscombe, 2001). AA inhibited currents at all voltages regardless of the CaVβ subunit (Fig. 5 A). G-V plots revealed no shift in the activation curve by AA compared with CTL plots (Fig. 5 B), consistent with inhibited channels becoming unavailable to open (stabilized closed) and thus not contributing to the G-V relationship. Therefore, AA inhibition is not the result of altering the voltage sensitivity of CaV1.3b.
Because these initial measurements are consistent with a model where AA stabilizes a deep closed state, we next sought to understand the observed changes in activation and inactivation described in Figs. 1 and and2.2. We first quantified current remaining at the end of the 40-ms test pulse (r40), respectively, for CaV1.3b with each CaVβ subunit. For currents elicited from a holding potential of −90 to −10 mV, r40 (Fig. 6 A) significantly decreased after a 1-min exposure to AA for currents containing β1b, β3, or β4, but not β2a (n = 6–8; P < 0.05 for β1b, β3, and β4; P ≥ 0.48 for β2a). Although less inhibition by AA was observed at −10 mV from a holding potential of −60 mV, the decrease in r40 was still apparent for channels with β1b, β3, and β4; no kinetic changes were observed for channels with β2a (n = 7–13; P < 0.05 for β1b, β3, and β4; P ≥ 0.48 for β2a) (Fig. 6 B). BSA reversed both the kinetic change and current inhibition regardless of holding potential (not depicted). Additionally, we found no correlations between CTL current amplitude, CTL r40, or percentage of AA-induced inhibition of CaV1.3b with any CaVβ subunit (Fig. S2).
TTP was also measured across a range of test potentials from a holding potential of −60 mV. After AA application, TTP was generally faster for β1b-, β3-, or β4-containing channels across voltages (Fig. 6 C). In contrast, TTP for CaV1.3b with β2a, which is slower than with the other CaVβ subunits, was only faster in the presence of AA at −20 mV. To confirm these data, activation time constants were measured at multiple test potentials before and after 1 min of AA (Fig. S3). The activation of CaV1.3b was best fitted with one exponential (τACT) for all CaVβ subunits, similar to previous monoexponential fits of CaV1.3 with β3 (Koschak et al., 2001). The τACT for CaV1.3b, β2a currents was approximately threefold slower than that for the other CaVβ subunits. Overall, AA decreased the mean τACT across voltages for CaV1.3b currents with all CaVβ subunits, consistent with the decrease in TTP after AA. TTP kinetic changes were also apparent after AA with the shorter 10-ms test pulses (not depicted). These findings suggest that although the kinetic changes observed with β1b-, β3-, or β4-containing channels occur with inhibition over the range of voltages tested, the TTP change produced by AA is not the result of a shift in voltage dependence of activation.
Kinetic changes produced by AA were also mimicked by ETYA. When comparing CTL currents to currents exposed to ETYA, TTP significantly decreased for all CaVβ subunits (n = 5–8; P < 0.05) (Fig. 6 D). Significant decreases in r40 were also observed for CaV1.3b with β1b, β3, and β4, but not with β2a (Fig. 6 E). For CaV1.3 with β3, TTP and r40 significantly decreased for ETYA + AA compared with CTL, but it did not significantly differ from ETYA alone. Thus, in addition to inhibition, the kinetic changes observed after AA application are not produced by an AA metabolite.
At least two explanations may account for the kinetic changes and the decrease in current amplitude after AA. In the first scenario, after exposure to AA, all of the channels shift to more reluctant gating where a greater voltage step is required to open channels. If this is the case, the activation curve will exhibit a rightward shift to more positive voltages after AA. Moreover, if AA shifts all channels toward reluctant gating, we would expect to observe a slower TTP. This scenario seems unlikely because no shift in the G-V curve occurs (Fig. 5 B), and TTP is faster at −10 mV in the presence of AA compared with CTL currents, despite our previous findings that AA increases dwell times in closed conformations and increases first latency to 90 ms (Liu and Rittenhouse, 2000). Because the test pulse duration for the whole cell current recordings was 40 msec, the inhibited channels are unlikely to open.
