Mechano- and chemosensitivity of rat nodose neurones – selective excitatory effects of prostacyclin


Corresponding author F. M. Abboud: Department of Internal Medicine, University of Iowa, 200 Hawkins Drive, Iowa City, IA, 52242, USA. Email:


Nodose ganglion sensory neurones exert a significant reflex autonomic influence. We contrasted their mechanosensitivity, excitability and chemosensitivity in response to the stable prostacyclin (PGI2) analogue carbacyclin (cPGI) in culture. Under current clamp conditions we measured changes in membrane potential (ΔmV) and action potential (AP) responses to mechanically induced depolarizations and depolarizing current injections before and after superfusion of cPGI (1 μm and 10 μm). Chemosensitivity was indicated by augmentation of AP firing frequency and increased maximum gain of AP frequency (max. dAP/dΔmV), during superfusion with cPGI. Results indicate that two groups of neurones, A and B, are mechanosensitive (MS) and one group, C, is mechanoinsensitive (MI). Group A shows modest depolarization without AP generation during mechanical stimulation, and no increase in max. dAP/dΔmV, despite a marked increase in electrical depolarization with cPGI. Group B shows pronounced mechanical depolarization accompanied by enhanced AP discharge with cPGI, and an increase in max. dAP/dΔmV. Group C remains MI after cPGI but is more excitable and markedly chemosensitive (CS) with a pronounced enhancement of max. dAP/dΔmV with cPGI. The effect of cPGI on ionic conductances indicates that it does not sensitize the mechanically gated depolarizing degenerin/epithelial Na+ channels (DEG/ENaC), but it inhibits two voltage-gated K+ currents, Maxi-K and M-current, causing enhanced AP firing frequency and depolarization, respectively. We conclude that MS nodose neurones may be unimodal MS or bimodal MS/CS, and that MI neurones are unimodal CS, and much more CS to cPGI than MS/CS neurones. We suggest that the known excitatory effect of PGI2 on baroreceptor and vagal afferent fibres is mediated by inhibition of voltage-gated K+ channels (Maxi-K and M-current) and not by an effect on mechanically gated DEG/ENaC channels.

In this study we address the mechanosensitivity and chemosensitivity of vagal afferent neurones isolated from nodose ganglia. For over 40 years the question of bimodal activation (i.e. mechanical and chemical) of vagal afferents has been studied mostly at the level of single afferent fibres (Coleridge et al. 1964; Page et al. 2002; Veelken et al. 2003; Page et al. 2005). Having identified specific mechanically activated non-voltage-gated ion channels (Cunningham et al. 1997; Drummond et al. 1998; Kraske et al. 1998; Price et al. 2000) as well as voltage-gated potassium channels sensitive to PGI2 in nodose neurones (Li et al. 1997; Snitsarev et al. 2005b), we now address the issue of bimodality at the level of specific conductances in the neurones rather than by measuring nerve activity at the single fibre level.

Aortic baroreceptor neurones as well as other nodose sensory neurones innervating the heart, lung, gastrointestinal tract and large arteries are mechanosensitive (MS) (Coleridge et al. 1964; Higashi et al. 1982; Armour et al. 1994; Cunningham et al. 1997; Kraske et al. 1998; Abysique et al. 1999; Snitsarev et al. 2002; Page et al. 2005) and/or chemosensitive (CS) (Higashi et al. 1982; Leal-Cardoso et al. 1993; Undem & Weinreich, 1993; Armour et al. 1994; Smid et al. 2001; Zhu et al. 2001; Linz & Veelken, 2002; Page et al. 2002, 2005). In response to mechanical deformation of visceral organs, cardiac distension, and a rise in arterial pressure, or in response to chemical stimulation by autocrine or paracrine factors such as bradykinin and prostaglandins (McDowell et al. 1989; Zucker et al. 1989; Xie et al. 1990; Wang et al. 1993; Ustinova & Schultz, 1994; Weinreich et al. 1995; Schultz et al. 1997; Schultz, 2001; Snitsarev et al. 2005b), the nerve endings in these organs initiate action potential (AP) discharges that propagate along vagal afferents and aortic depressor nerves through the nodose ganglia to the central nervous system. Action potential discharge in afferent fibres is measured away from the site of mechano- or chemoelectrical transduction in the sensory nerve terminals. The small size and complex architecture of the baroreceptor or other sensory terminals embedded in the vascular wall or in tissues has generally prevented the direct measurement of membrane potential in the terminals and limited investigation into mechanisms of baroreceptor activation or sensory signalling in vivo. Furthermore, the presence of endothelium and vascular muscle along with mucosal and other cell types in the vascular wall make it difficult to attribute changes in baroreceptor (BR) or sensory nerve activity solely to direct actions on the nerve terminals. Factors released from nearby cells and changes in vascular smooth muscle tone can alter BR activity (Chapleau & Abboud, 2004). These limitations motivated us and others to develop an in vitro preparation of isolated nodose neurones in culture. Thus we have demonstrated that cultured BR neurones are mechanosensitive. Mechanical stimulation of isolated BR neurones evokes an inward cationic current (voltage clamp), depolarizes the membrane (current clamp), and increases cytosolic calcium concentration (Cunningham et al. 1995, 1997; Sullivan et al. 1997; Drummond et al. 1998; Snitsarev et al. 2002). Mechanosensitive channels have been identified in isolated BR neurones at the single channel level (Kraske et al. 1998). Similar to our findings in vivo (Hajduczok et al. 1994), we have shown that gadolinium blocks responses to mechanical stimulation of BR neurones in vitro (Cunningham et al. 1995, 1997; Sullivan et al. 1997).

We have also reported that the molecular components of the mechanosensitive ion complex in BR neurones include members of the degenerin/epithelial Na+ channel (DEG/ENaC) family that we localized in nodose neurones and more importantly in the aortic baroreceptor terminals (Drummond et al. 1998, 2001; Ma et al. 2001b). DEG/ENaC proteins constitute important components of a MS cationic complex that is non-voltage gated, non-ligand gated, amiloride sensitive and conserved through evolution (Huang & Chalfie, 1994; Tavernarakis & Driscoll, 1997).

Following a mechanically induced depolarization at the nerve terminal, the activation of voltage-gated channels at the ‘spike initiating zone’ near the terminals (Katz, 1950; Grigg, 1986) leads to the generation of APs. These two electrical events, mechanically evoked depolarization (through DEG/ENaC channels) and AP generation (through voltage-gated channels), may be distinguished pharmacologically by blocking the former with amiloride or gadolinium (Hamill et al. 1992; Hajduczok et al. 1994; Cunningham et al. 1997; Sullivan et al. 1997; Drummond et al. 1998, 2001) while APs triggered by depolarizing current injections are preserved (Snitsarev et al. 2002).

We and others have also reported that PGI2 and its analogues increase the sensitivity of baroreceptor and vagal afferent nerves in vivo (McDowell et al. 1989; Zucker et al. 1989; Chen et al. 1990; Xie et al. 1990; Wang et al. 1993), that it is an autocrine enhancer of excitability in nodose neurones (Snitsarev et al. 2005b), and that the stable PGI2 analogue carbacyclin (cPGI) (Adaikan et al. 1980; Aiken & Shebuski, 1980; Whittle et al. 1980) inhibits the charybdotoxin (ChTX)-sensitive calcium-dependent K+ current, Maxi-K (Li et al. 1997).

