Calcium regulation of a slow post-spike hyperpolarization in vagal afferent neurons

Abstract

Activation of distinct classes of potassium channels can dramatically affect the frequency and the pattern of neuronal firing. In a subpopulation of vagal afferent neurons (nodose ganglion neurons), the pattern of impulse activity is effectively modulated by a Ca²⁺-dependent K⁺ current. This current produces a post-spike hyperpolarization (AHP_slow) that plays a critical role in the regulation of membrane excitability and is responsible for spike-frequency accommodation in these neurons. Inhibition of the AHP_slow by a number of endogenous autacoids (e.g., histamine, serotonin, prostanoids, and bradykinin) results in an increase in the firing frequency of vagal afferent neurons from <0.1 to >10 Hz. After a single action potential, the AHP_slow in nodose neurons displays a slow rise time to peak (0.3–0.5 s) and a long duration (3–15 s). The slow kinetics of the AHP_slow are due, in part, to Ca²⁺ discharge from an intracellular Ca²⁺-induced Ca²⁺ release (CICR) pool. Action potential-evoked Ca²⁺ influx via either L or N type Ca²⁺ channels triggers CICR. Surprisingly, although L type channels generate 60% of action potential-induced CICR, only Ca²⁺ influx through N type Ca²⁺ channels can trigger the CICR-dependent AHP_slow. These observations suggest that a close physical proximity exists between endoplasmic reticulum ryanodine receptors and plasma membrane N type Ca²⁺ channels and AHP_slow potassium channels. Such an anatomical relation might be particularly beneficial for modulation of spike-frequency adaptation in vagal afferent neurons.

Activation and sensitization of primary afferent nerve fibers during allergic inflammation are orchestrated by inflammatory mediators released from various cells, including tissue mast cells. Inflammatory mediators provoke excitability changes in sensory nerves through diverse mechanisms, including (i) modification of the density and coupling efficacy of ligand-gated ionic channels; (ii) alteration in voltage-gated sodium, potassium, and calcium channels; and (iii) manipulation of cellular mechanisms that control spike-frequency adaptation.

After immunologic activation of mast cells in airway in vivo or in sensory ganglia in vitro, a wide range of electrophysiological changes can be detected in peripheral sensory nerve terminals of the vagus (1) and in vagal primary afferent somata (located in the nodose and jugular ganglia) (2). These changes range from transient (minutes) membrane depolarizations that sometimes reach action potential (AP) threshold (3) to a sustained (days) unmasking of functional NK-2 tachykinin receptors (4, 5). One electrical membrane property that is particularly sensitive to inflammatory mediators is a slow post-spike afterhyperpolarization (AHP_slow; see Fig. 1) (3). This slow afterpotential influences neuronal excitability and determines the frequency and pattern of neuronal discharge. We have found that the amplitude and duration of the AHP_slow are exquisitely sensitive to known inflammatory mediators such as prostanoids, amines, and kinins applied exogenously (Table 1) or released endogenously (i.e., after immunologic activation of mast cells) (3, 6). Inhibition of the AHP_slow is accompanied by a loss of spike-frequency adaptation. Thus, modulation of the AHP_slow amplitude and duration provides a mechanism for neuronal sensitization.

Figure 1.

A single AP can evoke three types of AHP in nodose neurons. (Top) A neuron with a single-component afterpotential lasting ≈30 ms. This AHP is designated AHP_fast. All neurons have this short duration afterpotential. (Middle) Example of a neuron with two afterpotentials, an AHP_fast followed by a longer lasting afterpotential (≈300 ms), the AHP_medium. In approximately half of the neurons, the AHP_medium is Ca²⁺-dependent. (Bottom) In a subset of C fiber type nodose neurons, a slowly developing (hundreds of ms) and long-lasting (2–15 s) afterpotential is observed. This slow afterpotential (AHP_slow) is always Ca²⁺-dependent. Intracellular recordings were obtained at room temperature from adult neurons isolated from rabbit nodose ganglia. The values near the horizontal lines are resting membrane potentials. The calibration in the Top also applies to the Middle. Similar results have been recorded in guinea pig and ferret nodose neurons.

Table 1.

Inflammatory mediators that block AHP_slow in vagal afferent neurons

Mediator	Receptor type	EC50, nM
Bradykinin	B2	72
Histamine	H1	2,000
Serotonin	nd	300
PGD2, PGE2	nd	∼20
Leukotriene C4	nd	∼100

Bradykinin (26), histamine (27), serotonin (28), PGD₂ and PGE₂ (12), and leukotriene C₄ (3) block the AHP_slow. nd, not determined; PG prostaglandin. 