In a second scenario, at least two populations of L-channels are present under CTL conditions: one population of channels is reluctant to open (reluctant closed channel conformation [RC]), whereas another population is willing to open (willing closed channel conformation [WC]), with the RC transition time to the open channel conformation (O) slower than the WC→O transition time (Jones and Elmslie, 1997). This description of reluctant gating has been described previously for CaV2 channels modulated by G proteins (Bean, 1989) or by phosphatidylinositol-4,5-bisphosphate (PIP2) binding (Wu et al., 2002). CTL TTP can be expressed as the sum of RC→O plus WC→O transition times. After AA, a greater percentage of channels reside in the RC conformation longer and, therefore, will not open during the 40-ms test pulse due to the observed first latency increase to 90 ms (Liu and Rittenhouse, 2000). Consequently, the residual current displays only the faster WC→O transition. This model is consistent with the decreased TTP after AA. Similarly, RC channels show slower inactivation. Therefore, if RC channels are inhibited and WC channels remain active, an apparent increase in inactivation would occur. This model fits the observed kinetic changes in TTP and r40 after AA.
This simple model predicts that AA would only need to stabilize one or more deep closed states to confer the observed changes in channel gating; no change in fast or slow inactivation need occur. The above data rule out changes in fast inactivation. Therefore, we tested whether AA changes slower forms of inactivation. To test AA's effect on holding potential–dependent inactivation, a protocol was used in which channels were exposed to a series of “conditioning pulses” ranging from −100 to +50 mV (in 10-mV steps) for 2.2 s, and then repolarized to the test potential of −10 mV for 40 ms. Representative current traces for CaV1.3 with β2a (Fig. 7, A and C) or β3 (Fig. 7, B and D) are shown before (CTL) or after exposure to AA for 1 min. The long, 2.2-s conditioning pulse highlights the striking differences in inactivation kinetics between β2a- and β3-containing channels. Inactivation currents for CaV1.3 with β1b, β3, or β4 were best fit with two exponentials (τINACT-fast and τINACT-slow), whereas CaV1.3 β2a currents were best fit with one exponential (τINACT-slow). Fits of inactivation are shown overlying the representative traces (Fig. 7, C and D). Fig. 8 (A and B) summarizes the inactivation time constants for voltages from −20 to +40 mV. AA significantly decreased τINACT-fast at positive test potentials (0 to +40 mV) for β1b-, β3-, or β4-containing currents. As a general trend, AA did not effect τINACT-slow for β1b-, β3-, or β4-containing currents (Fig. 8 B), although AA decreased τINACT-slow for currents with β1b at +20 mV, β2a at 0 and +20 mV, β3 at +20 to +40 mV, and β4 at 0 mV (n = 5–7; P < 0.05). Overall, CaV1.3 currents with β1b, β3, or β4 appear to fast inactivate more rapidly in the presence of AA, consistent with the r40 data.
To measure the voltage sensitivity of holding potential–dependent inactivation, currents were normalized to peak inward current during the test pulse and plotted against the conditioning test potential (Fig. 8 C). The CTL inactivation curve for CaV1.3 currents with β2a was shifted ~10 mV more positive than the other CaVβ subunits. From fits to the Boltzmann equation (Fig. 8 C), we found that AA had no effect on the Vinact1/2 of the CaV1.3 currents coexpressed with β1b (CTL, −41.1 ± 1.2 mV; AA, −44.1 ± 0.8 mV), β2a (CTL, −29.8 ± 0.7 mV; AA, −30.4 ± 0.8 mV), β3 (CTL, −38.5 ± 0.8 mV; AA, −41.9 ± 1.1 mV), or β4 (CTL, −38.5 ± 0.8 mV; AA, −40.8 ± 0.4 mV). Similar values for CTL CaV1.3 β3 Vinact1/2 have been reported (Scholze et al., 2001). These results show that AA has no effect on the voltage dependence of holding potential–dependent inactivation for CaV1.3 currents with any CaVβ subunit.