In this work we contrasted the ionic mechanisms that determine mechanosensitivity, excitability, and chemosensitivity of nodose neurones in culture. We tested whether chemical activation by cPGI enhances the mechanically induced depolarizing potential (the mechanically gated channels) of the neurones, or the voltage-gated AP firing (increased membrane excitability) or both, and whether cPGI induces mechanosensitivity in neurones that are constitutively mechanoinsensitive (MI). Our results indicate that the mechanically induced depolarization through the DEG/ENaC channels is not enhanced by PGI2 whereas the AP generation may be enhanced in approximately 40% of the mechanosensitive (MS) neurones. Thus, bimodality in some neurones is achieved by a selective chemical modulation of voltage-gated ion channels and not by sensitization of MS conductances. We also found that two voltage-gated potassium channels, Maxi-K and M-currents, appear to mediate the effect of PGI2 on nodose neurones. Although cPGI had a much greater excitatory effect on MI neurones, it never induced mechanosensitivity in these neurones. Given the dominant role of nodose neurones in modulating autonomic control, and the autocrine influence of PGI2 on these neurones, the results provide insights into mechanisms of altered mechano- and chemosensitivity in pathophysiological states such as heart failure and hypertension.


All experimental protocols have been approved by the University of Iowa Animal Care Committee in conformance to regulatory standards established by the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC).

Cultured nodose neurones

Neurones from rat nodose ganglia were isolated and cultured as previously described (De Koninck et al. 1993; Cunningham et al. 1997; Li et al. 1997). Briefly, Sprague–Dawley rats were anaesthetized with halothane (Halocarbon Laboratories, River Edge, NJ, USA) and decapitated. Nodose ganglia were dissected, transferred to ice-cold modified L-15 medium, minced with a sharp razor blade, and incubated for 1 h at 37°C in medium supplemented with 0.1 mg ml−1 of DNAse, 1 mg ml−1 of trypsin, and 1 mg ml−1 of collagenase (all from Worthington, Lakewood, NJ, USA). The digested ganglia were triturated to dissociate neurones. The cell suspension was diluted with medium containing 5% rat serum (Atlantic Biological, Norcross, GA, USA), 2 mg ml−1 of soybean trypsin inhibitor (Worthington), 1 mg ml−1 of bovine serum albumin, and 3 mm of CaCl2, and was centrifuged. The neurones were re-suspended in modified L-15 medium supplemented with 5% rat serum and 2% chick embryo extract (Gibco, Paisley, Scotland), and plated on poly l-lysine coated cover slips. Experiments were performed at 37°C in a PDMI-2 heated bath (Harvard Apparatus, Holliston, MA, USA) within 1–3 days after dissociation.

A total of 107 neurones were used for testing mechanosensitivity: 49 (18 MS and 31 MI) for experiments with 10 μm cPGI, 37 (9 MS and 28 MI) for experiments with 1 μm of cPGI, and 21 (9 MS and 12 MI) for experiments with 10 μm of cPGI in the presence of 10 μm of tetrodotoxin (TTX). Seven neurones were used for experiments with ChTX and cPGI, 17 for experiments with linopirdine and cPGI, 5 for current clamp experiments and 12 for voltage clamp experiments. For current clamp experiments, the extracellular buffer contained (mm): NaCl (116), KCl (5.4), NaH2PO4 (1), MgSO4 (0.8), MgCl2 (1), CaCl2 (1.8), d-glucose (5.6), NaHCO3 (26), pH 7.4 (5% CO2).


Membrane potential (MP) and AP discharge were recorded with sharp microelectrodes (100–250 MΩ), pulled with a P- 2000 puller (Sutter, Novato, CA, USA); and filled with KCl (1 m) solution. Membrane conductance (MG) was calculated from the MP response to hyperpolarizing current injections of 0.1 nA for 1 s (Fig. 1C). Neuronal excitability was determined by counting the number of APs in response to mechanically induced depolarization and to electrically induced depolarizations by current injections ranging from 0.1 to 0.5 nA for 1 s before and after cPGI. Data were collected using Axoprobe-1A and Clampex 8 software and were analysed using Clampfit 8 (Axon Instruments, Union City, CA, USA) and plotted with Origin 7 software (OriginLab Corp., Northampton, MA, USA).

Figure 1.

Experimental preparation and responses to mechanical stimulation and to superfusion with cPG1
A, the experimental set-up shows a nodose neurone with a sharp microelectrode and a buffer ejecting pipette for mechanical stimulation. B, mechanical stimulation (horizontal bars) with a jet of extracellular buffer ejected under pressure of 5 psi depolarized this neurone. With a stronger stimulation of 10 psi, the depolarization was greater and triggered transient action potentials (APs). Membrane potential (MP) returned gradually to the baseline after the end of mechanical stimulation. This neurone belonged to the mechanosensitive (MS) and chemosensitive (CS) group. C, membrane conductance (MG) was calculated from the change in MP from MPo to MPinj (ΔMP) during 1 s current injection of a hyperpolarizing current of 0.1 nA. D, the beginning of carbacyclin (cPGI) superfusion is indicated by the arrow. The maximal depolarizations caused by cPGI (10 μm) occurred over a period of approximately 5 min and averaged 6.0 ± 2.4 mV in MS neurones (n= 18) and 7.8 ± 2.0 mV in mechanoinsensitive (MI) neurones (n= 31) and were not statistically different in these two groups. The response to cPGI was sustained and a wash-out period of 15 min reversed it and allowed studies in the recovery period. A dose of 1 μm of cPGI did not cause depolarization (not shown).

Most experiments were carried out in the current clamp mode to assess depolarizations and AP generation in response to mechanical, chemical and electrical stimuli. Experiments to define the effect of cPGI on M-current were performed under voltage-clamp conditions.


Studies of electrophysiological response to mechanical stimulation at a single cell level are technically challenging (Morris & Horn, 1991). The main difficulty is maintaining a tight electrical seal between a cell and a recording micropipette while the cell body is mechanically deformed. In this case, sharp microelectrodes present a significant advantage over patch clamp pipettes for several reasons. First, the area of contact between the sharp microelectrode (outer tip diameter is about 0.1 μm) and the cellular membrane is much smaller than in the case of a patch pipette (outer tip diameter is about 2 μm). Second, after impalements, the microelectrode can be retracted or moved in the lateral direction many micrometres without affecting resting MP (Brown & Flaming, 1986). Hence, the mechanical deformation of the impaled neurone does not disturb electrophysiological recording.

Neurones were mechanically stimulated by extracellular buffer ejected at various pressures from a micropipette using a pneumatic PicoPump (Model PV830) (WPI, Sarasota, FL, USA). The tip of the pipette (2 μm in diameter) was placed 10 μm from the cell surface (Fig. 1A). By puffing buffer at 5, 10, and 20 psi, we obtained graded depolarizations that were blocked selectively by gadolinium (Sullivan et al. 1997) or amiloride (Snitsarev et al. 2002). In the present experiments, however, we report data with injections at 10 psi, which we found more optimal than 5 psi to cause depolarizations that would reach the threshold of excitability and AP generation and less likely than 20 psi to dislodge the microelectrode. The 10 psi puffs gave us consistent and reversible depolarizations in MS cells (Fig. 1B) although the variability in responses among MS neurones was high. This is to be expected since the nodose neurones receive sensory signals from diverse visceral organs with variable mechanical properties. Similar to our previous study (Snitsarev et al. 2002), neurones were considered MS if the depolarization was greater than 3 mV. Approximately one-third of neurones tested were MS and all other neurones were considered MI. MS neurones generated a transient burst of APs during impalement with the sharp microelectrode and then became quiescent. Only a few neurones fired APs during the subsequent mechanically induced depolarizations in the absence of cPGI.


Neuronal excitability was assessed by measuring AP responses to depolarizing current injections (0.1, 0.2, 0.3, 0.4 and 0.5 nA for 1 s). Because of the variability in the magnitude of changes in MP during current injections among different groups of neurones, we calculated the number of APs generated as a function of the magnitude of depolarization (i.e. ΔmV) and used the normalized number of APs per millivolt as the index of neuronal excitability. An augmentation of the gain of this index of excitability over the range of current injections from 0.1 to 0.5 nA by cPGI reflected chemosensitivity. Changes in the MP (ΔmV) with each injection were measured as the difference between the averaged MP during the last 100 ms of current injection and the 100 ms before the injection and the resulting APs during the 1 s injections were counted.