We are interested in identifying the ionic channels and second-messenger transduction pathways that participate in the initiation and maintenance of the AHP_slow in vagal primary afferent neurons. In this report, we describe the general properties of this slow afterpotential and our progress in its characterization. Our working hypothesis is that a close functional proximity between three separate channels [N type voltage-sensitive calcium channels, ryanodine (RY)-sensitive Ca²⁺-induced Ca²⁺ release (CICR) calcium channels, and AHP_slow K⁺ (SK) channels that underlie the AHP_slow] is essential for the initiation of the AHP_slow.

RESULTS

General Properties of Vagal Afferent AHP_slow.

The AHP_slow is observed in a wide variety of peripheral and central neurons (for review, see ref. 7). In nodose neurons, AHP_slow is always preceded by a fast post-spike afterhyperpolarization (AHP_fast, 10–50 ms) that occurs at the end of the AP repolarization. In some neurons, the AHP_fast is followed by a second afterpotential that lasts 50–300 ms (AHP_medium). The AHP_medium is voltage- and Ca²⁺- dependent and blocked by 10 mM tetraethylammonium in ≈50% of neurons, suggesting that it is mediated by large-conductance Ca²⁺-activated K⁺ channels (BK channels) (8).

In vagal afferent somata, the AHP_slow is particularly robust. After a single AP, the AHP_slow displays a delayed onset (100–500 ms), a slow rise time to peak (0.3–5 s), and a long duration (2–15 s; see Fig. 1). The proportion of AHP_slow neurons within nodose ganglia varies among species: ≈20% in the guinea pig, ≈35% in rabbit, and ≈85% in ferret. Only nodose neurons classified as C fibers (conduction velocity <1 m/s) possess AHP_slow. To date, there have been few species differences in the pharmacological or physiological properties of the AHP_slow. An analogous slow AHP has also been recently described in ≈25% of C type dorsal root ganglion neurons of the rat (9, 10).

The AHP_slow in vagal afferent neurons influences cellular excitability and controls AP frequency over the physiological range from 0.1 Hz to 10 Hz (11, 12). One interesting property of the AHP_slow is that its amplitude is tuned to both AP number and frequency. Over the range of 1–100 Hz, the amplitude of the AHP_slow increases with the number of APs until it plateaus after ≈15 APs (Fig. 2); similar results were observed when the current underlying the AHP_slow was monitored. For reasons still unresolved, 10 Hz (100-msec interspike intervals) consistently evokes the largest responses.

Figure 2.

Effects of varying numbers of APs and frequency on the amplitude of the AHP_slow. All data points were recorded from a single acutely dissociated adult rabbit nodose neuron at room temperature. Resting potential and membrane input resistance were −55 mV and 53 MΩ, respectively. APs were evoked by transmembrane depolarizing current pulses (2 nA, 3 ms) at the frequencies indicated. Similar results were obtained when measuring I_AHP by using the hybrid voltage-clamp technique in rabbit, guinea pig, and ferret nodose neurons.

The current generating the AHP_slow (I_AHP) is a voltage-insensitive Ca²⁺-dependent K⁺ current (13, 14) that is unaffected by a wide range of K⁺ channel antagonists: 100 nM apamin, 10 μM d-tubocurarine, 5 mM Cs⁺, 30 mM tetraethylammonium, 10 mM Ba²⁺, 4 mM 4-aminopyridine, and 10 nM charybdotoxin. The magnitude of the AHP_slow (or the I_AHP) is linearly related to the concentration of extracellular Ca²⁺ (Fig. 3) and requires a rise in cytosolic free Ca²⁺ ([Ca²⁺]_i) for activation. Buffering intracellular Ca²⁺ with 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) abolishes the AHP_slow (Fig. 4). Noise analysis of the I_AHP suggests a single-channel conductance of ≈10 pS (unpublished observations). These features are consistent with the properties of a small-conductance Ca²⁺-activated K⁺ channel (SK channel; ref. 8). Of the several SK channels recently cloned from mammalian brain (15), the hSK1 channel has a pharmacological and biophysical profile compatible with the K⁺ current underlying the AHP_slow in nodose neurons.

Figure 3.

Effects of varying [Ca²⁺]_o on the amplitude of the AHP_slow recorded in isolated nodose neurons. (A) Sample traces of AHP_slow evoked by a train of four APs in the presence of different [Ca²⁺]_o. APs are evoked by transmembrane depolarizing current pulses (2 nA, 3 ms, 10 Hz) and are truncated. [Ca²⁺]_o was varied from 2.0 to 0.0 mM in 0.5 mM decrements. The AHP_slow is completely blocked when [Ca²⁺]_o is reduced to nominally zero. On returning to 2.0 mM [Ca²⁺]_o, the AHP_slow recovers to its original amplitude. (B) Relation between [Ca²⁺]_o and AHP_slow amplitude recorded in several neurons. Values are means ± SEM of the number of observations indicated near each data point. Data are normalized to the maximum response recorded in a given neuron. Linear regression analysis yields the solid line (r = 0.993).

Figure 4.