Closer inspection of the CaV1.3 inactivation curves for β1b-, β3-, or β4-containing channels revealed that inactivation occurred at negative test potentials before channels began to open, suggesting that channels were inactivating from a closed state similar to T- (Frazier et al., 2001) and N-type Ca2+ channels (Patil et al., 1998). For example, at the conditioning pulse of −60 mV, ~10–15% of the channels were inactivated even though the channels do not open at this voltage. At the conditioning pulse of −90 mV, only 5% of the channels were inactivated. The slope between −100 and −60 mV, before channels begin to activate, does not appear to be an artifact or an accumulation of inactivation because testing longer intervals between the final test pulse and the following conditioning pulse had no effect on the voltage dependence of the inactivation curves with and without AA (not depicted). Regardless, this observed inactivation profile does not change in the presence of AA.
Although the voltage sensitivity did not shift after AA, significantly fewer β2a-containing channels were available to open per voltage (−10 to +50 mV) in the presence of AA (Fig. 8 C). This finding correlates with the amount of current remaining at the end the 2.2-s conditioning pulse (r2200). Here, AA had no effect on the r2200 for β1b-, β3-, or β4-containing channels because >90% of CTL and inhibited current had already undergone fast inactivation. However, AA decreased the r2200 for β2a-containing channels (Fig. 8 A, inset in box). Therefore, over a longer time scale, currents with β2a subunits also displayed more inactivation in the presence of AA. By decreasing the number of channels available to open during a given test pulse, fast inactivation of β1b-, β3-, or β4-containing channels was accentuated, whereas holding potential–dependent inactivation of β2a-containing channels was accentuated by AA. Thus, AA may act via a common mechanism of stabilizing deep closed states independently of which CaVβ subunit is coexpressed with CaV1.3b.
Because an apparent increase in channels residing in closed conformations could be due to an increase in recovery time from the inactivated state, we next examined recovery from inactivation using two separate protocols. First, peak current was measured every 5 s after the inactivation protocol in the absence and presence of AA (Fig. 9 A). Current recoveries were fit to a single exponential time constant and did not significantly differ between CTL and AA for CaV1.3 with any CaVβ subunit (Fig. 9 B). Second, in another test for changes in recovery from inactivation, the test pulse was placed at various time points (Δt) after a 1-s conditioning pulse (Fig. 9 C). In this protocol, we chose a 1-s conditioning pulse to +20 mV based on where maximal holding potential inactivation occurred (Fig. 8 C, β3). Recovery from inactivation was best fitted with two time constants: τ1 = 291 ms and τ2 = 13.6 s. Recovery from inactivation in the presence of AA also was best fitted with two time constants: τ1 = 371 ms and τ2 = 17.2 s (Fig. 9 D). This experiment allowed for the measurement of recovery from fast inactivation, which was overlooked in fitting the recovery from the time courses during which currents were evoked every 5 s (Fig. 9, A and B). Consistent with the first protocol, the time constants were not significantly different between CTL and AA (Fig. 9 D). However, the fractional amplitudes A1 and A2 associated with the recovery time constants revealed an important difference. The CTL values for A1 and A2 were 0.52 and 0.34, respectively, whereas the values for AA were 0.67 and 0.18, respectively. The finding suggests that in the presence of AA, there are more channels recovering from fast inactivation and fewer channels recovering from slow inactivation. Therefore, AA does not change the rate constant between inactivated and closed conformations, but rather alters the number of channels recovering from fast and holding potential–dependent inactivation.
