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Calcium ions (Ca2+) released from inositol trisphosphate (IP3)-sensitive intracellular stores may participate in both the transient and extended regulation of neuronal excitability in neocortical and hippocampal pyramidal neurons. IP3 receptor (IP3R) antagonists represent an important tool for dissociating these consequences of IP3 generation and IP3R-dependent internal Ca2+ release from the effects of other, concurrently stimulated second messenger signaling cascades and Ca2+ sources. In this study, we have described the actions of the IP3R and store-operated Ca2+ channel antagonist, 2-aminoethoxydiphenyl-borate (2-APB), on internal Ca2+ release and plasma membrane excitability in neocortical and hippocampal pyramidal neurons. Specifically, we found that a dose of 2-APB (100 µM) sufficient for attenuating or blocking IP3-mediated internal Ca2+ release also raised pyramidal neuron excitability. The 2-APB-dependent increase in excitability reversed upon washout and was characterized by an increase in input resistance, a decrease in the delay to action potential onset, an increase in the width of action potentials, a decrease in the magnitude of afterhyperpolarizations (AHPs), and an increase in the magnitude of post-spike afterdepolarizations (ADPs). From these observations, we conclude that 2-APB potently and reversibly increases neuronal excitability, likely via the inhibition of voltage- and Ca2+-dependent potassium (K+) conductances.
An important determinant of neuronal activity is the intracellular concentration of calcium ions ([Ca2+]i). Changes in [Ca2+]i are mediated by entry through voltage-gated Ca2+ channels (VGCCs) and ligand-gated channels (e.g., NMDA receptors), and by release of Ca2+ from intracellular stores such as the endoplasmic reticulum. There are two major classes of intracellular Ca2+ release channels in neurons, ryanodine receptor channels and IP3Rs . Ryanodine receptors are stimulated by cytosolic Ca2+, while IP3Rs are sensitive both to IP3 and to Ca2+. The dependence of IP3Rs on both IP3 and Ca2+ endows these receptors with the ability to respond differentially to a broad range of stimulus combinations and intensities [2,3]. For example, stimuli that mobilize threshold levels of IP3 may nonetheless evoke robust internal Ca2+ release when accompanied by a VGCC-mediated rise in [Ca2+]i [4–6]. This characteristic of IP3Rs complicates interpretation of the consequences of IP3-mobilizing stimuli, as it suggests that IP3R-mediated [Ca2+]i rises and [Ca2+]i rises dependent on the activation of VGCCs or NMDA receptors are not necessarily mutually exclusive.
IP3 in neurons is generated following activation of gq/11-coupled GTP-binding protein-coupled seven transmembrane domain receptors (GPCRs) such as group I metabotropic glutamate receptors (mGluRs) and M1-type muscarinic acetylcholine receptors (mAChRs) or gi/o-coupled GPCRs such as M2-type mAChRs and α2 adrenergic receptors. Following activation, these receptors stimulate g-protein guanine exchange and dissociation into gα and gβγ subunits. Gq/11α and gi/oβγ subunits, in turn, stimulate phospholipase C in the plasma membrane, which cleaves the membrane phospholipid phosphatidylinositol bis-phosphate into IP3 and diacylglycerol (DAG). Mobilized IP3 diffuses through the cytosol to bind IP3Rs, while DAG binds to and activates protein kinase C. Disentangling the particular effects of IP3 mobilization and subsequent IP3R activation downstream from GPCR stimulation is thus further complicated by the fact that the events leading to IP3 production also generate at least two additional second messengers, gq/11βγ or gi/oα and DAG.