Effects of BAPTA on the AHP_slow and on the excitability of an acutely dissociated rabbit nodose neuron. (A) Bath-applied BAPTA/acetomethylester (10 μM) blocks the AHP_slow within 5 min without changing the resting membrane potential or membrane input resistance. APs were evoked by transmembrane depolarizing current pulses (4 nA, 1.5 ms, 10 Hz) and are truncated. (B) Responses recorded at a faster sweep speed to illustrate the kinetics of the AHP_fast and AHP_medium, which precede the AHP_slow. The AHP_fast is unaffected by 10 μM BAPTA/acetomethylester (compare a with b). The Ca²⁺ dependence of the AHP_medium is illustrated in c, where the neuron is superfused with 100 μM CdCl₂ for 30 s, which blocks most of the AHP_medium. The residual component of the AHP recorded in CdCl₂ is the AHP_fast, which is mediated by delayed rectifier K⁺ channels. (C) Depression of the AHP_slow markedly increases neuronal excitability. The average AP firing frequency induced by a current ramp protocol (1 nA, 2 s) increased from 1 to 5.5 Hz when the AHP_slow was blocked. Similar loss of spike-frequency adaptation was observed with bradykinin, prostaglandin D₂, histamine, and other inflammatory autacoids (see Table 2). The scale bar represents 3 mV, 2 s in A; 15 mV, 0.25 s in B; and 15 mV, 0.5 s in C. The dashed line represents the resting membrane potential (−60 mV). Resting membrane input resistance was 70 MΩ. Data is from ref. 19 with permission from the American Physiological Society.

Ca²⁺ Injection Evokes Two Temporally Distinct Outward Currents.

To test whether the K⁺ channels associated with the AHP_medium and the AHP_slow are directly activated by Ca²⁺, we iontophoretically injected Ca²⁺ into nodose neurons. Independent of the AHP_slow, a large outward current with rapid activation and decay kinetics was elicited by Ca²⁺ injection. This current (I_K-medium) was evoked at holding potentials between −2 mV and −45 mV. It was completely blocked by 5 mM tetraethylammonium but unaffected by inhibitors of the AHP_slow (100 nM prostaglandins D₂ or E₂ or 1 μM forskolin). I_K-medium was strongly voltage-dependent, requiring membrane holding potentials more positive than −55 mV. Assuming a reversal potential of −80 mV, I_K-medium had an e-fold increase in peak conductance for each 8.0 ± 1.0 mV (mean ± SEM; n = 8) depolarization, as calculated from semilogarithmic plots of peak chord conductance versus voltage-clamp holding potential. These properties are similar to those of large-conductance BK (AHP_medium) channels.

In neurons that exhibited AHP_slow, Ca²⁺ injection provoked a slowly developing and protracted outward current (I_K-slow). Fig. 5 shows an overlay of the outward current responses evoked by Ca²⁺ injection in a single nodose C type neuron at holding potentials of −20 mV and −50 mV. The kinetic differences between I_K-medium and I_K-slow after Ca²⁺ injection are dramatic. In contrast to the rapid activation of I_K-medium, the onset of I_K-slow is delayed, and the decay of I_K-medium is nearly complete before the peak amplitude of the I_K-slow is reached. These two outward currents mirror the temporal and pharmacological differences between AHP_medium and AHP_slow. I_K-slow, like the AHP_slow, was blocked by 100 nM prostaglandin D₂. The data shown in Table 2 summarize quantitative differences between these two Ca²⁺-induced outward currents.

Figure 5.

Comparison of two outward K⁺ currents evoked by intracellular Ca²⁺ injection. Recordings were made in a single acutely isolated adult rabbit nodose neuron. A slow outward current (I_K-slow) was activated by a 5-nA, 1-s iontophoretic Ca²⁺ injection at a holding potential of −50 mV. A second outward current (I_K-medium) was activated at −20 mV (5 nA, 0.5 sec). I_K-medium activates and decays completely before I_K-slow reaches peak amplitude. I_K-medium was blocked by 10 mM tetraethylammonium; I_K-slow was blocked by 100 nM prostaglandin D₂. The iontophoretic pipette was filled with a 0.2 M CaCl₂ solution. Voltage-clamp currents were recorded with a second intracellular pipette. The discontinuous (switched) current injection mode of an Axoclamp II amplifier was used for both currentand voltage-clamp applications. The larger calibration value is for I_K-medium. Population data is shown in Table 2.

Table 2.