Chemosensitivity and response to cPGI

In these experiments we did not use PGI2 because at physiological pH it hydrolyses within minutes to form its inactive derivative 6-keto-PGF. We used instead cPGI, which is a stable PGI2 analogue and an excellent PGI2 mimetic, even though it possesses less than one-tenth the potency of PGI2, and hence the higher doses of cPGI (Adaikan et al. 1980; Aiken & Shebuski, 1980).

cPGI (10 μm) depolarized MS and MI neurones, but it rarely generated APs on its own. In both MS and MI neurones, however, cPGI enhanced AP firing that occurred during mechanical stimulation and/or depolarizing current injections. Superfusion of cPGI at 1 μm was insufficient to cause depolarization but it enhanced excitability. Superfusion of 10 μm induced a gradual depolarization that peaked within 5 min and was maintained until the wash-out period. The enhanced excitability by cPGI was mediated by inhibition of Maxi-K but its depolarizing action was not. We therefore tested the possible role of another K+ current, the neuronal M-current, as the mediator of depolarization.

The neuronal M-current was first described in 1980 as a slowly activating, non-inactivating K+ current inhibited through stimulation of muscarinic receptors (Brown & Adams, 1980). It is voltage dependent, opens at potentials of about −60 mV and has a slow time constant for both activation and deactivation (Jensen et al. 2007). It contributes to the maintenance of resting MP. The M-current was measured in the whole cell patch-clamp configuration according to Suh & Hille (2002). MP was clamped at −20 mV and 500 ms hyperpolarizing steps to −60 mV were applied every 4 s. A slow inward current relaxation began within 50 ms following the initiation of the hyperpolarizing command and progressed during the hyperpolarizing step of 500 ms reflecting time-dependent deactivation of M-current. The effects of cPGI on this current were tested before and after oxotremorine-M (10 μm), a selective muscarinic agonist that blocks the current (Birdsall et al. 1978), or before and after linopirdine (30 μm), which inhibits the current (Aiken et al. 1996; Lamas et al. 1997; Schnee & Brown, 1998).

Several protocols were designed to define the putative ionic mechanisms involved in the sensitization of responses of nodose neurones by cPGI. We determined whether:

  • 1cPGI sensitizes the mechanosensitive DEG/ENaC channels that mediate the mechanically induced depolarization. This defined the ionic selectivity of the excitatory effect of cPGI.
  • 2The AP response to mechanical and electrical depolarizations (i.e. excitability) is augmented by cPGI (1 μm and 10 μm). This helped define a subpopulation of MS neurones and the MI neurones as ‘chemosensitive’ to cPGI.
  • 3The excitability of MI neurones differed from that of MS neurones. This helped define the much higher level of excitability and chemosensitivity of MI neurones to cPGI.
  • 4Modulation of Maxi-K mediates the chemosensitivity and increased current-induced AP generation in response to cPGI. Experiments were done before and after blockade of Maxi-K with charybdotoxin (ChTX).
  • 5M-current mediates the depolarization by cPGI. Voltage-clamp experiments were done before addition of M-current blockers oxotremorine and linopirdine.

In contrasting the magnitude of changes in MP (ΔmV) and the resulting APs during mechanical and electrical depolarizations before and after cPGI the following two precautions were taken.

Measurements from a fixed MP at −60 mV The variability in resting MP and in the depolarizing action of cPGI among nodose neurones could alter the magnitude of mechanical and electrical depolarizations and the number of generated APs. To limit this variable, a hyperpolarizing current was injected to maintain MP at −60 mV during all interventions before and after cPGI. Thus, differences in resting levels of MP could not account for altered responsiveness.

Blockade of AP generation by TTX The generation of APs tends to hyperpolarize the MP. It is difficult to estimate the direct effect of an intervention on MP (ΔmV) in the presence of increased excitability and AP generation. Therefore, the MP responses to mechanical and electrical stimulation before and after cPGI were also studied in the presence of the Na+ channel blocker TTX (10 μm) in the bathing solution.

Statistical analysis

Student's t test for paired and unpaired data and analysis of variance were used as appropriate using Origin (OriginLab Corp.) and GB-STAT (Dynamic Microsystems, Silver Spring, MD, USA). Group data are expressed as means ± standard error of the mean (s.e.m.). Logistical functions describing the relationship between changes in MP (ΔmV) and AP discharge frequency were analysed before and after chemical interventions over the range of depolarizing current injections. Differences were considered significant at P < 0.05.


cPGI (6,9α-methylene-11α,15S-dihydroxyprosta-5E,13E-dien-1-oie acid) was purchased from Cayman Chemical (Ann Arbor, MI, USA). All other reagents (amiloride, oxotremorine-M, linopirdine) were from either Sigma (St Louis, MO) or Fisher Scientific (Pittsburgh, PA, USA).


Mechanically induced depolarization of nodose neurones

Nodose neurones were isolated, impaled with sharp microelectrodes and tested for mechanosensitivity (Fig. 1A and B). They all had characteristically round somata ranging from 25 to 35 μm in diameter, and elicited a transient burst of APs upon impalement. We had shown previously that MS nodose neurones respond incrementally to increasing levels of mechanical stimulation ranging from 5 to 20 psi (Sharma et al. 1995; Cunningham et al. 1997; Sullivan et al. 1997; Kraske et al. 1998; Snitsarev et al. 2002). As explained in Methods, here we report responses to mechanical stimulation at 10 psi. These averaged 9.8 ± 1.6 mV in 36 MS neurones while 71 neurones had essentially no response (0.0 ± 0.1 mV). The latter group of neurones were considered mechanoinsensitive (MI). There were no differences between MS and MI neurones in size (33.3 ± 0.7 μm and 31.9 ± 0.6 μm, respectively), in resting MP (−49.2 ± 2.4 and −49.6 ± 1.3 mV, respectively), or in resting MG (18.5 ± 2.3 and 20.9 ± 1.7 nS, respectively).

Effect of cPGI on mechanosensitive neurones

In 18 MS neurones cPGI (10 μm) caused gradual depolarization from −48.5 ± 2.4 to −42.5 ± 2.2 mV (P < 0.001) with a corresponding decrease in MG from 18.7 ± 3.2 to 13.7 ± 2.0 nS (P < 0.01) (Fig. 1C and D). Mechanically induced depolarizations were not augmented by cPGI even when neurones were tested at a constant MP of −60 mV (n= 18) (to offset the depolarizing effect of cPGI) or after TTX (n= 9) (Table 1).

Table 1.  Changes in membrane potential (ΔMP in mV) during mechanical stimulation (10psi) before and after cPGI (10 μm)
 nBefore cPGIAfter cPGI
  1. Data represent means ±s.e.m. Depolarizations (ΔmV) of MS nodose neurones during mechanical stimulation were measured at their resting membrane potential (RMP), at a constant MP of −60 mV, and after tetrodotoxin (TTX 10 μm). Hyperpolarizing currents were injected to maintain neurones at −60 mV before mechanical stimulation and offset the depolarization by cPGI. Mechanically induced depolarizations were unchanged (P > 0.05) after cPGI when measured at the RMP, or at −60 mV, or after TTX. These 18 neurones include both MS and MS/CS ones. Fig. 2 shows the separate responses to mechanical stimulation at RMP.

