Ileitis modulates potassium and sodium currents in guinea pig dorsal root ganglia sensory neurons

Abstract

Intestinal inflammation induces hyperexcitability of dorsal root ganglia sensory neurons, which has been implicated in increased pain sensation. This study examined whether alteration of sodium (Na⁺) and/ or potassium (K⁺) currents underlies this hyperexcitability. Ileitis was induced in guinea pig ileum with trinitrobenzene sulphonic acid (TBNS) and dorsal root ganglion neurons innervating the site of inflammation were identified by Fast Blue or DiI fluorescence labelling. Whole cell recordings were made from acutely dissociated small-sized neurons at 7–10 days. Neurons exhibited transient A-type and sustained outward rectifier K⁺ currents. Compared to control, both A-type and sustained K⁺ current densities were significantly reduced (42 and 34 %, respectively; P < 0.05) in labelled neurons from the inflamed intestine but not in non-labelled neurons. A-type current voltage dependence of inactivation was negatively shifted in labelled inflamed intestine neurons. Neurons also exhibited tetrodotoxin-sensitive and resistant Na⁺ currents. Tetrodotoxin-resistant sodium currents were increased by 37 % in labelled neurons from the inflamed intestine compared to control (P < 0.01), whereas unlabelled neurons were unaffected. The activation and inactivation curves of these currents were unchanged by inflammation. These data suggest ileitis increases excitability of intestinal sensory neurons by modulating multiple ionic channels. The lack of effect in non-labelled neurons suggests signalling originated at the nerve terminal rather than through circulating mediators and, given that Na⁺ currents are enhanced whereas K⁺ currents are suppressed, one or more signalling pathways may be involved.

Abdominal pain is a major cause of morbidity for patients who suffer from intestinal disorders, such as inflammatory bowel disease (IBD) and irritable bowel syndrome. Antecedent infection or inflammation has been implicated in the visceral hyperalgesia associated with the irritable bowel syndrome (IBS). The cell bodies of sensory neurons which initiate these sensations originate in the dorsal root ganglia (DRG) (Mayer & Gebhart, 1994). Sub-populations of DRG neurons respond to innocuous and noxious mechanical stimulation, as well as chemical and thermal stimulation, and thus are thought to be polymodal sensory neurons (Mayer & Gebhart, 1994). These polymodal neurons play a central role in responding to conditions that are potentially injurious to tissues. Inflammatory mediators cause a reduction in the threshold and an increase in the gain of the transduction process of these neurons in a process referred to as ‘peripheral sensitization’. Also, under certain pathological conditions, there is evidence that an additional population of normally mechanically insensitive nociceptive fibres can be recruited (Gebhart, 2000). Following inflammatory stimulation these ‘silent afferent’ fibres become active at rest and begin respond to mechanical stimulation. Together, these afferent pathways have been suggested to contribute to disproportionate pain states in response to injury and may persist even after the inflammatory state has resolved (Al Chaer et al. 2000; Collins et al. 2001).

In studies in the somatic nervous system, peripheral sensitization appears to involve both acute and chronic mechanisms (Woolf & Costigan, 1999). Acute mechanisms of increased nociceptive stimulation include both direct depolarization of nerve terminals by neuroactive agents and alterations in ionic currents in the membrane terminals (Rang et al. 1991). Also, later and longer lasting transcription-dependent changes occur and appear to be evoked by either signalling molecules, such as nerve growth factor and/or activity-dependent second messenger cascades (McCleskey & Gold, 1999). These transcription-dependent events can result in altered expression of voltage-gated ion channels, such as increased expression of TTX-resistant sodium channels, and increased TTX-resistant currents (TTX-R I_Na) (Khasar et al. 1998; Tanaka et al. 1998) or decreased availability of voltage-gated K⁺ channels (Yoshimura & de Groat, 1999).