β2a uniquely stands apart from other CaVβ subunits because it contains two palmitoyl groups at cysteine residues in the N terminus (Chien et al., 1996), conferring unique non-inactivating properties to CaV current (Qin et al., 1998). To test whether the non-inactivating properties of β2a-containing channels are responsible for less AA inhibition, we tested another β2 splice variant, β2e, which is not palmitoylated, yet still has non-inactivating properties. After exposure to AA for 1 min, β2e channels were inhibited 24 ± 8% (n = 6; P ≥ 0.26 compared with β2a) (Fig. 10 A). Even though percent inhibition by AA approximately doubled for β2e compared with β2a, the difference was not significant. Therefore, we tested a β2a subunit in which amino acid residues 3 and 4 have been mutated from cysteines to serines (β2a C3,4S) resulting in a β2a subunit that cannot be palmitoylated (Restituito et al., 2000). After exposure to AA for 1 min, β2a C3,4S channels were inhibited 32 ± 5% (n = 6; P < 0.05 compared with β2a) (Fig. 10 A). To examine whether the loss of the palmitoyl groups or the loss of non-inactivating properties resulted in more inhibition by AA, we tested a β2a C3,4S subunit with a CD8 sequence attached. CD8 is a transmembrane protein, and thus attaching CD8 to the N terminus of β2a C3,4S restores the non-inactivating properties (Restituito et al., 2000). After exposure to AA for 1 min, β2a C3,4S-CD8 channels were inhibited 44 ± 9% (n = 9; P < 0.05 compared with β2a) (Fig.10 A). Fig. 10 B shows the r40 for currents with β2e, β2a C3,4S, and β2a C3,4S-CD8 before and after AA application. Representative sweeps for each condition are displayed in Fig. 10 C. β2e and β2a C3,4S-CD8 currents both display non-inactivating properties yet are inhibited more by AA than β2a channels, suggesting that the palmitoyl groups of β2a, and not the non-inactivating properties, are responsible for decreased AA inhibition of CaV1.3b.
A compelling idea is that palmitoyl groups of β2a may compete for a “fatty acid” or “AA” binding site on CaV1.3. Therefore, we tested if excess PA would block AA inhibition. A 1-min exposure to 10 µM PA inhibited β2a C3,4S currents by 8 ± 4% (n = 3; P < 0.05 compared with 1 min exposure of β2a C3,4S currents to 10 µM AA). Cells were then preincubated in 10 µM PA in Tyrode's solution before establishing a seal. After breakthrough, bath solution containing 10 µM PA was washed in and a baseline current was measured before adding 10 µM PA + 10 µM AA. After a 1-min exposure to 10 µM AA, currents were inhibited 11 ± 7% (n = 3; P < 0.05 compared with 1 min exposure of β2a C3,4S currents to 10 µM AA). Fig. 11 A is a summary bar graph of mean inhibition by AA. Fig. 11 B shows representative sweeps for each condition. A summary bar graph for current remaining for each condition is shown in Fig. 11 C. Overall, these results suggest that the palmitoyl groups of β2a or excess PA may compete with AA for an inhibitory site on CaV1.3b.
Here, we show that AA inhibits CaV1.3b, a short form of CaV1.3 with a truncated C terminus, ruling out its importance in modulation by AA. Because we had previously characterized AA's effects on native L-current from SCG neurons (Liu and Rittenhouse, 2000; Liu et al., 2001, 2006), we used CaV1.3b, which was cloned from SCG neurons (Xu and Lipscombe, 2001), in these studies. Using ETYA, a nonmetabolizable form of a fatty acid similar to AA, we show that inhibition and kinetic changes of CaV1.3 are due to AA itself, and not a metabolite (Figs. 4 and 6, D and E). Current inhibition occurs regardless of the CaVβ subunit coexpressed. However, the magnitude of inhibition after 1 min by either AA or ETYA was less with coexpression of β2a than β1b, β3, or β4. In addition, AA decreased r40, a measure of fast inactivation, and TTP of channels containing β1b, β3, or β4 (Figs. 1, ,2,2, and and6).6). In contrast, β2a-containing channels showed no differences in TTP after exposure to AA. These complex changes are manifested in the presence of AA due to the influence of different CaVβ subunits coexpressed with CaV1.3b on gating kinetics. Because the kinetic changes do not correlate with percent inhibition by AA over time (Figs. 1 D and 3 C), AA may act at multiple sites on L-channels.