ΔmV (RMP)1812.3 ± 3.011.6 ± 2.8
ΔmV (−60 mV)1810.1 ± 2.910.5 ± 2.9
ΔmV (TTX) 9 8.4 ± 1.6 7.9 ± 1.8

In 11 of these 18 neurones (Group A) mechanical depolarization was small, averaging 6.4 ± 2.1 mV from a control potential of −60 mV; it did not increase with cPGI, nor did it generate APs either before or after cPGI. In the remaining seven neurones (Group B), mechanical depolarization was significantly greater (23.3 ± 5.2 mV) but still did not increase with cPGI. However, AP generation increased significantly in this subgroup after cPGI (Fig. 2). Thus cPGI did not enhance the mechanically induced depolarization in either subgroup, but it did enhance AP generation in the subgroup of seven, indicating increased excitability by cPGI and that these neurones are also chemosensitive (CS). Increased excitability of this subgroup was evident also from the results obtained with electrical depolarizations during current injections.

Figure 2.

Responses to mechanical stimulation (10 psi for 3 s) before and after cPGI (10 μm)
Tracings represent depolarizations (ΔmV) and action potential (AP) spikes of 3 individual neurones in response to mechanical stimulation from their resting membrane potential. Bar graphs show means ±s.e.m. of changes in membrane potential (ΔmV), and the number of AP spikes generated during mechanical stimulation before (Control – open bars) and after cPGI (filled bars). Three groups of neurones are represented. Two groups, A and B, were mechanosensitive (MS) and one group, C, was mechanoinsensitive (MI). Group A (MS) neurones (n= 11) depolarized less than Group B and did not generate AP spikes before or after cPGI. Group B (MS/CS) (n= 7) depolarized significantly more and generated AP spikes. cPGI did not increase the magnitude of mechanically induced depolarizations in either A or B but enhanced the AP generation in B. *Significant difference (P < 0.05). Because of their enhanced APs after cPGI, group B neurones were considered also chemosensitive (CS). Group C (MI/CS) neurones were mechanoinsensitive (MI) before cPGI and remained MI after cPGI. cPGI sensitized their AP responses to electrical depolarization hence their label as CS (see Fig. 3).

A sigmoidal function reflecting neuronal excitability was noted between the number of APs and the progressive increments in electrical depolarizations (Table 2 and Fig. 3). In Group A (MS) neurones, electrical depolarizations were markedly increased by cPGI but without a significant increase in the maximum gain of the AP frequency response curve. In contrast, Group B (MS/CS) neurones showed an opposite response, i.e. there were no significant increases in depolarizations after cPGI, but there were greater increments in AP frequency resulting in a significant increase in the maximum gain with a shift of the curve to the left. This enhancement of excitability by cPGI indicates that MS neurones in Group B are also CS or bimodal (MS/CS) whereas MS neurones in Group A are not CS (unimodal). Responses returned to control levels after washout of cPGI (Fig. 4).

Table 2.  Current-induced depolarizations and corresponding APs During 1 s injections of 0.1–0.5 nA in three groups of neurones (MS, MS/CS, and MI/CS) before and after cPGI (10 μm)
Current (nA)Before cPGIAfter cPGI
  1. Data represent means ±s.e.m. of changes in membrane potentials (ΔMP in mV) and corresponding action potentials (APs) in spikes s−1. Significant differences (P < 0.05) between groups were calculated for ΔMP and APs using logistical equations and ANOVA. *Significant difference of MI/CS and MS/CS from MS; †significant difference of MI/CS from MS/CS; §significant differences of responses after cPGI compared to before cPGI; NS, no statistical difference from MS. The differences in depolarizations between groups (MS versus MI) before and after cPGI were not altered by TTX (data not shown) 1Derivatives of the AP firing frequency/ΔmV sigmoidal curves obtained during current injections in each group were calculated before and after cPGI. The maximum gain for each group and the ΔmV at which that gain was recorded are shown. See Fig. 3 for portrayal of the curves and their derivatives.

 0.114 ± 3 8 ± 310 ± 119 ± 411 ± 313 ± 1
 0.227 ± 7 16 ± 6 17 ± 237 ± 920 ± 524 ± 3
 0.344 ± 1124 ± 8 23 ± 3 69 ± 1628 ± 836 ± 5
 0.460 ± 1832 ± 1232 ± 4 96 ± 31 38 ± 1149 ± 7
 0.583 ± 2142 ± 1740 ± 6138 ± 39 51 ± 1964 ± 9
 0.10 ± 00 ± 0 1 ± 1 2.5 ± 2.3 3.2 ± 2.714 ± 4
 0.20.9 ± 0.80.6 ± 0.6 5 ± 2 5.7 ± 4.3 6.0 ± 4.321 ± 5
 0.34.4 ± 2.62.2 ± 1.2 8 ± 3 9.9 ± 5.2 8.6 ± 5.028 ± 7
 0.46.8 ± 4.04.0 ± 2.110 ± 415.6 ± 7.310.4 ± 6.333 ± 7
 0.57.2 ± 4.15.2 ± 2.713 ± 515.8 ± 7.412.8 ± 7.339 ± 8
 Maximum gain0.250.240.450.200.342.16
 (dAP/d ΔmV)(@38 mV)(@24 mV)(@18 mV)(@31 mV)(@10 mV)(@1 mV)
  (NS)(*†) (*§)(*†§)
Figure 3.

Current-induced depolarizations and corresponding AP generations in three groups of nodose neurones: MS, MS/CS and MI/CS
(Refer to group data in Table 2.) The upper tracings from 3 representative neurones show depolarizations and AP firings in response to current injections of 0.3 nA for 1 s before (control) and after cPGI. Resting MPs were maintained at −60 mV. The increase in APs after cPGI in an MS neurone (left tracings) was induced by a pronounced increase in depolarization. In the MS/CS neurone (middle tracings), the increase in APs after cPGI occurred without a significant increase in depolarization. In MI/CS neurone (right tracings) the increase in APs was significantly greater than in the other two neurones before cPGI and much greater after cPGI. Analyses of group data shown in Table 2 are presented here in two graphs under each of the 3 groups: MS, MS/CS and MI/CS. The upper graphs portray the sigmoid curves calculated from the AP spikes during the depolarizations induced by current injections. The circles and dashed lines represent the control mean values of ΔMP (mV) and the corresponding mean values of AP obtained during each of the 5 current injections (0.1, 0.2, 0.3, 0.4 and 0.5 nA) before cPGI (see Table 2 for s.e.m. of each value). The squares and continuous lines represent the values obtained with the same interventions after cPGI (10 μm). The lower graphs portray the derivatives dAPs/dΔMP (number s−1 mV−1), obtained in each group before (dashed lines) and after (continous line) cPGI. The following significant differences with respect to the effect of cPGI were determined by analyses of logistical equations and ANOVA (see also Table 2). In the MS group, ‘control’ depolarizations were greater than in both other groups (P < 0.05) and cPGI caused a significantly greater increase in the magnitude of depolarization without an increase in the maximal gain in APs. In the MS/CS group, cPGI did not increase the magnitude of depolarization but it caused a significant increase and a shift to the left of the maximal gain. In the MI/CS group, the control gain was higher than the other 2 groups (P < 0.05) and cPGI caused a most pronounced enhancement of the maximal gain of APs with a significant shift to the left.

Figure 4.

Recovery from the enhanced responses to current injections (0.5 nA) caused by cPGI
Bars indicate means ±s.e.m. of ΔMP (mV) and APs in response to 0.5 nA current injections in MS (n= 11), MS/CS (n= 7) and MI/CS (n= 31) neurones that were obtained before cPGI (open bars), during cPGI (filled bars), and during recovery following the washout period (shaded bars). Although we show responses to only 0.5 nA for simplicity, differentially enhanced responses to 0.1, 0.2, 0.3 and 0.4 nA by cPGI were qualitatively similar (as shown in Table 2). *Responses in MS/CS and MI/CS neurones were significantly different (lesser for ΔMP and greater for APs) from those seen in MS neurones; †greater AP responses in MI/CS neurones than MS/CS neurones; §significant increases with cPGI compared to corresponding controls in each group which were reversed back to control values after washout periods of 15 min. Absence of symbols indicates lack of significant differences.