IP3R activation in neocortical and hippocampal pyramidal neurons triggers propagating waves of internally released Ca2+ [7–11]. Two particularly intriguing consequences of these Ca2+ waves are the activation of a transient SK-type Ca2+-dependent K+ channel-mediated membrane hyperpolarization and the generation of a prolonged membrane depolarization (e.g., Fig. 1A and B; [11–15]). In vitro, these changes in membrane potential have been observed to alter the excitability and spiking patterns of individual pyramidal neurons [11–14]. Whether IP3R-mediated Ca2+ waves can similarly alter neuronal excitability in vivo, and how IP3R-dependent excitability changes might influence patterns of network activity and, ultimately, animal behavior are important questions. The importance of understanding how IP3R-dependent Ca2+ signaling contributes to normal cognitive function is underlined by a recent series of studies implicating dysregulated internal Ca2+ release in the pathophysiology of Alzheimer’s disease [16–19]. The results of these studies furthermore raise the question of whether IP3Rs might represent an important therapeutic target for the treatment of cognitive dysfunction. Our ability to address these and other questions whose aim it is to identify and understand the particular consequences of IP3R-dependent internal Ca2+ release is naturally limited by the selection of available experimental tools. One of the most valuable of these tools is a pharmacological IP3R antagonist. Ideally, any pharmacological agent employed in the study of IP3R signaling would be potent, specific, and membrane-permeant.
There are two commonly employed classes of commercially available and membrane-permeant IP3R antagonists: the marine sponge toxin xestospongin C and the boron compound 2-APB. Neither of these pharmacological agents acts only on IP3Rs, however. Xestospongin C, while a potent IP3R antagonist in some preparations [20,21], has in other preparations failed entirely to inhibit IP3R-mediated internal Ca2+ release [22,23]. Furthermore, it has been suggested to antagonize store-operated Ca2+ channels (SOCs) , and to interfere with Ca2+ store loading via antagonism of sarco-endoplasmic reticulum Ca2+ (SERCA) pumps [20,22,25, but see 26]. Xestospongin C has also been reported to inhibit voltage-dependent Ca2+ and K+ currents in smooth muscle cells, although this finding has not been verified in neuronal preparations . 2-APB has a similar and similarly large repertoire of targets, including IP3Rs [27–30, but see 22], SOCs [27–29,31], SERCA pumps [27–29,32], and transient receptor potential channels [33–35]. Furthermore, 2-APB has been shown to inhibit voltage-gated K+ channels , and is suggested to interfere with Ca2+ signaling mediated by voltage-gated Ca2+ channels .
Here we describe our observations that 2-APB consistently inhibits, but inconsistently blocks IP3-evoked internal Ca2+ release and Ca2+ waves and that 2-APB increases neuronal excitability in neocortical and hippocampal pyramidal neurons. Based on these findings, we conclude that 2-APB is of limited utility for studies seeking to examine how IP3R-mediated Ca2+ signaling and subsequent alterations in neuronal excitability might influence activity patterns in individual neurons, neuronal networks and systems, and, ultimately, animal behavior.
Brain slices were prepared from P22–P35 male Sprague–Dawley rats (n = 19 animals) as previously described . Briefly, animals were anesthetized with a mixture of ketamine, xylazine, and acepromazine, and decapitated when no longer responsive to a foot pinch. Coronal slices of the medial prefrontal cortex (320 µm) and horizontal hippocampal slices (350 µm) were cut using a Vibratome in a peltier-cooled slicing chamber filled with dissecting solution and maintained at a temperature between 0.5 and 3.0 °C. The dissection solution was continuously bubbled with 95% O2/5% CO2, and contained (in mM): 87 NaCl, 75 sucrose, 10 dextrose, 2.5 KCl, 25 NaHCO3, 1.3 NaH2PO4, 7.0 MgCl2, 0.5 CaCl2, adjusted with sucrose to 295–305 mOsm. After cutting, the brain slices were incubated for 10–20 min in 34–37 °C dissecting solution, and then transferred to 34–37 °C recording artificial cerebrospinal fluid (ACSF) and allowed to cool to room temperature for at least 1 h prior to recording. Standard recording ACSF contained (in mM): 124 NaCl, 10 dextrose, 2.5 NaHCO3, 1.3 NaH2PO4, 2.0 MgCl2, and 2.0 CaCl2, adjusted with sucrose to 295–305 mOsm.