AA inhibits native L-current in a variety of systems: cardiac muscle (Petit-Jacques and Hartzell, 1996; Xiao et al., 1997; Liu, 2007), skeletal muscle T-tubule membranes (Oz et al., 2005), smooth muscle (Shimada and Somlyo, 1992), SCG (Liu and Rittenhouse, 2000; Liu et al., 2001), photoreceptors (Vellani et al., 2000), and retinal glial cells (Bringmann et al., 2001). No consensus of mechanism exists as to how AA inhibits L-channel activity, whether directly, through AA metabolites, or via AA activating a phosphatase. Varying effects of AA on L-current may be due to expression of different L-CaVα1 subunits. For example, ventricular myocytes and smooth muscle cells primarily express CaV1.2, whereas SCG neurons, atrial myocytes, and cochlear hair cells express CaV1.3 (Lin et al., 1996; Platzer et al., 2000; Mangoni et al., 2003; Catterall et al., 2005). Additionally, CaV1.2 and CaV1.3 undergo alternative splicing (Hell et al., 1993; Safa et al., 2001), which may contribute to the differences in L-current kinetics observed with varying cell types. Notably, the lack of AA-induced kinetic changes in cardiac myocytes and SCG neurons may be due to high levels of β2a subunit mRNA found in these cells (Lin et al., 1996; Birnbaumer et al., 1998).
In this study of whole cell recombinant CaV1.3b L-current, we found changes induced by AA complementary to our previous single-channel data. Analysis of unitary L-channel gating in SCG neurons revealed that AA-induced inhibition had no effect on mean open time or on unitary current amplitude (Liu and Rittenhouse, 2000). Consistent with this finding, the results from three experiments indicate that the open conformation of CaV1.3b is not necessary for inhibition by AA. First, test pulse length did not affect the magnitude of current inhibition by AA (Fig. 3 E). In fact, reducing the test pulse length increased the amount of AA-induced current inhibition. Second, β2a-containing currents, which do not exhibit fast inactivation and therefore gate open and closed longer, exhibited less inhibition (Figs. 1–5). Third, CaV1.3b channels exhibited robust inhibition when held closed for 1 min during AA application and then allowed to open (Fig. 3 E). Thus, AA does not confer inhibition by binding to the open state.
Single-channel analysis of L-current in SCG neurons also revealed that AA decreased channel open probability by increasing the number of null sweeps. Moreover, when channels were active, AA decreased the number of openings and increased first latency and mean closed time (Liu and Rittenhouse, 2000). Together, the changes in the unitary activity indicate that AA stabilizes L-channels in one or more nonconducting states: either closed or inactivated conformations.
We hypothesize that AA decreases channel availability by increasing the number of L-channels stabilized in a deep closed state, as opposed to an inactivated state, based on several findings. First, we observe more AA inhibition from channels held at a membrane potential of −90 mV than at −60 mV (Fig. 3, A and B); at a membrane potential of −90 mV, channels undergo less closed state inactivation than at −60 mV (Fig. 8 C, β1b, β3, and β4). Second, the voltage sensitivity of holding potential–dependent inactivation does not change with AA (Fig. 8 C). Third, when measuring both fast and slow recovery for β3-containing channels, we observe no change in recovery from inactivation in the presence of AA (Fig. 9), suggesting that the transition rate from inactivated to closed conformations remains unchanged. Moreover, inactivated channels regardless of voltage will not respond to positive test pulses until given enough time at a negative holding potential to recover. On the other hand, stabilizing a deep closed state would require more depolarized voltage and, hence, more time to transition the channel to an open state, consistent with increases in first latency (Liu and Rittenhouse, 2000). The first latency of 30 ms fits well with the notion that L-channels in SCG neurons couple to β2a. Liu and Rittenhouse (2000) showed that in the presence of AA, this time increases to 90 ms. In these single-channel experiments, the effect of AA on a single channel's activity is directly monitored. Here, we hypothesize that when measuring whole cell current in the presence of AA, the observed current arises from channels not inhibited by AA.