Effect of cPGI on mechanoinsensitive neurones

cPGI (10 μm) depolarized 31 MI neurones from −51.1 ± 1.4 to −43.3 ± 2.0 mV (P < 0.001) with corresponding decreases in MG from 19.6 ± 2.7 to 11.4 ± 1.7nS (P < 0.01). These effects tended to be greater than those seen in MS neurones but they did not provoke any response to mechanical stimulation.

Electrical depolarizations from a constant level of −60 mV in these MI neurones were significantly less than those seen in MS neurones and generated a greater number of APs than those seen in MS and MS/CS neurones (Figs 3 and 4 and Table 2). The excitability of MI neurones was further enhanced by cPGI as indicated by a marked increase in the maximum gain of the curve and its shift to the left (Fig. 3). These results reflect a significantly greater degree of chemosensitivity of MI neurones than seen in the bimodal MS/CS neurones. Responses returned to control levels after washout of cPGI (Fig. 4).

In another group of 28 MI neurones, a smaller dose of 1 μm cPGI had no significant effect on resting MP and MG (data not shown). That dose did provoke significant increases in AP responses during current injections but without the significant increases in the depolarizing potentials that were seen with 10 μm cPGI (Fig. 5). Thus chemosensitivity was evident from the excitatory effect of cPGI on AP generation which was observed in the absence of its depolarizing effect (with 1 μm cPGI) as well as at a constant MP of −60 mV (with 10 μm cPGI).

Figure 5.

Dose-related enhancement of responses to electrical depolarization by 1 μmversus 10 μm cPGI
Bars indicate means ±s.e.m. of the increases in ΔMP (mV) and in APs in response to incremental current injections that were provoked by 1 μm cPGI (open bars) and by 10 μm cPGI (black bars) over corresponding responses seen in the absence of cPGI. The small dose of cPGI (1 μm) enhanced the AP responses (*P < 0.05) without causing a significant increase in ΔMP (NS) over the changes observed in the absence of cPGI. The larger dose (10 μm) caused a more pronounced enhancement of APs (*P < 0.05) with a significant increase in ΔMP (*P < 0.05) over control values without cPGI.

Role of Maxi-K in mediating the chemosensitivity to cPGI

We had reported earlier that cPGI inhibits the large conductance Ca2+-activated potassium channel (Maxi-K) in aortic baroreceptor neurones (Li et al. 1997). We reasoned that this action may contribute to the enhanced AP generation by cPGI in response to electrical depolarizations and be partly responsible for the chemosensitivity to cPGI in nodose neurones.

ChTX (100 nm), a blocker of Maxi-K, did not change significantly resting MP (ΔMP =−2.0 ± 4.8 mV, n= 8, P > 0.05) and conductance (ΔMG = 5.5 ± 8.0 nS, n= 8, P > 0.05) but increased significantly the AP firing frequency in response to depolarizing current injections without increasing the magnitude of depolarization significantly (Fig. 6, Table 3).

Figure 6.

Effect of ChTX and cPGI on the ΔMP (mV) and APs responses to current injections
The tracings show representative responses to 3 current injections of 0.4 nA over 1 s in the same neurone from a constant MP of −60 mV. ChTX (100 nm) increased AP generation from ‘control’ of 4 spikes to 11 spikes without an increase in current-induced depolarization. After the additional superfusion of cPGI (10 μm), current-induced depolarizations increased but without a further increment in APs (10 spikes). Table 3 includes responses in 8 neurones to graded current injections that indicate that ChTX enhances APs and abrogates further increases in APs by cPGI despite the enhanced effect on depolarization (ΔMP).

Table 3.  Current-induced depolarizations and corresponding APs during 1 s injections of 0.1–0.5 nA before (control) and in the presence of ChTX (100 nm) and with the subsequent addition of cPGI (10 μm) (n= 8)
Current (nA)Depolarization ΔMP (mV)APs (spikes s−1)
ControlChTXcPGI after ChTXControlChTXcPGI after ChTX
  1. Data represent means ±s.e.m. of ΔMP in mV and APs in spikes s−1 in response to current injections with MPs held at −60 mV prior to current injections. Compared to Control the AP generation with current injections was increased significantly by ChTX (*P < 0.05) without a corresponding increase in depolarization (NS). The addition of cPGI in the presence of ChTX caused significantly greater depolarizations (P < 0.05) with current injections than in Control (*) and ChTX alone (†). But for equivalent changes in depolarization, the AP responses were not enhanced by the addition of cPGI to ChTX compared with ChTX alone. Using logistical function analyses for comparison of regression curves, responses to ChTX were significantly greater than Control (F value = 7.988 and P= 0.0401) but responses to cPGI + ChTX were not different than those with ChTX alone (F value = 3.548 and P= 0.129).

0.1 8.4 ± 2.2 9.7 ± 1.412.1 ± 2.4 0.5 ± 0.5 2.8 ± 1.3 6.4 ± 3.4
0.213.1 ± 3.615.9 ± 3.021.5 ± 4.4 4.6 ± 3.3 8.9 ± 4.311.4 ± 5.4
0.317.8 ± 5.022.4 ± 4.832.1 ± 7.6 7.8 ± 4.313.5 ± 5.416.9 ± 6.8
0.423.2 ± 7.529.3 ± 7.6 43.7 ± 12.110.0 ± 5.617.3 ± 6.321.6 ± 7.8
0.528.4 ± 9.7 37.4 ± 11.1 54.5 ± 15.412.4 ± 6.420.0 ± 7.026.9 ± 8.5
  (NS)(*†) (*)(*)

The addition of cPGI (10 μm) in the presence of ChTX depolarized the cells by 5.0 ± 4.6 mV, decreased MG by 15.6 ± 8.1 nS (n= 8, P < 0.05), enhanced the current-induced depolarizations but did not increase the AP firing over what was observed with ChTX at the same level of ΔmV (Table 3, Fig. 6). Thus, suppression of Maxi-K mediates the enhancement of AP responses to current injections caused by both ChTX and cPGI but does not mediate the enhanced cPGI-induced depolarization. This suggested that an alternative conductance is responsible for cPGI-induced depolarization.

Role of M-current in mediating the depolarizing effect of cPGI

In current-clamp experiments oxotremorine-M did not depolarize the isolated nodose neurones (not shown), whereas linopirdine did by 17.6 ± 5.8 mV (n= 5, P < 0.05). Linopirdine also decreased MG by 7.4 ± 3.1 nS (n= 5, P < 0.05) (Fig. 7). In the presence of linopirdine, and with the MP held at −60 mV, the addition of cPGI had no further effect on MP and MG suggesting that inhibition of the M-current mediates the cPGI-induced depolarization (Fig. 7).

Figure 7.