Individual slices selected for recordings were continuously perfused with 95% O2/5% CO2-saturated recording ACSF (flow rate 1–2 ml/min) and maintained at 31–34 °C throughout the course of each experiment. Visualized whole-cell patch clamp recordings were performed using infrared differential interference contrast (DIC) microscopy on an upright microscope (Zeiss Axioskop or Olympus BX51WI) with thick-walled borosilicate glass patch pipettes (2–5MΩ). The pipette solution contained (in mM): 134 KMeOSO3, 10 HEPES, 3.0 KCl, 1.0 MgCl2, 4.0 Mg-ATP, 0.5 Na-GTP, 5.0K2-phosphocreatine, 5.0 Na2-phosphocreatine, pH adjusted with KOH to 7.52–7.55, 285–290 mOsm, as well as 50 units/ml creatine phosphokinase, 5–15 µM Alexa 488 or Alexa 568 for visualization of filled processes under fluorescent illumination, and one of the following Ca2+ indicator dyes: 100 µM bis-fura-2, 200 µM fura-2ff, 100 µM fluo-4, or 100 µM Oregon Green 488 BAPTA-2. KMeOSO3, Mg-ATP, Na-GTP, Na2-phosphocreatine, and creatine phosphokinase were obtained from Sigma–Aldrich (St. Louis, MO). K2-phosphocreatine was obtained from Calbiochem (San Diego, CA). Alexa 488, Alexa 568, bis-fura-2, fluo-4, and Oregon Green 488 BAPTA-2 were purchased from Molecular Probes/Invitrogen (Carlsbad, CA). For uncaging experiments, the internal solution was supplemented with 97 µM NPE-caged IP3 (Calbiochem/EMD Biosciences; San Diego, CA). Electrical signals were acquired at 2 kHz using an SEC 05LX amplifier (npi electronic; Tamm, Germany) in bridge or discontinuous voltage-clamp mode, digitized, and analyzed on- and off-line using custom software developed in IGOR Pro (WaveMetrics; Portland, OR). Whole-cell series resistance ranged from 5 to 37 MΩ and was compensated for by adjusting the bridge balance. Data were not corrected for junction potential (~10 mV). Cells were held at approximately −65 mV (prefrontal cortical neurons) or −63 mV (CA1 hippocampal pyramidal neurons) for the duration of the experiment. The 26 prefrontal cortical and three CA1 hippocampal pyramidal neurons included in this study had an average resting membrane potential of −59.1 ± 0.9 mV (cells were discarded if > −50 mV), an average input resistance of 84.6 ± 5.2 MΩ, and exhibited spike frequency adaptation in response to prolonged (300 ms) suprathreshold current injection. 2-APB (100 µM; Tocris Bioscience; Ellisville, MO) was prepared as a 100 mM stock solution in DMSO.
Ca2+ indicator dye diffused into the recorded neuron via the patch pipette. Dye fluorescence was imaged using a cooled CCD camera (Quantix 57 or Cascade 512B; Photometrics; Tucson, AZ). Images were collected at 50 or 25 Hz with 2 × 2 to 5 × 5 pixel binning. Epifluorescence illumination was provided by a 150 W short-arc xenon bulb (Optiquip; Highland Mills, NY). Relative changes in [Ca2+]i were quantified as changes in ΔF/F = |F(t) − F|/F, where F represents baseline fluorescence intensity prior to stimulation and ΔF represents the magnitude of fluorescence change during activity. Optical data were corrected for tissue autofluorescence and smoothed with 5- to 10-frame averaging. Relative changes in dye fluorescence in regions of interest over the soma and dendrites are displayed as changes in the amplitude of correspondingly colored optical traces.