Compared to the somatic nervous system, there is considerably less known about the ionic mechanisms underlying inflammation-induced changes in sensory neurons innervating the gastrointestinal tract. Recent studies in the rat stomach have shown that experimentally induced gastric ulcers increase TTX-R I_Na in vagal and DRG neurons innervating the stomach (Bielefeldt et al. 2002a) and to a lesser degree in mild gastritis (Bielefeldt et al. 2002b). This latter study, however, found that outward K⁺ currents were unchanged. In contrast, studies in viscera outside the gastrointestinal (GI) tract suggest that inflammation may also modulate K⁺ currents (Yoshimura & de Groat, 1999) Taken together, it is likely that changes in ionic mechanisms underlying inflammation-induced plasticity depend on the organ involved and the inflammatory repertoire which follows the initiation of inflammation.

We have recently used trinitrobenzene sulphonic acid (TNBS) ileitis in the guinea pig, as a model of inflammatory bowel disease (Moore et al. 2002), to examine the effects of intestinal inflammation on DRG neurons. We found hyperexcitability of dissociated DRG neurons innervating the intestine, manifested by decreased threshold and repetitive firing, properties consistent with hyperalgesia and allodynia. This study employs whole cell voltage clamp techniques in this model to determine if changes in Na⁺ and/ or K⁺ currents underlie this hyperexcitability.

METHODS

Animal model and neuron isolation

Guinea pigs (140–225 g) of either sex were obtained from Charles River Laboratories (Montreal, Quebec, Canada). Experiments were performed according to the guidelines of the Canadian Council of Animal Care and approved by the Queen's University animal care committee.

Animals were anaesthetized using a combination of Hypnorm (0.315 mg ml⁻¹ fentanyl citrate and 10 mg ml⁻¹ fluanisone) and midazolam (5 mg ml⁻¹) (0.0025 ml g⁻¹ each, i.p.), and surgery was performed under aseptic conditions to exteriorize the terminal ileum as described previously (Moore et al. 2002). Ileal-projecting neurons were labelled by injecting the retrograde tracer Fast Blue (dissolved in distilled water) or DiI (dissolved in EtOH) into the ileal wall using a Hamilton syringe fitted with a 30 Ga needle. A total injection volume of 15–20 μl was given via 10–15 injection sites. DiI was used in later experiments since Fast Blue became unavailable. Ileitis was induced by injection of 0.5 ml of TNBS (25 mg ml⁻¹ in 25 % EtOH; TNBS kindly provided by Dr G. Morris, Department of Biology, Queen's University, Kingston, Ontario, Canada K7L 5G2) into the ileal lumen using a 30 Ga needle. The intestine was replaced in the abdominal cavity and the wound sutured with 4–0 silk. Buprenex (Buprenophine 0.0225 mg g⁻¹i.p., Reckitt and Colman) was given to all animals to control post-operative pain. Animals recovered from the anaesthetic in a quiet environment, on an electric thermal blanket to maintain normothermia. The post-operative condition of the animals was monitored at least twice daily by trained animal care staff. Animals that were failing to thrive or showed behaviour suggestive of persistent pain were killed. After 7–10 days, animals were anaesthetized by inhalation of isofluorane and killed by cervical dislocation and exsanguination.

The terminal ileum was removed from each animal to establish the degree of TNBS induced ileitis. The ileum was opened along the mesenteric border and pinned flat with the mucosal surface uppermost. Inspection of the mucosa revealed mucosal hemorrhage, ulceration and thickening of the tissue, as described in our previous studies (Moore et al. 2002). In the current study, inspection of the mucosa revealed similar damage following inflammation.