Our hypothesis that AA modulates channels in deep, closed conformations, but not channels that are in a conformation near to the open state, is reminiscent of the two populations of channels described as willing and reluctant. Historically, willing-reluctant gating terminology has been used to describe gating that occurs when G proteins bind to high voltage–activated Ca2+ (CaV2) channels (Bean, 1989). G protein Gβγ subunits can bind CaV2 α1 subunits and inhibit CaV2 current by stabilizing “reluctant gating” conformations. This inhibition can be relieved by a strong depolarizing prepulse (typically to +120 mV), releasing the Gβγ subunit and allowing the channel to occupy “willing gating” conformations. Although CTL CaV1.3 currents display facilitation after a strong depolarizing prepulse, the mechanism is not mediated by Gβγ binding CaV1.3 (Bell et al., 2001; Safa et al., 2001) and remains unknown. Determining whether AA promotes facilitation under certain conditions as well as inhibition will require further detailed experimentation.
More recently, the willing-reluctant terminology has been used to describe how PIP2 increases channel availability of the CaV2 family (Wu et al., 2002; Gamper et al., 2004). PIP2 has been hypothesized to act at two unidentified sites termed the “S” and “R” sites on CaV2 channels (Wu et al., 2002). When PIP2 binds to the high affinity S site, channel activity is stabilized and whole cell currents are protected from rundown. When PIP2 binds to the lower affinity R site, channels open more slowly during weak depolarizations, similar to the reluctant gating observed with G proteins (Wu et al., 2002; Michailidis et al., 2007). Exogenous application of AA appears to exert the opposite effects of PIP2's dual actions on CaV2.2 channels where AA decreases current magnitude but increases the activation and inactivation kinetics (unpublished data). These two actions of AA are distinguishable both biophysically and pharmacologically (Barrett et al., 2001; Liu et al., 2001). Because AA is a product of PIP2 breakdown by several phospholipases, these findings raise an interesting possibility that AA also acts at two analogous sites on CaV1.3 to antagonize PIP2's putative actions on L-current (Michailidis et al., 2007). A similar bidirectional relationship between PIP2 and AA has been documented for Kir and KV channels in which PIP2 and AA are hypothesized to act at distinct sites (Oliver et al., 2004; Tucker and Baukrowitz, 2008). Whether exogenously applied AA competitively displaces PIP2 or acts directly or indirectly to alter a distinct site on CaV1.3 channels to elicit inhibition has not yet been determined; however, free AA interacting at the same location(s) of PIP2's fatty acid tail(s) to confer inhibition is an appealing idea in its simplicity.
In addition to current inhibition, AA decreases TTP and the τACT becomes faster. We present the following scenario to explain our interpretation of this result. At rest, channels reside in a series of closed conformations whereby τACT describes the rate of channels transitioning from different closed conformations to the open conformation: RC + WC→O. Under CTL conditions, RC channels may have PIP2 bound to the R site as well as the S site (Wu et al., 2002), resulting in channels available to open but at a slower rate of transition to the open conformation than WC channels. However, the overall rate will be the average of RC + WC channels and is dependent on the proportion of channels occupying each closed conformation. If AA stabilizes a deep, closed conformation, RC channels, which transition through the closed states more slowly than WC channels, would be more susceptible to inhibition (Fig. 12 A). Thus, AA does not cause reluctant or willing gating, but rather AA is more likely to act on reluctant gating channels.