Effects of cPGI and linopirdine on MP and MG under current clamp and on M-current under voltage clamp conditions
Current clamp: linopirdine (Lin) depolarized the neurone by 20.1 mV and decreased MG by 7.7 nS. When MP was adjusted back to −60 mV by a hyperpolarizing current injection (−0.58 nA) (see arrow), the addition of cPGI in the presence of Lin had no further effect on MP and MG. During Lin applications, occasional spontaneous APs could be observed. Negative deflections are responses to −0.1 nA current injections of 0.1 s used to calculate MG, which was decreased by Lin. Voltage clamp (M-current): A, tracings from one neurone show current excursions produced by a series of four, 500 ms hyperpolarizing voltage steps from a holding potential of −20 mV to −60 mV obtained before (Control), after the addition of cPGI (10 μm), after further addition of linopirdine (30 μm) (cPGI + Lin), and after the washout period (Wash). B, tracings show a tenfold magnification of the bottom part of the first and second current excursions in A while the MP was held at −60 mV (during the 500 ms duration of the hyperpolarizing voltage step) in Control (IM+Ires), and after cPGI (Ires). An electronically derived subtraction of the current after cPGI from the control current (Control–cPGI) is also shown (IM). C and D are similar to tracings A and B except that they were acquired from a different neurone to which Lin was added first followed by cPGI and the washout period. At the holding potential of −20 mV before the hyperpolarizing step, the M-current was partly activated contributing to a steady outward current (A and C, Control), which was reduced dramatically by cPGI (A, cPGI) or by linopirdine (C, Lin). During the 500 ms hyperpolarizing step at −60 mV, the deactivation of the M-current resulted in a net slow inward current relaxation (IM+Ires). After cPGI (10 μm) (A and B, cPGI) there was a reversal of the slow inward current relaxation during the hyperpolarizing step giving rise instead to a gradual outward current (Ires). The addition of linopirdine in the presence of cPGI had no significant additional effect on the current. Subtraction of the gradual outward current (Ires) from the slow net inward current relaxation in control (IM+Ires) resulted in a total inward current (IM) of over 30 pA. When linopirdine was applied first (n= 6) (C and D) the inward current relaxation in control (IM+Ires) was reversed (C and D Lin), and was not affected by further addition of cPGI in the presence of linopirdine (C, Lin + cPGI). The washout period restored both the steady outward current and the slow inward current relaxation (A and C, Wash).

In voltage-clamp experiments, the 500 ms hyperpolarizing step from −20 to −60 mV deactivates the M-current resulting in a net slow inward current relaxation (IM/Ires) averaging 25.0 ± 5.9 pA (n= 12) (Fig. 7, Control tracings). After cPGI (Fig. 7A and B) there was a reversal of the slow inward relaxation to a gradual outward current (Ires) of 18.8 ± 6.2 pA (n= 6) which was not altered by further addition of linopirdine (15.9 ± 5.5 pA, n= 6). The electronically derived total current blocked by cPGI (IM) averaged 47.3 ± 14.5 pA (n= 6) (Fig. 7B). Conversely when linopirdine (Lin) was applied first in six other neurones (Fig. 7C and D) the inward current relaxation (IM+Ires) which was 23.0 ± 7.4 pA was reversed to an outward current (Ires) of 32.9 ± 17.4 pA, which was not altered significantly by the further addition of cPGI. The derived total current blocked by linopirdine averaged 56.0 ± 12.1 pA (Fig. 7D).

Thus the additions of linopirdine after cPGI or cPGI after linopirdine did not alter the Ires significantly (P > 0.05 for both groups of n= 6 each) indicating that cPGI inhibits the outward M-current known to be blocked by linopirdine and vice versa. Oxotremorine-M (10 μm) had no effect on the slow inward current relaxation (not shown).


The ion channels involved in depolarization of nodose neurones may be mechanically gated, voltage gated or activated through ligand coupled receptors.

Our results indicate the following. (1) The excitatory effect of cPGI on nodose neurones is selective. The mechanically induced depolarization, which is the first ionic event in mechanoelectrical transduction, is not sensitized by cPGI whereas AP generation resulting from mechanically or electrically induced depolarizations is enhanced by cPGI. (2) cPGI activates nodose neurones by inhibiting two K+ channels. One is Maxi-K which may determine AP generation and neuronal excitability and the other is M-current which mediates neuronal depolarization. (3) Nodose neurones may be separated into three groups as outlined below.

(a) ‘Unimodal’ mechanosensitive In a significant number of MS neurones, mechanical stimulation causes a modest depolarization and does not generate APs even with cPGI. Moreover, despite significant increases in electrical depolarization after cPGI the normalized AP firing frequency is not enhanced by cPGI. These neurones are cPGI insensitive and predominantly ‘unimodal’ mechanosensitive.

(b) ‘Bimodal’ mechanosensitive and chemosensitive In 40% of MS neurones mechanical stimulation causes a pronounced depolarization, and the APs generated either by mechanical or electrical depolarizations are enhanced significantly by cPGI. Thus these MS neurones are considered also chemosensitive (CS) to cPGI.

(c) ‘Unimodal’ chemosensitive The mechanoinsensitive (MI) neurones generated significantly more APs during current injections than MS neurones. They were very sensitive to cPGI which enhanced their AP responses severalfold more than in MS neurones and they never exhibited mechanosensitivity.

The discussion will focus on mechanisms of sensory transduction in vagal afferents and the use of nodose neurones to assess mechano- and chemosensitivity; the mechanisms that determine sensitization of nodose neurones to PGI2; and the physiological and pathological significance of the result.

Mechanoelectrical transduction in nodose neurones

The majority of baroreceptor and vagal afferents of nodose neurones are unmyelinated fibres that are much more responsive to chemical factors than myelinated fibres (Higashi et al. 1982). However, either type of nerve fibres may be mechanosensitive. An important concept from the results of this study is that mechanosensitivity or chemosensitivity should not be defined simply in terms of AP responses of a particular nerve fibre recorded from a site far from where the mechano- or chemotransductions occur. The determinants of these responses need to be defined in terms of ionic conductances.

The mechanotransduction process involves a dual channel activation in tandem. First is the mechanosensitive depolarization that initiates the ‘receptor potential’ or ‘generator potential’. Second is activation of voltage-gated ion channels that initiate the spike firing which is eventually recorded as the nerve activity. Since the terminals are embedded in vascular or visceral tissues or muscles, the recorded activity is subject not only to mechanical deformation but also to modulation by paracrine factors from the endothelium or vascular muscle or other cells. Thus a defect in baroreceptor nerve activity may be caused by impairment of specific neuronal ion channels and/or their abnormal regulation by chemical factors. Some neuronal channels may be components of the primary transducer and others may be voltage gated and secondarily activated by the depolarizing potentials.

The ability to distinguish the two separate ionic events is necessary to better understand the basic mechanisms leading to impairment of baroreceptor activity and its pathological consequences. Unfortunately, the small size and complex architecture of the terminals makes the recording of the mechanosensitive potential from these nerve terminals practically impossible (Matsuura, 1973). Over 10 years ago we first used aortic baroreceptor nodose neurones in culture to characterize mechanosensitive conductances and single channel units that were blocked by Gd3+ (Hajduczok et al. 1994; Cunningham et al. 1995, 1997; Sharma et al. 1995; Kraske et al. 1998). We then found in these somata, and more importantly in their terminals, subunits of the DEG/ENaC superfamily of cationic, non-voltage-gated, non-ligand-gated, amiloride sensitive, mechanosensitive channels (Drummond et al. 1998; Snitsarev et al. 2002; Chapleau & Abboud, 2004). At present a subunit of the acid sensing ion channels subfamily (ASIC2) has been described as a mechanosensor in mammalian peripheral sensory nerve, and our preliminary work with genetic deletion of this subunit ASIC2 reveals a significant impairment of baroreceptor afferents and the baroreflex (Price et al. 2000; Ma et al. 2001a; Chapleau & Abboud, 2004).

Our first conclusion from this study is that cPGI does not sensitize mechanosensitive conductances in nodose neurones. The justifications for extending observations made at the perikaryon to the sensory nerve terminal have been addressed in the past. These have included the fact that specific ligand receptors and ion channel proteins are common to both the sensory endings and the cultured nodose neurones (Fowler et al. 1985; Stansfeld et al. 1986; Christian et al. 1989; Drummond et al. 1998; Kraske et al. 1998). We have also shown that Gd3+ and the amiloride analogue, benzamil, block the carotid sinus nerve activity in vivo as well as the mechanically induced depolarization or inward currents of nodose baroreceptor neurones in culture (Hajduczok et al. 1994; Cunningham et al. 1997; Sullivan et al. 1997; Drummond et al. 1998; Snitsarev et al. 2002). Consequently the cell body has served us and others as a useful model of the nerve terminal allowing the separate identification of mechanosensitive as well as chemosensitive conductances that would not be accessible in the terminals (Undem & Weinreich, 1993; Weinreich et al. 1995; Marrion, 1997; Schild & Kunze, 1997; Sullivan et al. 1997; Lancaster & Weinreich, 2001; Snitsarev et al. 2002, 2005b; Wladyka & Kunze, 2006).