DRG neurons were isolated from thoracic vertebra Th10–13 as described previously (Moore et al. 2002). Briefly, isolated DRG were dissected free of adherent connective tissue and then incubated at 37°C in HBSS with 0.2 mg ml⁻¹ papain and 0.4 mg ml⁻¹ cysteine for 10 min. This was followed by washing in L-15 medium (GIBCO-BRL) with 10 % fetal bovine serum. The DRGs were then incubated in HBSS containing 1 mg ml⁻¹ Type 1 collagenase (Worthington) and 4 mg ml⁻¹ dispase II (Boeringer Manneheim). The ganglia were then titurated with a fire-polished Pasteur pipette. Neurons were maintained in MEM culture medium with Earle's salts and HCO₃ (GIBCO-BRL) containing 1 % penicillin-streptomycin, 2 mm glutamine and 0.2 % (w/v) glucose. The cell suspension was plated onto rat tail collagen-coated glass coverslips and stored in a humidified incubator at 37°C under 95 % air and 5 % CO₂ until they were retrieved for use in electrophysiological experiments 4–24 h later.

Solutions

For current clamp experiments, the control solution used to superfuse the cells contained (mm): NaCl, 140; KCl, 5; MgSO₄, 1; CaCl₂, 1; Hepes, 5; pH adjusted to 7.4 using NaOH. Identical control solutions were used to superfuse the cells in K⁺ current voltage clamp experiments, except for the equimolar replacement of NaCl with N-methyl-d-glucamine. Potassium channel recording solutions were adjusted to pH 7.4 using HCl. The control filling solution contained (mm): KCl, 140; Hepes, 5; MgSO₄, 1; EGTA, 1; pH adjusted to 7.2 using KOH. For the recording of sodium currents, solutions of the following composition were used: extracellular solution (mm): NaCl, 100; NMDG, 50; Hepes, 10; MgCl_2, 10; d-Glucose, 10; pH adjusted to 7.4 with HCl. Pipette solution (mm): Cs, 115; NaCl, 25; Hepes, 10; MgCl_2, 3; EGTA, 11; pH adjusted to 7.2 with CsOH.

Electrophysiological procedures

Coverslips supporting adherent DRG neurons were placed in a RC-26 recording chamber (Warner Instrument Corporation) and mounted on an inverted microscope (IX-70, Olympus, Japan) fitted for both fluorescence and bright field microscopy. Labelled neurons were identified by their fluorescence under brief (< 15 s) exposure to ultraviolet light, using a U-MWIG2 filter for Fast Blue or a U-MWU2 (Olympus, Japan) for DiI, after which cells were viewed under bright field illumination. Whole cell recordings were made using variations of the patch clamp technique (Hamill et al. 1981) and an Axopatch-200B amplifier (Axon Instruments). Patch electrodes were fabricated using thin-walled borosilicate glass (Kimble Products) and a PP-830 electrode puller (Narishige Instruments) or P-97 (Sutter Instruments). After fire polishing, pipettes used for whole cell voltage clamp experiments had DC resistances of 1.5–3.0 MΩ when filled with the control filling solution. Electronic compensation of total series resistance was used in all experiments. Capacitive transients were corrected using analog circuitry. Drugs were added to the bath solution, or delivered by a fast-flow solution switching system (VC-6, Warner Instrument Corporation). Membrane currents were filtered at 2 kHz and sampled at 5 kHz and stored on a microcomputer for later analysis using pCLAMP 8.2 software (Axon Instruments). Cells were included for analysis if in current clamp mode the resting membrane potential was more negative than −40 mV and the cells displayed overshooting action potentials or in voltage clamp mode the series resistance error (after compensation) was less than 5 mV and seal and access resistances remained stable. The mean number of cells studied per animal was 3 (range 1–10).

Drugs and chemicals

All chemicals and drugs were from Sigma Chemical Co. 4-Aminopyridine (4-AP) was dissolved in distilled water to a stock concentration of 100 mm and the pH was adjusted to 7.4 using 2 n HCl. Tetrodotoxin (TTX) was dissolved in water at a stock concentration of 1 mm. Drugs were diluted to their final concentration in the bath solution. Drugs were administered either by direct addition to the bath solution, or directly to the cell under study using a fast-flow solution switching system (Warner Instruments).

Statistical analysis

Student's t test or ANOVA with Student-Newman-Keuls post hoc test were used where appropriate. P values < 0.05 were considered significant.