Synaptic afferents were electrically stimulated using a glass microelectrode (5–10 µm tip diameter) with a fine tungsten rod glued to its side and filled with standard recording ACSF. Stimulating electrodes were placed ~20–60 µm away from the recorded cell body and ~20–50 µm to one side of its primary apical dendrite. Trains of unipolar pulses (20–50 stimuli, 15–150 µA, 0.1 ms, 50–100 Hz,) were delivered to elicit internal Ca2+ release. For focal pharmacological mGluR stimulation, a 2–5 MΩ pipette was filled with the group I/II mGluR agonist (±)-1-aminocyclopentanetrans-1,3-dicarboxylic acid (ACPD, 400 µM; Tocris Bioscience) in standard recording solution or in recording solution where 10 mM HEPES replaced 10 mM dextrose, and positioned ~20–60 µm away from the soma and <10 µm to one side of the primary apical dendrite. Focal stimulation of ryanodine receptors was similarly accomplished using a 2–5 MΩ pipette filled with caffeine (50 mM; Sigma–Aldrich) in standard recording solution and placed adjacent to the primary apical dendrite. NPE-caged IP3 diffused into the recorded neuron via the patch pipette. Photolysis of NPE-caged IP3 was achieved with 50–500 ms flashes of UV light (320–400 nm) produced by a 100 W mercury lamp (HBO ebq 100 isolated; Carl Zeiss, Inc.; Thornwood, NY). The photolysis beam (~20 µm diameter) was directed onto the soma or proximal apical dendrite of the recorded cell using a custom-made fiber optic spot illumination system (Rapp OptoElectronic GmbH; Hamburg, Germany) fitted to the aperture stop port in the epi-illumination pathway of an Olympus BX51WI microscope.
Input resistance was calculated from the average voltage response to a train of hyperpolarizing current pulses (150–250 pA, 300 ms duration). Voltage responses were measured during the last 20 ms of each 300 ms square pulse current injection and compared to baseline membrane potential in the 20 ms prior to current injection. Action potential heights were measured from the average membrane potential 20 ms prior to action potential onset and to the peak of the membrane potential deflection. Action potential widths were calculated at half-maximum amplitude. Single spike ADPs were calculated as the difference between the average membrane potential in the 5–20 ms following the onset of the action potential and the average membrane potential in the 20 ms prior to action potential onset. Single spike AHPs were calculated as the difference between the minimum membrane potential >1 ms following the peak of the action potential and the average membrane potential in the 2–7 ms prior to action potential onset. The time at which an action potential occurred was defined as the time at which the somatically recorded membrane potential reached its peak. Control, drug, and wash conditions typically corresponded to the following time periods: 15–0 min prior to 2-APB application, 10–30 min following addition of 2-APB to the recording ACSF, and 15–30 min after the start of 2-APB wash-out. Data from control cells were analyzed such that “control” and “drug” time periods were defined as 0–15 min and 30–45 min following the experimenter’s initial assessment of cellular health and/or internal Ca2+ release potential.
As there were no obvious differences in responses across ages or cell types, data were pooled. Data are presented as mean ± S.E.M. Statistical significance (p < 0.05) was assessed using two-tailed unpaired Student’s t-tests.
We first examined the effects of 2-APB on IP3R-dependent internal Ca2+ release and Ca2+ waves in neocortical and hippocampal pyramidal neurons in an in vitro slice preparation. Three different stimulation techniques were used to trigger internal Ca2+ release: focal pharmacological stimulation of mGluRs with the group I/II mGluR agonist ACPD (ACPD puffing), focal photolysis of NPE-caged IP3 (IP3 uncaging), and electrical stimulation of synaptic afferents. Regardless of the technique employed, we observed that 100 µM 2-APB consistently inhibited internal Ca2+ release, typically within less than 15 min (n = 15/17; inhibition first observed after 11.7 ± 1.3 min 2-APB exposure). More specifically, we found that 2-APB attenuated the amplitude and extent of Ca2+ waves in 4/9 neurons stimulated with puffs of ACPD (Fig. 1A), in 2/3 neurons where release was evoked by IP3 uncaging (Fig. 1B), and in 3/5 synaptically stimulated neurons. In one cell stimulated with puffs of ACPD and in another cell stimulated with uncaged IP3, 2-APB had no apparent effect on internal Ca2+ release (data not shown). 2-APB blocked internal Ca2+ release and Ca2+ waves in each of the remaining neurons tested (n = 6/17; block first observed after 11.3 ± 2.9 min 2-APB exposure; Fig. 1C). We did not observe recovery of internal Ca2+ release during 2-APB wash-out (n = 2). These findings are consistent with reports suggesting that 2-APB inhibits internal Ca2+ release by antagonizing IP3Rs, and are also in agreement with the observation that 2-APB only inconsistently blocks internal Ca2+ release evoked by IP3R stimulation . Importantly, the inhibition by 2-APB of IP3-induced internal Ca2+ release depends on the relative concentrations of 2-APB and IP3 . Our observation that 2-APB failed to block Ca2+ waves in over half of all cells stimulated synaptically or with puffs of ACPD (n = 8/14; release still present after 22.4 ± 2.7 min 2-APB exposure) and in every cell for which internal release was triggered by IP3 uncaging (n = 3/3; release still present after 30.3 ± 0.9 min 2-APB exposure) may therefore suggest that, in some cells, 2-APB is being out-competed by IP3.