Comparison of I_K-slow and I_K-medium

Current	Peak conductance, nS	n	Holding potential, mV	n	Time-to-peak, ms	n	Decay time constant, ms	n	Duration, s	n
IK-slow	27.9  ±  6.5	14	−55.4  ±  2.7	14	6,570  ±  1085	12	6,735  ±  789	5	23  ±  3.4	14
IK-medium	53.2  ±  16.5	6	−20  ±  3.7	6	958  ±  56	6	818  ±  97	6	2.5  ±  0.16	6

 I_K-slow and I_K-medium are outward currents elicited by iontophoretic injection Ca²⁺ into acutely isolated nodose neurons of the rabbit. The peak conductance is the largest conductance elicited, independent of membrane potential. The holding potential is the potential at which the peak conductance was measured. The decay time constant was measured by fitting a line, by eye, to the log transform of the decay of the current. The duration was calculated from the onset of Ca²⁺ injection to the time at which the current had decayed to 20% of its peak value. Data are summarized as the mean ± SEM. 

It is possible that the delayed onset of I_K-slow compared with I_K-medium results from unequal Ca²⁺ diffusion distances from the injection site to the two types of K⁺ channels. This cause seems unlikely because the orientation of impalement was random, and the plasma membranes of dissociated nodose neurons appear devoid of processes that would provide semi-isolated regions where I_K-slow might be generated. An alternative possibility is that additional intermediate steps, such as the synthesis or release of a second messenger, are required to activate I_K-slow. The large Q₁₀ (>3.0; ref. 14) supports the latter alternative. One candidate is mobilization of intracellularly stored Ca²⁺.

Ca²⁺ Released by the CICR Pool Is Essential for the Generation of the AHP_slow.

Single APs produce transient increases in [Ca²⁺]_i (ΔCa_t) as measured by the fluorescent indicator fura-2. The magnitude of the ΔCa_t depends on both [Ca²⁺]_o and the number of APs. Over the range of one to eight APs, there is an approximately linear relation between the magnitude of the ΔCa_t and the number of APs (Fig. 6). In the presence of drugs that block CICR but do not significantly affect AP-induced Ca²⁺ influx [(RY, 10 μM), 2,5,-di(t-butyl)hydroquinone (DBHQ, 10 μM), or thapsigargin (TG, 100 nM)], we found that at least eight APs were required to evoke a detectable ΔCa_t (Fig. 6). In the presence of RY, DBHQ, and TG, the ΔCa_t–AP relation exhibits slopes of 0.5, 1.1, and 0.8 nM per AP, respectively. When compared with the slope of 9.6 nM per AP in control neurons, Ca²⁺ influx produced by a single nodose AP is amplified by 5- to 10-fold by CICR (16). Nodose neurons demonstrate a relatively low stimulus threshold for eliciting CICR. For instance, a robust CICR response can be observed after a single AP stimulus in nodose neurons, whereas many tens of APs are required in dorsal root ganglion neurons (17). The greater CICR response in nodose neurons is not due to greater Ca²⁺ influx through voltage-dependent Ca²⁺ channels (VDCCs); a single AP produces comparable Ca²⁺ influx in nodose and dorsal root ganglion neurons (39 vs. 49 pC, respectively; refs. 16 and 18). Rather, the more responsive CICR pool in nodose neurons may reflect either a closer proximity between plasma membrane Ca²⁺ influx channels and endoplasmic reticulum RY receptors or a more sensitive RY receptor.

Figure 6.

(Upper) Effect of RY on AP-induced Ca²⁺ transients. Traces are Ca²⁺ transients evoked by varying numbers of APs, as indicated below each trace. In control neurons, distinct Ca²⁺ transients can be elicited by very few APs. In contrast, in the presence of 10 μM RY, a CICR inhibitor, at least eight APs are required to generate a discernible change in [Ca²⁺]_i. Suppression of the Ca²⁺ transient by RY is due to its effect on CICR and not the result of nonspecific effects on Ca²⁺ channels; the kinetics and amplitude of I_Ca elicited by APs are completely unaffected by RY. (Lower) Effect of RY on the relation between the amplitude of Ca²⁺ transients and number of APs. ○ and ● are mean amplitudes of Ca²⁺ transients evoked by varying numbers of action potentials for control (n = 10) and for RY-treated nodose neurons (n = 3), respectively. Linear regression of data from control (≤4 action potentials) and RY-treated cells yielded slopes of 9.6 ± 0.01 and 0.5 ± 0.23 nM per AP, respectively. Comparison of the slopes illustrates that CICR is capable of amplifying the “trigger” Ca²⁺ resulting from AP-induced Ca²⁺ influx by 20-fold. Data is modified from ref. 16 with permission from Journal of Physiology (London).

By using physiological stimuli (APs) in conjunction with pharmacological manipulations of CICR, we have demonstrated that CICR is essential for the development of the AHP_slow. Over the range of 1–16 APs, the magnitudes of the AP-induced AHP_slow and the ΔCa_t (a monitor of CICR in these neurons) were highly correlated (r = 0.985). Simultaneous recordings of ΔCa_t and AHP_slow before and during bath application of CICR inhibitors (RY, TG, DBHQ, or 10 μM cyclopiazonic acid) revealed that both responses were blocked in a parallel fashion (Fig. 7; see also Table 1 in ref. 19). These data indicate that a CICR pool is essential for the generation of the AHP_slow. They also provide a potential explanation for the slow kinetics of the AHP_slow, namely Ca²⁺ mobilization from CICR.