RESULTS

General properties

Successful recordings were obtained from 149 DRG neurons. Three groups of neurons were examined in this study; Fast Blue- or DiI-labelled neurons from control animals (Control) or TNBS-treated animals (TNBS) and neurons from TNBS-treated animals which were not labelled (non-labelled TNBS). Approximately 3–5 % of the dissociated cells were labelled with Fast Blue or DiI. The mean resting membrane potential in current clamp recordings from labelled control neurons was −58.1 ± 1.1 mV (range −48 to −64 mV; n = 13). Our previous studies of TNBS ileitis using intracellular recording techniques demonstrated that small cell size correlated very closely with the following properties: TTX-resistant action potentials with inflections on the repolarizing phase and capsaicin sensitivity (Moore et al. 2002). These properties have been shown to be present in small-diameter somatic and visceral unmyelinated afferents, a proportion of which are nociceptive afferents (Sengupta & Gebhart, 1994; Blackshaw & Gebhart, 2002). Using whole cell current clamp recordings, we found that small neurons had similar properties, i.e. TTX-resistant action potentials with a prominent inflection of the repolarizing phase (5/5). Measurements of cell capacitance were used to monitor size, and only cells with capacitance < 40 pF were examined, as these cells consistently expressed these properties. Larger cells (mean capacitance 81 ± 7.4 pF) had narrow, TTX-sensitive action potentials (4/4). In our previous work using the same model we demonstrated inflammation-induced changes in excitability of this sub-population of small neurons. In these studies, intracellular recordings were obtained from neurons from control (n = 12) and animals with ileitis (n = 17) and we found that TNBS ileitis caused significant reductions in rheobase, and increased the number of action potentials evoked at 2 × rheobase. Furthermore TNBS ileitis resulted in increased action potential upstroke velocity and increased input resistance. (Moore et al. 2002). In the present study, using whole cell recording techniques, we demonstrated a similar effect of TNBS ileitis on rheobase (> 65 % decrease compared to control and non-labelled cells) and number of action potentials elicited at twice rheobase (> 2.7 times compared to control and non-labelled cells) (Fig. 1).

Figure 1. Effects of TNBS ileitis on intestinal sensory neuron excitability.

A, representative current clamp traces of action potentials elicited at rheobase and two times rheobase in control labelled, TNBS labelled and TNBS non-labelled neurons. TNBS results in a reduction of the rheobase, while increasing the number of spikes evoked by a 2 × rheobase current injection. Inflections on the falling phase of the action potential are not obvious due to the time scale. B, mean rheobase in control labelled (n = 13), TNBS labelled (n = 4) and TNBS non-labelled (n = 5) neurons (*P < 0.05). C, mean number of spikes elicited by a 2 × rheobase current injection in these same groups of neurons (*P < 0.05).

Characterization of voltage-gated K⁺ currents

Previous studies of unidentified DRG neurons have identified two kinetically distinct voltage-dependent K⁺ currents, a transient ‘A-type’ current (I_A) and ‘sustained delayed rectifier type’ (I_K) (McFarlane & Cooper, 1991; Akins & McCleskey, 1993; Gold et al. 1996; Everill et al. 1998). In the present study, using a voltage clamp protocol with 5 mV steps (400 ms) from a holding potential of −100 mV, both the I_A and sustained I_K were evident (Fig. 2A). The reversal potential of these currents was −75.5 ± 2.2 mV, near the predicted reversal potential for potassium (−84 mV). Inactivating I_A currents were adequately fitted by a monoexponential decay with a mean time constant of 171 ± 16 ms (range 133–210 ms).

Figure 2. Voltage-gated potassium currents and the effect of TNBS ileitis on current density.