Our observations have focused on the mechanosensitivity mediated through DEG/ENaC ion channels that are blocked by amiloride. In recent years the transient receptor potential (TRP) channels have been implicated in mechanotransduction (Lin & Corey, 2005; O'Neil & Heller, 2005). TRPV1 and several TRPC proteins are expressed in sensory neurones and in rat nodose ganglia (Helliwell et al. 1998; Glazebrook et al. 2005). They are not sensitive to amiloride and may certainly contribute to mechanoelectrical transduction. In our earlier studies there is a residual activation of the carotid sinus nerve even after 10 μm of benzamil (Drummond et al. 1998). Similarly, in the isolated nodose neurones, although amiloride (1 μm) reduced significantly the mechanosensitive depolarization and AP generation (Snitsarev et al. 2002), there was still a residual depolarization that might represent activation of TRP channels.

Specificity of amiloride blockade A characteristic of the DEG/ENaC conductance has been its blockade by amiloride. The mechanically induced Ca2+ transient through DEG/ENaC is blocked by 1 μm of amiloride. This dose is insufficient to block voltage-gated Ca2+ channels (Tang et al. 1988). At an even lower dose of 100 nm it inhibits the mechanically induced non-voltage-gated Ca2+ transient (Drummond et al. 1998). On the other hand, Gd3+, which is another blocker of mechanosensitivity, is a more potent blocker of voltage-gated Ca2+ channels and thus would be less specific than amiloride.

Diversity of mechanosensitivity We found that MS nodose neurones varied in the magnitude of their depolarization in response to the mechanical stimulus. This may reflect the fact that their sensory nerve endings supply a variety of organs. Some may trigger continuously and vigorously mechanically activated conductances, e.g. the aortic arch and the cardiac vagal afferents, while others may induce slow and less intense mechanical deformation of the nerve endings. The former may have a greater expression of the MS conductances. This is suggested by the marked differences in mechanically induced depolarizations (23.3 ± 5.2 mV versus 6.4 ± 2.1 mV) and corresponding increases in conductances (6.1 ± 2.9 nS versus 2.6 ± 1.2 nS) seen in the two groups of MS neurones. It is also reasonable to anticipate that different mechanosensory conductances besides DEG/ENaC may be involved in mechanoelectrical transduction.

Distribution of the three groups of neurones We can also speculate that the three groups of nodose neurones that we were able to identify receive signals from different terminals in different tissues and organs. Thus the relative distribution of the predominantly MS, the predominantly CS and the bimodal neurones may be different depending on the origin of their sensory terminals in a particular organ or tissue. We have previously reported for example that over 90% of labelled aortic baroreceptor neurones in nodose ganglia are mechanosensitive (Drummond et al. 1998), whereas only 60% of the non-labelled neurones are mechanosensitive. Veelken et al. (2003) have reported that ∼73% of cardiac vagal afferent fibres in rat responded to both left ventricular distension (mechanical) and phenylbiguanide (chemical) injected into the pericardial sac and were classified as bimodal whereas 9 of 57 fibres were solely mechanosensitive and 6 of 57 were solely chemosensitive. Similar studies on vagal afferents from mouse stomach and oesophagus by Page et al. (2002) indicate a different distribution with two types of MS fibres with approximately 50% responding to chemical stimuli. A third type of fibre was mechanosensitive but responded only to bile.

These results emphasize the notion that mechanosensitivity and chemosensitivity are better defined in terms of specific conductances which could differ in distribution, density and sensitivity in various terminals. There is a need to focus future studies on specific populations of nodose neurones.

Chemosensitivity of nodose neurones to cPGI

Of all the putative modulators of nodose neuronal activity, we have chosen PGI2 because of several indications that it is significantly involved in the functional regulation of vagal afferent activity especially from baroreceptors and cardiac vagal afferents. We had demonstrated that PGI2 enhances carotid sinus nerve activity, that it is released from the carotid sinus during its mechanical activation, that indomethacin decreases BR activity, that PGI2 may be involved in baroreceptor resetting, and that in hypertensive animals its reduced release may contribute to the decreased baroreceptor activity (McDowell et al. 1989; Chen et al. 1990; Xie et al. 1990). We have also shown in cultured nodose neurones that PGI2 is an autocrine regulator of the recovery from post-excitatory inhibition (Snitsarev et al. 2005b). Others have implicated its involvement in sensitization of cardiac vagal afferents in heart failure, coronary occlusion, and the myocardial ischaemia–reperfusion response (Zucker et al. 1989; Brandle et al. 1994; Schultz, 2001). It is certainly true that several other chemicals including bradykinin, ATP, substance P and serotonin may be involved and their effects may vary in various vagal afferents, e.g. from the heart, the gut or the baroreceptors.

We also recognize that our interpretation of what constitutes chemosensitivity needs further clarification. First, chemosensitivity was not defined by the ability of cPGI by itself to generate APs in the nodose neurones in culture. In our study, 10 μm cPGI caused a short train of APs in only two out of a total of 49 neurones (18 MS and 31 MI). Although we and others have shown that activity in vagal and carotid sinus afferent fibres in vivo may reveal AP firing with several chemicals including PGI2, serotonin, ATP, histamine, bradykinin, thromboxane, PGE2, endothelin, leukotrienes, etc. (Coleridge et al. 1964; Christian et al. 1989; Chen et al. 1990; Xie et al. 1990; Chapleau et al. 1992; Wang et al. 1993; Armour et al. 1994; Ustinova & Schultz, 1994), most of these agents do not cause sufficient depolarization of nodose neurones in culture to trigger APs by themselves except for serotonin (Higashi et al. 1982; Leal-Cardoso et al. 1993; Undem & Weinreich, 1993). The dispersion and isolation process may increase the threshold for AP generation in vitro. However, once an initial depolarization was triggered, in vitro, with mechanical stimulation or current injection, the chemosensitivity to cPGI manifested itself with a significant enhancement of AP generation.

Chemosensitivity can be clearly uncoupled from mechanosensitivity. We could not define it in terms of an enhancement of the mechanically gated conductances and mechanically induced depolarizations because we could not demonstrate any such enhancement by cPGI. Although cPGI did enhance significantly the electrical depolarization in some (unimodal) MS neurones, their AP firing remained proportionate to the depolarization without evidence of sensitization or increase in the gain of AP firing frequency. However, in other MS neurones (bimodal) distinct chemosensitivity and enhanced gain of AP firing frequency were evident without any enhancement of mechanical or electrical depolarization by cPGI (Figs 3 and 4, Table 2). Moreover, chemosensitivity to cPGI was pronounced in MI neurones in the absence of any evidence of mechanosensitivity. The gain of AP firing frequency was dramatically increased by cPGI in these neurones with minimal or no increase in electrical depolarization. They were most excitable and predominantly or solely CS.

These observations are not inconsistent with the notion that PGI2 enhances baroreceptor nerve activity or mechanically activated vagal afferent activity as mentioned above in vivo. The reason is that PGI2 enhances nerve activity not by virtue of sensitization of the mechanically activated DEG/ENaC conductances but by enhancing AP generation as a result of blockade of voltage-gated Maxi-K conductances. Thus chemosensitivity of a baroreceptor or vagal afferent fibre is dependent on whether that nerve fibre (or neurone) has voltage-gated conductances that respond to the particular chemical and induce APs (in our case Maxi-K inhibited by cPGI).