Figure 7.

Effect of DBHQ, a functional CICR inhibitor, on the AP- induced Ca²⁺ transient and on the AHP_slow recorded simultaneously in an acutely isolated rabbit nodose neuron. Upper traces represent superimposed Ca²⁺ transients evoked by a train of four APs (10 Hz) recorded in control Locke solution and 7 min after switching to Locke solution containing 10 μM DBHQ. The lower pair of traces shows AHP_slow. DBHQ treatment completely blocked both the Ca²⁺ transient and the AHP_slow. Resting [Ca²⁺]_i was 91 nM. Fluorescence data were acquired at 10 Hz. Resting membrane potential was −67 mV. AP amplitudes are truncated. Data are from ref. 19 with permission from the American Physiological Society.

Effects of Changing [Ca²⁺]_o on the AHP_slow, ΔCa_t, and Ca²⁺ influx.

If the AHP_slow depends on Ca²⁺ released from the CICR pool triggered by AP-induced Ca²⁺ influx, it would follow that changes in [Ca²⁺]_o should produce corresponding effects on both the AHP_slow and the ΔCa_t. The data shown in Fig. 3A illustrate the effects of progressively lowering [Ca²⁺]_o from 2.0 mM to nominally zero on the amplitude of the AHP_slow recorded in a single nodose neuron. As [Ca²⁺]_o was decreased, the amplitude of the AHP_slow was reduced proportionally. When the results from this and five additional neurons were plotted (Fig. 3B), the relation between [Ca²⁺]_o and the amplitude of the AHP_slow was linear (r = 0.993; n = 6, pooled data from three current-clamp and three hybrid voltage-clamp experiments).

Next, we examined the relation between [Ca²⁺]_o and the magnitude of the AP-induced ΔCa_t. Fig. 8A illustrates ΔCa_ts elicited by varying numbers of APs recorded from a single neuron in Locke solution containing 2.2 or 1.1 mM Ca²⁺. The population results relating the normalized amplitude of the ΔCa_ts recorded in four neurons to the number of APs is shown in Fig. 8B. In 1.1 mM [Ca²⁺]_o, the first few APs did not elicit a measurable ΔCa_t. For the neuron shown in Fig. 8A, at least eight APs were necessary to evoke a detectable ΔCa_t. In three additional neurons, the minimum number of APs necessary to elicit a detectable ΔCa_t ranged from 4 to 32. The ΔCa_t–AP relation recorded in 1.1 mM [Ca²⁺]_o, as in Locke solution containing normal [Ca²⁺]_o, followed a hyperbolic relation (χ² = 6.75 and 0.31; r = 0.988 and 0.999 for 2.2 and 1.1 mM Ca²⁺,respectively; Fig. 8B and see also Fig. 1 in ref. 16). Given the hyperbolic nature of the ΔCa_t–AP relation, deducing the effects of altered [Ca²⁺]_o on the magnitude of the ΔCa_t clearly depends on where along this relation the comparison is made. At one extreme, there is a ≈2-fold change when comparing the plateau phases of the curves in normal and one-half normal [Ca²⁺]_o. It is also possible to calculate the limiting initial slopes for the rising phase of the curves (dashed lines in Fig. 8B). The limiting slopes, which represent the full Ca²⁺ release potential of the CICR pool before any release has actually occurred, were 15 ± 3.8 and 2 ± 0.7 nM per AP in 2.2 and 1.1 mM [Ca²⁺]_o, respectively. Thus, reducing [Ca²⁺]_o by a factor of 2 results in a reduction of the ΔCa_t by a factor of 7 ± 2.8 when the rising phases of the two curves are compared. The ≈7-fold reduction of the ΔCa_t associated with halving [Ca²⁺]_o is much larger than the 2-fold reduction in the AHP_slow amplitude (Fig. 3), suggesting that some, but not all, of the Ca²⁺ released from the CICR pool is required for the generation of the AHP_slow.

Figure 8.

Effect of varying [Ca²⁺]_o on the amplitude of AP -induced Ca²⁺ transients. (A) Representative traces of Ca²⁺ transients evoked by varying numbers of APs in normal (2.2 mM) and reduced (1.1 mM) [Ca²⁺]_o. APs were elicited by transmembrane depolarizing current pulses (2 nA, 1.5 ms, 10 Hz). The number of APs is indicated below each trace. (B) The normalized (mean ± SEM) amplitude of Ca²⁺ transients recorded in four neurons is plotted against varying numbers of APs. Data are normalized to the maximal response recorded in a given neuron. ○ represents Ca²⁺ transients recorded in 2.2 mM [Ca²⁺]_o; ● represents Ca²⁺ transients recorded in the same neurons in 1.1 mM [Ca²⁺]_o. Continuous curves are rectangular hyperbolas fit to the data (χ² = 6.75 and 0.31, r = 0.988 and 0.999 for 2.2 and 1.1 mM [Ca²⁺]_o, respectively). The dashed lines represent the limiting initial slopes (15 ± 3.8 and 2 ± 0.7 nM per AP for 2.2 and 1.1 mM [Ca²⁺]_o, respectively).