If AA selectively stabilizes RC channels in a closed state, making them unlikely to open during a test pulse, only the WC channels, with the faster activation rate, will open and determine the whole cell current kinetics. The decreases in TTP (i.e., peak inward current occurs faster) after exposure to AA is consistent with this model. Notably, in experiments with β2a-containing channels, TTP changes are not observed with 40-ms test pulses as this time frame is not long enough to account for the changes in activation; however, TTP changes are observed for β2a in the 2.2-s pulses that are long enough to account for first latency changes from 30 to 90 ms observed in SCG neurons. In several experiments, the membrane potential was held at −60 mV, near the threshold for opening (−50 mV) for CaV1.3 (Xu and Lipscombe, 2001). At this voltage, more channels would be predicted to reside in conformations closer to the open state. We observed less inhibition at −60 mV than with a holding potential of −90 mV, at which a greater percentage of channels would reside in deeper closed conformations (Fig. 12 B). Thus, in the presence of AA, the observed smaller current with a faster τACT arises from willing channels residing closer to the open state and “above” the closed state(s) stabilized by AA. It will be interesting to confirm these kinetic changes produced by AA for each of the subunit combinations in future studies of single-channel gating.
To address whether AA stabilizes inactivated as well as closed conformations, we measured whole cell CaV1.3 currents under various inactivation protocols in the absence and presence of AA. Although r40 decreased for β1b-, β3-, and β4-containing channels, the difference current was relatively non-inactivating. Because CaV1.3, β2a currents only undergo slow inactivation, the decrease in r2200 revealed that there is a common mechanism for inhibition by AA that occurs regardless of the β subunit that is coexpressed. AA may preferentially inhibit the RC population of channels, and thus for β2a-containing channels, the contribution of reluctant gating channels undergoing slow inactivation is decreased. Therefore, the percentage of WC channels, which remain available to open and available to undergo inactivation, is greater in the normalized holding potential–dependent inactivation curves. These findings suggest that when CaV1.3b exhibits willing gating, L-channels will be less susceptible to inhibition by AA. Taken together, AA decreases the contribution of reluctant gating channels to the whole cell current, resulting in an increased proportion of the remaining current displaying inactivation kinetics of willing channels. Moreover, a shift in percentage of willing versus reluctant channels is manifested in the current kinetics in a CaVβ subunit–dependent manner after AA; fast inactivation increases for β1b-, β3-, and β4-containing channels, whereas holding potential–dependent inactivation increases for β2a-containing channels.
Why is less inhibition of CaV1.3b current observed after 1 min with AA for channels coexpressed with β2a? One possibility is that the site of inhibition by AA occurs on the CaVα1 subunit. In this case, the palmitoylated β2a may alter the conformation of the channel to noncompetitively regulate the availability of an AA binding pocket. Second, residues in the S6 transmembrane domains affect inactivation, suggesting an overall change in the pore structure (Stotz et al., 2004), a concept that has been compared with pore collapse or C-type inactivation describing slow inactivation in Na+ and K+ channels (Lopez et al., 1994; Ogielska et al., 1995; Vilin et al., 1999). Channels coassembled with β2a have a higher open probability than β1b or β3 (Colecraft et al., 2002) and would provide greater access for AA to contact these pore residues, promoting greater holding potential–dependent inactivation at more depolarized voltages.
Indeed, AA, or molecules with AA moieties in their structure including anandamide, can occupy the binding site for L-channel agonists and antagonists, such as dihydropyridines (DHPs) (Shimasue et al., 1996; Jarrahian and Hillard, 1997). DHPs interact with several critical amino acid residues in the pore of CaV1.2 and CaV1.3 (Sinnegger et al., 1997; Wappl et al., 2001; Liu et al., 2003; Yamaguchi et al., 2003). Thus, pore residues overlapping with the DHP site may constitute a site for AA interaction. However, if AA interacted with pore residues, coexpression of CaV1.3b with β2a should increase rather than decrease the magnitude of CaV1.3 current inhibition by AA. Alternatively, slow open-channel pore block by AA is a third possible mechanism of inhibition that would explain an increase in the rate of inactivation as well as a decreased TTP. This possibility seems unlikely because AA inhibits channels held closed in the absence of test potentials. Regardless, a role for CaVβ subunits modulating the AA site, whether by occupying a DHP site in the pore or by pore block, is plausible because these accessory subunits can fine-tune the affinity of L-channel blockers (Hering, 2002).