Previous reports have suggested that, in addition to antagonizing IP3Rs, 2-APB may also influence intracellular Ca2+ signaling via inhibition of SERCA pumps [27–29,32]. Given the dependence of internal Ca2+ release in pyramidal neurons on prior VGCC-mediated Ca2+ influx and presumed loading of the readily releasable intracellular Ca2+ pool [11,37,38], it seems possible that the inhibition by 2-APB of IP3R-mediated internal Ca2+ release may result from 2-APB-dependent SERCA pump antagonism and consequent slow depletion of intracellular Ca2+ stores [27–29,32]. To further explore this possibility, we tested whether 2-APB inhibited internal Ca2+ release triggered by focal pressure application of the ryanodine receptor agonist, caffeine (caffeine puffing). We observed a reduction in the amplitude of caffeine-triggered internal Ca2+ release following 2-APB exposure (n = 3/3; inhibition first observed after 23.3 ± 1.9 min 2-APB exposure; Fig. 1D). However, the duration of 2-APB exposure prior to this inhibition was significantly greater than the duration of 2-APB exposure prior to inhibition or block of IP3R-mediated internal Ca2+ release (p < 0.0001 for both measures). Moreover, 2-APB did not completely block Ca2+ waves evoked by caffeine puffing in any of the cells tested (n = 3/3; release still present after 33.3 ± 1.3 min 2-APB exposure). While 2-APB-mediated SERCA pump antagonism may contribute to the inhibition of internal Ca2+ release in pyramidal neurons, it seems unlikely that this effect could, on its own, account for the inhibition by 2-APB of IP3R-mediated internal Ca2+ release.
In addition to its effects on internal Ca2+ release, we also observed that 2-APB elevated pyramidal neuron excitability. This increased excitability was readily identified as a change in the characteristics of action potential waveforms and spike trains. We examined action potentials evoked by prolonged (300 ms) and brief (2 ms) suprathreshold current injections. Action potential trains evoked by prolonged current injection under control conditions initiated after a slight delay (delay to 1st spike, 46.8 ± 1.4 ms, n = 12) and exhibited spike frequency adaptation (Fig. 2A). Subsequent to 2-APB application, delays preceding the onset of action potential trains were reduced. A comparison of averaged measurements made during the control period (in the 15 min prior to 2-APB application) and the drug period (between 10 and 30 min following 2-APB application) revealed that this change was significant. In particular, we observed that the delay to 1st recorded action potential was 29.8 ± 3.6 ms sooner in the presence of 2-APB than it was under control conditions (n = 12; p < 0.001; Fig. 2A and D). Spike frequency adaptation was also inhibited following 2-APB application. This inhibition was apparent as an elevation in the rate of action potential generation immediately following the first action potential (Fig. 2A and D), and was quantified using the firing frequency in the first 100 ms of a spike train. This firing frequency was significantly greater in the presence of 2-APB than during the control period (13.4 ± 3.8 Hz faster, n = 11; p < 0.05). In pyramidal neurons, both the delay preceding the onset of the first action potential in a spike train evoked by prolonged square pulse current injection and the frequency of action potentials at the start of the spike train are regulated by voltage-gated K+ currents [39,40]. The changes we observed in both these quantities therefore suggest that 2-APB may inhibit voltage-dependent K+ conductances. This hypothesis is further supported by our observation that action potentials triggered by both prolonged and brief current injection were significantly broader in the presence of 2-APB than action potentials evoked under control conditions (Figs. 2A, B, E and 3C, E). The mean width of the 4th action potential triggered by prolonged current injection, for