The disproportionate effect of reduced [Ca²⁺]_o on the ΔCa_t versus the AHP_slow could arise from a nonlinear reduction of Ca²⁺ influx per AP and/or from a decreased Ca²⁺ release from CICR pool per unit Ca²⁺ influx. To study these possibilities, we examined the effect of lowering [Ca²⁺]_o on AP-induced Ca²⁺ influx. The amount of Ca²⁺ entering a neuron with each AP in normal and in reduced [Ca²⁺]_o was determined by using a prerecorded AP as whole-cell voltage-clamp command under experimental conditions where the major inward charge carrier is Ca²⁺ (for details, see Fig. 2 in ref. 16). When [Ca²⁺]_o was decrementally reduced from 2 mM to nominally zero, the magnitude of the I_Ca decreased proportionally. The charge movement caused by Ca²⁺ influx, normalized to cell membrane capacitance (pC/pF), was plotted against varying [Ca²⁺]_o for 12 neurons. Over the range of 0–2.0 mM [Ca²⁺]_o, Ca²⁺ influx varied linearly with [Ca²⁺]_o (r = 0.974). These results indicate that changes in Ca²⁺ influx alone cannot account for the disproportionate reduction in the ΔCa_t relative to the AHP_slow that is observed when [Ca²⁺]_o is reduced.

The disproportionate effect of reduced [Ca²⁺]_o on the ΔCa_t–AHP_slow relation could arise from a diminution in the amount of Ca²⁺ released from the CICR pool. Caffeine, a known agonist of CICR, is traditionally used to assess the releasable content of the CICR pool. In 8 of the 13 neurons studied, halving [Ca²⁺]_o reduced the caffeine-induced ΔCa_t by 20–79% (100% vs. 47 ± 7.2% in 2.2 and 1.1 mM [Ca²⁺]_o, respectively; P = 0.0002). In other words, decreasing [Ca²⁺]_o by a factor of 2 caused a 1.25- to 5-fold reduction in the caffeine response. On returning to normal Locke solution, the caffeine response was restored to near control values. In the remaining five neurons, the caffeine-induced ΔCa_t was unaffected by reducing [Ca²⁺]_o (100% vs. 112 ± 8.4% in 2.2 and 1.1 mM [Ca²⁺]_o, respectively; P = 0.690). There was no significant difference in resting levels of [Ca²⁺]_i between these two groups of neurons (93 ± 29.5 nM vs. 111 ± 29.7 nM; P = 0.530). Unfortunately, the wide variability in the effects of reduced [Ca²⁺]_o on the caffeine responses prevents a meaningful interpretation of the effect of [Ca²⁺]_o on the releasable content of the CICR pool.

Ca²⁺ Influx Through N Type Calcium Channels Selectively Elicits AHP_slow.

Six types of VDCCs have been described in neurons: L, N, P, Q, R, and T (20). Nodose neurons express several types of VDCCs. By using a panel of pharmacologic reagents that are selective for different types of VDCCs, we tested the contribution of each to the total AP-induced Ca²⁺ current. Our results, summarized in Table 3, reveal that ≈85% of the AP-induced inward Ca²⁺ current is shared by L and N type Ca²⁺ channels (Fig. 9). P, Q, and T type Ca²⁺ channel antagonists were ineffective, suggesting that the remaining Ca²⁺ current is associated with Ca²⁺ influx through R type channels. Nifedipine (10 μM), an L type Ca²⁺ channel blocker, produced no measurable effect on either the AHP_fast, the AHP_medium, or the AHP_slow. By contrast, ω-conotoxin-GVIA (0.5 μM), a selective N type Ca²⁺ channel blocker, always obliterated the AHP_slow, and in ≈50% of the neurons abolished the AHP_medium (about half of the AHP_medium are Ca²⁺-sensitive, see above), while leaving the AHP_fast unaffected (Fig. 9 and Table 4.). These results indicate that the SK and BK type K⁺ channels are both regulated by Ca²⁺ influx through N type channels. BK channels are gated by influx Ca²⁺ directly (8), whereas SK channels are affected by influx Ca²⁺ indirectly (i.e., Ca²⁺ entering through N type VDCC triggers RY receptors to release Ca²⁺ from CICR pools). Such a sequence implies a functional coupling between N type Ca²⁺ channels and RY channels in the endoplasmic reticulum. We tested this proposition by examining the effects of VDCC antagonists on the magnitude of AP-induced ΔCa_t.