A fourth possibility is that two sites of AA action could exist, and either the palmitoyl group(s) or the inability of β2a to fast inactivate prevents AA interaction with CaV1.3. In fact, the time course of inhibition of CaV1.3b, β2a currents appears slower than with the other CaVβ subunits (Fig. 1 E, top), suggesting that palmitoylated β2a may antagonize AA binding. Our findings suggest that the palmitoyl groups of β2a either directly or indirectly impede AA's ability to inhibit channel activity (Figs. 10 and and11).11). We propose a model whereby two AA modulatory or fatty acid sites exist on CaV1.3b. We have included two sites in our model based on a previous determination that AA inhibited T-channels with a Hill coefficient of 1.6 (Talavera et al., 2004). T-channel expression requires no other accessory subunit, suggesting that at least two sites of interaction exist on CaVα subunits. When CaV1.3b colocalizes with β1b, β3, or β4, both sites are available. When CaV1.3b colocalizes with β2a, the palmitoyl groups interfere or occupy at least one of the sites, resulting in less inhibition by AA (Fig. 12 C).
CaV1.3 channels colocalize with large-conductance Ca2+-activated K+ channels (BK channels) in the brain (Grunnet and Kaufmann, 2004). Increases in AA levels may directly affect Ca2+-activated K+ channel activity, resulting in an enhancement of K+ current (Denson et al., 2000; Clarke et al., 2002; Sun et al., 2007). The influence of AA on Ca2+ influx and K+ efflux, as well as several other ion channels (Meves, 2008), could profoundly alter membrane excitability. Additionally, proteins with two adjacent palmitoyl groups, such as β2a, may localize to cholesterol-rich microdomains (Pike, 2003), which may in turn alter CaV1.3 susceptibility to modulation by AA.
In certain pathophysiological conditions, such as ischemia and stroke, concentrations of AA, comparable to those used in the present study, are released in the brain (Lipton, 1999). Brain regions with high expression of β2a subunits (Birnbaumer et al., 1998; McEnery et al., 1998) may be less affected by the cytotoxic effects of AA release downstream of neurotransmitter activation because β2a-containing CaV1.3b currents show less inhibition. Conversely, AA could increase cell survival by inhibiting Ca2+ influx because large sustained influxes of Ca2+, such as that of the non-inactivating β2a current, can be cytotoxic (Lipton, 1999). Similarly, currents from β1b-, β3-, or β4-containing CaV1.3b channels show more AA inhibition; this overall depression of Ca2+ influx may decrease excitotoxicity. Whether AA acts indirectly or by binding to CaV1.3b or the CaVβ subunit remains unanswered. Because AA is hydrophobic, if inhibition is due to a direct binding effect, CaV1.3 with its 24 membrane–spanning segments seems the more likely candidate as opposed to the cytosolic CaVβ subunit. Future studies involving mutational analyses of CaVα or the CaVβ subunits should provide these answers.
The authors would like thank Liwang Liu, John Heneghan, and Tora Mitra-Ganguli for their assistance and discussions; TzuFeng Chung for maintaining the HEK M1 cell line; and José Lemos, William Kobertz, Haley Melikian, Elizabeth Luna, and Edward Perez-Reyes for thoughtful critique of the manuscript.
This work was supported by National Institutes of Health grants (NS34195 and NS07366) and funding from UMASS Medical School.
Lawrence G. Palmer served as editor.
Abbreviations used in this paper: AA, arachidonic acid; CTL, control; DHP, dihydropyridine; ETYA, 5,8,11,14-eicosatetraynoic acid; GPCR, G protein–coupled receptor; HEK, human embryonic kidney; L-, L-type; M1, muscarinic M1 receptor; O, open channel conformation; PA, palmitic acid; PIP2, phosphatidylinositol-4,5-bisphosphate; RC, reluctant closed channel conformation; SCG, superior cervical ganglion; T-, T-type; TTP, time to peak; WC, willing closed channel conformation.