Effect of ChTX and cPGI on Maxi-K channels Our prior work with voltage clamped neurones indicates that cPGI blocks the large conductance Ca2+ activated K+ current Maxi-K (Li et al. 1997). We did not know whether inhibition of Maxi-K by cPGI contributed to the augmented depolarization, or the enhanced AP generation, or both during current injections. The present results with current clamped neurones indicate that selective blockade of Maxi-K by ChTX enhanced the AP generation during depolarizing current injections without enhancing the magnitude of depolarization. They also indicate that once Maxi-K is blocked with ChTX any additional cPGI-mediated increase in gain of the AP frequency response to depolarization is abrogated (Table 3, Fig. 6), yet the enhanced electrical depolarization by cPGI is maintained. The preserved cPGI depolarization led us to postulate that the M-current may contribute to it.

Inhibition of M-current by cPGI The results indicate that cPGI causes depolarization and inhibits M-currents in nodose neurones. Both these actions are inhibited by linopirdine (30 μm), a known direct blocker of the M-current (Aiken et al. 1996; Lamas et al. 1997). Both the M-current and the Maxi-K current are important determinants of neuronal excitability (Wang & McKinnon, 1995; Wang et al. 1998a,b; Ma et al. 2006). First described in sympathetic neurone (Brown & Adams, 1980), the M-current was subsequently identified in central neurones (Marrion, 1997; Shapiro et al. 2000). Activation of muscarinic receptors inhibits the current. Several agonists inhibit the M-current and increase excitability (Suh & Hille, 2002). To our knowledge, during the performance of these studies the presence of the M-current in nodose neurones had not been reported. However, recently Wladyka & Kunze (2006) were first to demonstrate that KCNQ-M currents contribute to the resting membrane potential in rat visceral sensory neurones from the nodose ganglia. We now confirm their finding that M-currents are present in nodose neurones and show their inhibition by cPGI. The absence of an effect of oxotremorine, which is a muscarinic agonist that suppresses M-currents, may reflect the absence of muscarinic receptors in nodose neurones. Several downstream messengers responsible for inhibition of the M-current have been considered (Brown et al. 1989; Marrion, 1997; Brown & Yu, 2000; Suh & Hille, 2002). In a recent review Jensen et al. (2007) describe the modulation of various subtypes of the Kv7 family of K+ channels by a number of factors including PKA and PKC, phosphatidyl inositol phosphate (PIP2), G-protein coupled receptors, changes in cell volume, intracellular Ca2+ and muscarinic receptors. The M-channel (KCNQ1) has been linked to important K+ conductances in human disease (Jespersen et al. 2005).

Variation in neuronal excitability and sensitivity to cPGI

Nodose neurones were separated into three groups based on their sensitivity to mechanical stimulation, their ability to generate APs in response to graded levels of depolarizing current injections, and the degree to which cPGI enhanced their excitability and AP frequency response to depolarizations. In one group of mechanosensitive nodose neurones the mechanically induced depolarizations were larger and generated APs that were enhanced significantly by cPGI. Moreover, the maximal gain of AP firing frequency in response to electrical depolarizations was also enhanced significantly by cPGI. We have referred to this group as MS and CS, i.e. bimodal (Fig. 2). In contrast, in a second group of MS neurones, mechanical depolarization was only modest and did not generate APs even after cPGI. There was another significant difference in this group; the magnitude of electrical depolarization was significantly greater and was enhanced even more after cPGI, yet the maximal AP frequency response to depolarization was not enhanced by cPGI. We concluded that this MS group is predominantly unimodal and not CS. We speculate that the lack of chemosensitivity to cPGI and the failure of cPGI to enhance AP firing frequency may represent an impairment of Maxi-K. Conversely, the greater depolarization with cPGI in this may reflect greater M-current sensitivity. One might argue that the fact that cPGI depolarized those neurones and enhanced their electrical depolarization with current injections is an indication of chemosensitivity. As mentioned above, however, we chose instead to define chemosensitivity to cPGI in terms of enhanced maximal gain of AP firing frequency response to depolarization with more AP firing from the same or lesser change in MP.

The mechanoinsensitive neurones formed the third group. Their remarkable feature is their high levels of excitability and chemosensitivity. They exhibit a significantly larger number of APs than MS neurones and a very high maximal level of gain of the AP frequency response at relatively low levels of depolarization after cPGI. In contrast to MS neurones we speculate that there is a dominance of Maxi-K in MI neurones that may mediate the exaggerated APs and a reduced M-current sensitivity that may explain the lesser depolarization. Another important feature of the MI/CS neurones is that despite their high neuronal excitability they do not exhibit any mechanosensitivity even after addition of cPGI.

Physiological relevance

Our studies on nodose neurones do not represent a specific group of sensory afferents, and it has been reported that a majority of vagal afferents to the nodose neurones are unmyelinated fibres originating in abdominal viscera (Higashi et al. 1982). With that caveat we wish to review the functional relevance of our findings in terms of activation of baroreceptors and vagal afferents in hypertension, myocardial ischaemia and heart failure where the suppression of mechanosensitivity contributes to a detrimental enhancement of sympathetic nerve activity and neuro-humoral drive (Thames et al. 1978; Thames & Abboud, 1979; Brandle et al. 1994). The reason for the decrease in mechanosensitivity in these disease states is unclear. It could be caused by a defect in the mechanosensitive conductance, and if so the likelihood of enhancing that conductance by pharmacological means may be restricted in view of our finding of the ineffectiveness of PGI2 on this conductance. A more systematic assessment of potential modulators of mechanosensitivity and a determination in animal models of the degree of expression of the channel proteins and their functional responsiveness in isolated systems are necessary. For example, in preliminary studies we have found that a majority of aortic BR nodose neurones of the spontaneously hypertensive rat fail to depolarize in response to mechanical stimulation (Snitsarev et al. 2005a). A failure of expression of the mechanosensitive channels may certainly be a factor in their decreased baroreceptor sensitivity. Suppressed mechanosensitivity may also be a function of the absence of a sensitizing or the presence of an inhibitory chemical factor that modulates AP generation through Maxi-K and reduces the firing of the terminal despite an intact mechanosensitive conductance.

A second important functional perspective relates to the large number of mechanoinsensitive CS nodose neurones that are excitable and very sensitive to cPGI. In a clinical state where the mechanosensitive conductance at sensory terminals is suppressed and results in a detrimental disinhibition of sympathetic efferent activity, the enhanced activation of the chemosensitive conductances at the same or other sensory terminals by endogenous PGI2 may restore the sympathoinhibitory afferent signals from cardiac or aortic or carotid sinus terminals and provide a beneficial autonomic state. So, by enhancing the excitability of sensory nerve endings in these pathological states, PGI2 may have a salutary effect (Ustinova & Schultz, 1994). We have shown that a decreased rate of synthesis of PGI2 in the carotid sinus of hypertensive rabbits may contribute to decreased baroreceptor activity (Xie et al. 1990). Restoration of that activity would be beneficial. Several studies have demonstrated that mechanosensitivity of cardiac vagal afferents is suppressed in heart failure, while chemosensitivity of these afferents is enhanced (Brandle et al. 1994; Schultz et al. 1997; Schultz, 2001). These authors also postulated a potentially beneficial compensatory restoration of vagal afferent activity as a result of enhanced chemical sensitivity. The specific conductances that mediate the action of PGI2 as well as other important modulators of neuronal activity need to be defined and the functional relevance of their contribution to sensory signalling requires continued investigation.



This publication was made possible by grant number HL14388 from the National Institutes of Health and a VA Merit Review Award to MWC from the Department of Veterans Affairs. We are grateful to Shawn Roach for graphics assistance and to Shawn Averkamp and Cheryl Ridgeway for secretarial assistance.