example, was 0.67 ± 0.09 ms greater in the 10–30 min after 2-APB application than it was in the control period (n = 10; p < 0.001). Increases in action potential width did not result from a change in the kinetics of the action potential upstroke, as one might expect if Na+ conductances were affected, but rather from attenuation in the rate of membrane repolarization (Fig. 2A). Action potential repolarization in pyramidal neurons is controlled by voltage-dependent K+ conductances distinct from those involved in setting the delay to action potential initiation or the rate of firing during a spike train [39–42]. Based on this observation, we find it highly likely that 2-APB inhibits multiple families of voltage-dependent K+ conductances, including rapidly activating delayed-rectifier K+ channels of the KV class and slowly gated delayed-rectifier K+ channels of the KCNQ class. This result is in agreement with a report indicating that 2-APB inhibits two voltage-dependent K+ currents with very different kinetics in Limulus photoreceptors .
Afterhyperpolarizations were also significantly diminished in the 10–30 min following addition of 2-APB to the recording ACSF (Fig. 2A, C and F). The mean AHP of the 4th action potential triggered by prolonged current injection, for example, was 6.3 ± 0.8 mV smaller during 2-APB exposure than it was under control conditions (n = 7, p < 0.001). AHPs in pyramidal neurons are mediated predominantly by Ca2+-dependent K+ conductances [39–41,43,44]. Indeed, the reduction in firing frequency observed over the course of a prolonged suprathreshold current injection is thought to depend strongly on the accumulation of Ca2+ entering the neuron via VGCCs and subsequent activation of SK-type and other Ca2+-dependent K+ channels [39,40,43]. Antagonism of Ca2+-dependent K+ conductances therefore results in diminished AHPs and reduced late phase spike frequency adaptation. Inhibition by 2-APB of SK or other Ca2+-dependent K+ channels might thus account for the observed decrease in AHP magnitudes as well as a part of the increase in firing rates of action potentials triggered by prolonged current injection.
In addition to AHPs, neocortical and hippocampal pyramidal neurons frequently exhibit an afterdepolarization that appears as a small hump at the base of an action potential trace (Fig. 3A; [39,45,46]). We examined the magnitudes of ADPs following action potentials evoked with 2 ms current injections, and found that they were significantly greater following 10–30 min bath application of 2-APB than they were in the 15 min prior to 2-APB exposure (8.8 ± 1.9 mV greater, n = 14; p < 0.05; Fig. 3A, B and D). The magnitude of ADPs depends, in part, on the amplitude of AHPs, such that antagonism of K+ channels readily augments the ADP [45–47]. Our observation of an enhanced ADP is thus in line with the idea that 2-APB may inhibit the Ca2+-dependent K+ channels responsible for generating AHPs. Interestingly, the enhanced ADPs we observed were occasionally suprathreshold for action potential generation (n = 3/14 cells). Inhibition of SK channels alone is unlikely to explain the generation of these action potentials. In both layer V neocortical and hippocampal pyramidal neurons, SK channel immunoreactivity is strongest in the soma and proximal primary apical dendrites [48,49]. The smaller amplitudes and broader widths of ADP spikes suggest, however, that they were dendritically initiated. Moreover, activation of the currents underlying ADPs in pyramidal neurons requires the influx of Ca2+ via VGCCs. The enhanced ADPs we observe might therefore suggest an influence by 2-APB on Ca2+ signaling via VGCCs, a concept that is in keeping with the effects of 2-APB on SOCs [27–29,31] and TRPs [33–35]. Alternately, the larger and sometimes suprathreshold ADPs evoked in the presence of 2-APB may be secondary to the inhibition by 2-APB of dendritic voltage-dependent K+ conductances.