Representative traces from one cell showing currents separated biophysically by manipulating the holding potential (A) and pharmacologically by 4-AP (B). Currents were elicited in response voltage steps from −90 mV to +50 mV in 5 mV increments. A, left: at holding potentials of −100 mV, two currents were apparent, a transient, inactivating ‘A’ type current, and a non-inactivating sustained I_K type current. A, middle: resultant current when membrane potential is held at −60 mV. Note disappearance of the transient component. Only the sustained component remains, and was measured at 400 ms. A, right: subtraction of the sustained from the total current yields I_A. I_A amplitude was measured as the peak of the transient component. B, left: in the pharmacological experiments, the holding potential was −100 mV. B, middle: when the voltage steps were repeated in the presence of 1 mm 4-AP the transient component was significantly inhibited. B, right: results of subtracting current obtained in the presence of 4-AP from control, revealing the 4-AP-sensitive I_A current. C, effect of TNBS ileitis on mean peak I_A density obtained using either biophysical (left) or pharmacological separation (right). TNBS ileitis results in a significant reduction in I_A density compared to control, or non-labelled TNBS neurons (*P < 0.05). D, effect of TNBS ileitis on mean peak I_K density. TNBS ileitis results in a significant reduction in peak I_K density, compared to control and non-labelled cells (*P < 0.05).

I_A and I_K currents were isolated based on their contrasting biophysical and pharmacological properties. In preliminary experiments, we established that neither current was significantly inactivated when the membrane potential was held at −100 mV (Fig. 2). However, the I_A was selectively inactivated when the membrane potential was held at −60 mV, whereas inactivation of I_K at this holding potential was minimal (Fig. 2). Thus, the sustained current was isolated by holding the membrane potential at −60 mV, with the sustained I_K measured isochronally, 400 ms after the onset of the pulse, at which time I_A was largely inactivated, minimizing contamination by this current (Fig. 2). The I_A was isolated by subtracting the sustained I_K from the total K⁺ current recorded from a holding potential of −100 mV. Peak I_A was measured as the peak of the transient component of this subtracted current (Fig. 2).

I_A was also isolated pharmacologically using the K⁺ channel blocker 4-aminopyridine (4-AP) (Fig. 2B). Preliminary experiments demonstrated that 100 μm 4-AP (n = 6), and 600 μm 4-AP (n = 3) caused partial suppression of the I_A currents whereas 2 mm (n = 3) completely suppressed I_A and caused significant suppression of the I_K currents. 4-AP most selectively blocked the I_A at a concentration of 1 mm. Therefore the I_A was isolated by subtracting the current in the presence of 1 mm 4-AP from the total control current, yielding the 4-AP-sensitive I_A (Fig. 2B). Experiments using both of these approaches in the same control neuron revealed no significant difference in the I_A current density obtained by either method (80.3 ± 19.7 vs. 99.1 ± 10.0 pA pF⁻¹, n = 5)

I_A and I_K current density following TNBS-induced inflammation

The density of I_A and I_K elicited by a depolarizing pulse to +50 mV was compared in labelled neurons from control animals and both labelled and unlabelled neurons from TNBS animals (Fig. 2C and D). Current density was determined by normalizing the current amplitude by the cell's capacitance. The I_K amplitude was measured 400 ms following the onset of a depolarizing pulse from a holding potential of −60 mV. The I_A amplitude was measured as the peak current amplitude of the subtracted current as described above (Fig. 2A and B). Figure 2C-E illustrates the reduction of I_A and I_K in labelled neurons from TNBS animals. When I_A was isolated biophysically, the I_A density was 42 % less (P < 0.05) in labelled neurons from TNBS animals (n = 22) than in labelled neurons from control animals (n = 14) and 65 % less than in unlabelled neurons from TNBS animals (n = 19) (Fig. 2C). I_A was also isolated pharmacologically using 4-AP (1 mm). The 4-AP-sensitive I_A in labelled neurons was reduced by 33 % in labelled TNBS neurons (n = 6, P < 0.05) compared to the currents in labelled neurons from control animals (n = 4) and was 43 % less than in non-labelled TNBS neurons (n = 7, P < 0.01, Fig. 2D). The current density of I_K from labelled TNBS neurons (n = 20) was 34 % less (P < 0.05) than in labelled neurons from control animals (n = 15) and 31 % less (P < 0.05) than in unlabelled neurons from TNBS animals (n = 20) (Fig. 2E).