Table 3.

Effects of Ca²⁺ channel blockers on action potential-induced inward Ca²⁺ currents

Channel type	Channel blocker	Concentration μM	Reduction	n
T	Amiloride	500	0  ±  0	18
L	Nifedipine	10	44  ±  5.6	9
P/Q	ω-AGA IVA	0.2	0  ±  0	2
Q	ω-CTX MVIIC	0.25	0  ±  0	6
N	ω-CTX GVIA	1	40  ±  4.0	15

The blocking effect of amiloride, nifedipine, ω-agatoxin (AGA) IVA, ω-conotoxin (CTX) MVIIC, and ω-conotoxin (CTX) GVIA is expressed as percent reduction in the peak amplitude of the total calcium current ± SEM. n corresponds to the number of cells for each condition. 

Figure 9.

Effects of VDCC antagonists on AP-induced calcium currents, AHP_slow and AP-induced Ca²⁺ transients. (A) Inward calcium currents recorded in isolated nodose neurons evoked by a prerecorded AP waveform from a holding potential of −60 mV. From Left to Right, control inward current in the presence of 2 mM [Ca²⁺]_o and in the presence of 10 μM nifedipine. After reestablishing control conditions, the neuron was exposed to 1 μM ω-conotoxin-GVIA. The effects of 500 μM cadmium were recorded in another neuron; the control current for this cell was similar to the first trace. (B) AHP_slow evoked by a train of four APs (10 Hz) recorded in another nodose neuron. From Left to Right, AHP_slow evoked in control conditions, in the presence of 100 μM CdCl₂, after washout, in the presence of 500 nM ω-conotoxin-GVIA, and after washout. (C) AP-induced Ca²⁺ transients recorded in two nodose neurons. From Left to Right, Ca²⁺ transients evoked by a train of eight APs in normal Locke solution, and in Locke solution containing 10 μM nifedipine. In another neuron, 1 μM ω-conotoxin-GVIA reduced the Ca²⁺ transient ≈50% (see Table 4). APs were evoked by 2.5-ms, 10-Hz depolarizing current pulses.

Table 4.

Actions of specific Ca²⁺ channel blockers on the action potential-induced Ca²⁺ transient and the AHP_slow

Channel type	Channel blocker	Reduction, %
		Ca2+ transient	n	AHPslow amplitude	n
L	Nifedipine	57  ±  7.7	21	0  ±  0	5
N	ω-CTX GVIA	39  ±  6.2	4	100  ±  0	6
T, R	Nickel	nd		0  ±  0	5
All	Cadmium	100  ±  0	2	100  ±  0	6

The following concentrations of antagonists were used: nifedipine (10 μM), ω-conotoxin GVIA (0.5 μM or 1 μM), nickel (50−500 μM), and cadmium (100 μM). nd, not determined. 

Ca²⁺ influx through both L and N type Ca²⁺ channels triggers CICR. The magnitude of the ΔCa_t is a sensitive indicator of Ca²⁺ release from the CICR pool. To determine the relative influence of Ca²⁺ influx through L and N type channels on release from the CICR pool, we applied selective VDCC antagonists and monitored the amplitude of ΔCa_t. Nifedipine (10 μM) and ω-conotoxin-GVIA (0.5–1.0 μM) diminished the amplitude of the ΔCa_t by 57% and 39%, respectively (Fig. 9 and Table 4). These results reveal that Ca²⁺ entering through either L or N type Ca²⁺ channels provides “trigger” Ca²⁺ to stimulate CICR. Given that the amount of Ca²⁺ influx through L and N type Ca²⁺ channels is comparable (44% and 40%, respectively, of total AP-induced Ca²⁺ influx; see Table 3), there must be a remarkable spatial arrangement between plasma membrane N type Ca²⁺ channels, endoplasmic reticulum RY receptors, and plasma membrane SK channels. Our working hypothesis concerning the regulation of the AHP_slow by Ca²⁺ is illustrated schematically in Fig. 10.

Figure 10.

Schematic diagram of the relation between plasma membrane Ca²⁺ channels, BK, and SK potassium channels and endoplasmic reticulum RY receptors in primary vagal afferent neurons. Single APs evoke Ca²⁺ influx through L and N type VDCCs. Ca²⁺ influx through either of these channels can trigger release of Ca²⁺ from the endoplasmic reticulum via RY receptors. Whereas BK channels are activated directly by Ca²⁺ entering the neuron via N type VDCC, SK channels are activated indirectly. SK channels require Ca²⁺ from CICR pools released after Ca²⁺ influx through N type channels.