The altered firing patterns and action potential waveforms we observed after adding 2-APB to our recording ACSF were accompanied by an increase in the whole-cell input resistance (11.1 ± 6.0 MΩ increase, n = 8; p = 0.105). This increase would suggest that, in addition to influencing active and/or Ca2+-dependent K+ conductances, 2-APB might also antagonize passive or leak K+ currents. Alternately, a slight change in input resistance could simply reflect the antagonism of voltage-dependent K+ channels that are partially activated at resting potentials (e.g., KCNQ, Kir; ). A decrease in the resting K+ conductance, however, would also be expected to lower the potential difference across the plasma membrane. Indeed, we found that cells were slightly more depolarized following 10–30 min bath application of 2-APB than they were in the 15 min prior to 2-APB exposure (4.6 ± 2.6 mV increase, n = 5; p = 0.119).
In order to verify that the increases in excitability we observed were specific to 2-APB exposure, and not an artifact of our recording conditions, we evaluated the progression of each measure over the course of experiments in which 2-APB was not applied. With the exception of the post-spike ADP, all measures of intrinsic excitability were relatively stable (<10% change) over time in control cells. Control ADPs, however, decreased in magnitude over time, while ADPs following 2-APB exposure were, as described above, significantly increased. In sum, we found that nearly all measured changes in excitability in cells exposed to 2-APB were significantly different from those in control cells (Table 1). Moreover, the effects of 2-APB on neuronal excitability reversed during 2-APB wash-out (Figs. 2D, E and 3B–F).
On the basis of the findings presented here, we conclude that 2-APB, in addition to inhibiting IP3R-dependent internal Ca2+ release, also increases the excitability of neocortical and hippocampal and pyramidal neurons. Furthermore, we find – albeit by indirect means – that the mechanism of this increased excitability most likely involves the combined antagonism of multiple voltage-and Ca2+-dependent K+ conductances. Our findings have important implications for the interpretation of past and future studies using 2-APB to examine the consequences of IP3R signaling. Specifically, they suggest that observed changes in cellular, network, or animal behavior following the introduction of 2-APB might not be attributable to consequently impaired IP3R-mediated Ca2+ signaling, or even to changes in the activity of SOCs or SERCA pumps, but rather to unexpected or undesired changes in plasma membrane ion conductances and neuronal excitability.
We did not, in this study, evaluate the potential influence of the other membrane-permeant IP3R antagonist, xestospongin C, on neuronal excitability. The variety of Ca2+-carrying proteins and currents targeted by xestospongin C is strikingly similar to that also affected by 2-APB (IP3Rs, SOCs, SERCA pumps). In this light, and in view of its having been reported to antagonize voltage-gated K+ currents in smooth muscle , it might be sensible to test whether xestospongin C similarly alters the excitability of pyramidal neurons before turning to this drug as an alternative to 2-APB in the examination of IP3R-mediated internal Ca2+ release and its consequences.
In general, the results of this study highlight the limitations of pharmacological tools employed in the study of intracellular signaling. To a certain degree, these limitations may be overcome in animal models through the use of recombinant genetic strategies such as knock-in and knock-out mice and by the use of siRNAs. The utility of genetic manipulations for the treatment of human disease, however, is very limited for the present. We therefore find that both the study of IP3R-mediated neuronal signaling and the remediation of its dysfunction in disease would benefit highly from the development of a novel pharmacological agent specifically targeted to IP3 receptor ion channels.
We would like to thank Len Kaczmarek for a critical reading of the manuscript, as well as Keith Gipson and Joe Santos-Sacchi for many thoughtful discussions. This work was supported by the Kavli Foundation, the Dart Foundation, NIMH RO1-MH067830 and P50-MH068789, and NIAAA 1RL1AA017536-01 (MFY), an NIH-NHLBI/Yale BioSTEP Program Fellowship (CEB), and an NSF Graduate Research Fellowship (AMH).