DISCUSSION

Whether recorded in intact vagal sensory ganglia or in acutely isolated vagal afferent somata (nodose neurons), single APs can elicit an AHP_slow that exhibits a delayed onset (50–300 ms), a slow time to peak amplitude (0.3–0.5 s), and a particularly long duration (2–15 s) (14, 21). Inhibition of the AHP_slow by numerous inflammatory mediators (e.g., bradykinin, prostanoids, histamine, serotonin, leukotriene C₄; see Table 1) results in an increased neuronal excitability and a loss of spike-frequency adaptation. Thus, modulation of the AHP_slow by these mediators provides a mechanism for peripheral nociceptor sensitization that may underlie allergic inflammation-induced hyperalgesia.

One unresolved but important mechanistic question revolves around the delayed onset and protracted duration of the AHP_slow. Many of our studies of nodose AHP_slow were performed with acutely dissociated adult neurons, which are essentially spherical structures lacking dendritic and axonal processes. Thus, the delayed onset of the AHP_slow cannot be due to slow diffusion of Ca²⁺ from distal sites of influx to somal SK channels. The high temperature coefficient (Q₁₀ > 3.0) for the rising phase and the decay time constant of the nodose AHP_slow (14) also argues against simple Ca²⁺ diffusion as an explanation for the slow kinetics of the AHP_slow. The time course of the AHP_slow could arise from unusual channel kinetics of the SK channels. This also appears unlikely if SK channels in nodose neurons have activation kinetics similar to those cloned from rat brain (22). Recombinant SK channels from rat brain have activation time constants that are orders of magnitude shorter than the rise time of the AHP_slow. It is more likely that the time course of the AHP_slow is a consequence of the ΔCa_t because of CICR.

If the AHP_slow is directly dependent on Ca²⁺ released from the CICR pool, the AHP_slow and the AP-induced rise in [Ca²⁺]_i should display similar kinetics. Quantitative kinetic comparisons between these two variables are subject to some uncertainty, because the time course of the ΔCa_t reflects global changes in [Ca²⁺ ]_I, whereas the kinetics of the AHP_slow are determined by events at the plasma membrane. Nonetheless, we determined the time-to-peak and 10-to-90% decay time for both the AHP_slow and the ΔCa_t elicited by one to eight APs (19). The time-to-peak for AHP_slow was significantly slower than the ΔCa_t by nearly a factor of a two (1.0 s vs. 1.9 s); the ΔCa_t also decayed more rapidly than the AHP_slow (3 s vs. 7 s). Analogous temporal discrepancies have been reported between the ΔCa_t and AHP_slow in vagal motoneurons (23). Such temporal differences suggest that Ca²⁺ released from CICR pools does not act alone to gate AHP_slow K⁺ channels. Cloned SK channels contain many potential phosphorylation sites (15); Ca²⁺-dependent phosphorylation and/or dephosphorylation may thus be additional processes in the signal-transduction pathway of AP-evoked AHP_slow.

Unambiguous data now exist showing that Ca²⁺ can directly activate SK channels in hippocampal neurons (24) and in Xenopus oocytes (22). In nodose neurons, it is less clear whether Ca²⁺ alone is sufficient to activate and sustain the AHP_slow after an AP. In hippocampal neurons, flash photolysis of a “caged” Ca²⁺ chelator immediately truncates AP-induced AHP_slow, suggesting that elevated intracellular Ca²⁺ is required to maintain the AHP_slow (25). These results do not, however, distinguish between continuous Ca²⁺ gating of SK channel and the involvement of other Ca²⁺-dependent factors sustaining the longevity of the AHP_slow. It is also possible that Ca²⁺-dependent factors act synergistically with Ca²⁺ to control SK channels (23). The nearly spherical morphology and large size of acutely isolated adult nodose neurons provide a favorable preparation to determine the nature of second messengers required to activate and sustain the AHP_slow.

In conclusion, a subset of vagal primary afferent neurons possess a slowly developing and long-lasting spike afterhyperpolarization, the AHP_slow, that can profoundly affect the discharge frequency of these visceral afferent neurons. Although AP-evoked Ca²⁺ influx via both L and N type Ca²⁺ channels triggers CICR, only Ca²⁺ flux through N type channels activates the CICR-dependent AHP_slow. This type of specificity suggests that spatial coupling between N type Ca²⁺ channels and SK channels may be critical for the generation of the AHP_slow in nodose neurons. The exact mechanism coupling ΔCa_t to the AHP_slow current remains to be determined.

ABBREVIATIONS

AP

action potential

BK

large-conductance Ca²⁺-activated K⁺ channels

SK

small-conductance Ca²⁺ -activated K⁺ channels

CICR

Ca²⁺-induced Ca²⁺ release stores

RY

ryanodine

VDCC

voltage-dependent Ca²⁺ channels

L

N, R, L type, N type, and R-type VDCC

AHP

afterhyperpolarization

DBHQ

2,5,-di(t-butyl)hydroquinone