We also attempted to isolate I_K pharmacologically, using TEA. However, we found, as have others (Everill et al. 1998; Gold et al. 1996) that TEA in doses sufficient to block I_K, also cause significant blockade of the I_A.

Voltage dependence of activation and inactivation

Voltage dependencies of activation and inactivation were compared in labelled control cells and labelled TNBS cells to determine if changes in TNBS animals could contribute to the reduced current density.

To determine I_A K⁺ conductance and I_K K⁺ conductance, peak I_A amplitudes and isochronally measured end-pulse current amplitudes were measured, respectively. Conductance was then determined using the relation G = I/(V_m−E_K), where G is the conductance, I is the measured membrane current, V_m is the voltage step, and E_K is the equilibrium K⁺ potential, which was calculated to be −84 mV in control solutions. Average K⁺ conductance was plotted against membrane potential (Fig. 4) and the continuous line is an average of individual fits to a Boltzmann function of the form:

where G is the conductance, G_max is the fitted maximal conductance, V_1/2 is the membrane potential for half-activation, V_m is the command potential, and k is the slope factor. No differences in the voltage dependencies of activation for I_A or I_K were found between control and TNBS neurons (Fig. 3A and B).

Figure 4. Voltage-gated sodium currents and the effects of TNBS ileitis.

A, representative voltage-gated sodium currents from a small (25 pF) intestinal DRG neuron. Left: total inward current generated by stepwise 30 ms voltage pulses between −80 and +40 mV from a holding potential of −90 mV. Middle: TTX-R I_Na currents obtained using the same protocol in the presence of 1 μm TTX. Right: TTX-S I_Na currents obtained by subtracting TTX-R current from total. B, mean peak TTX-R I_Na current density. TTX-R I_Na density was increased by TNBS ileitis, *P < 0.05 compared to control and non-labelled neurons (n = 7 cells for each group). C, TTX-S I_Na current density was not significantly affected by TNBS ileitis (P > 0.05, n = 7 cells for each group).

Figure 3. Activation and inactivation curves of voltage-gated K⁺ currents.

Activation curves were generated by voltage pulses in 5 mV steps from −80 to +50 mV. Each curve represents the mean of curves fitted to the Boltzmann equation (n = 8 cells each). A, activation curves for I_A in control and TNBS neurons. TNBS did not affect the activation properties of I_A. B, inactivation curves for I_A. Inactivation curves were constructed using a two-pulse protocol, a 1 s prepulse varying between −120 and 0 mV, followed by a 400 ms test pulse of +50 mV. The peak of the transient component was measured. TNBS treatment resulted in a slight leftward shift in the inactivation curve of I_A. Activation (C) and inactivation (D) curves for I_K. Inactivation curves for I_K were generated using a two-pulse protocol, with an 8 s prepulse varying between −80 and 0 mV, followed by a 1 s test pulse of +50 mV. End-pulse I_K amplitude was measured. TNBS had no effect on voltage dependence of either activation or inactivation of I_K.

To examine the voltage dependencies of inactivation for I_A and I_K, we employed two-pulse voltage protocols as described by others (Philipson et al. 1991; Yoshimura & de Groat, 1999). Residual I_A currents were measured following short conditioning pulses (1 s duration) which allowed inactivation of only the rapidly inactivating I_A currents. Longer conditioning pulses (8 s duration) allowed inactivation of both I_A and I_K currents. Therefore residual currents were isochronally measured at the end of a 1 s test pulse to minimize the contribution of I_A currents to the measured residual current. The residual current amplitude was plotted against conditioning pulse potential and the continuous line is an average of fits to a negative Boltzmann function:

where I is the current, I_max is the maximal current, V_1/2 is the membrane potential for half-activation, V_m is the command potential, and k is the slope factor (Philipson et al. 1991). Significant differences in I_A were found between control and TNBS neurons with V_1/2 (control −65.4 ± 2.0 mV, TNBS 85.2 ± 2.4 mV, P < 0.05) and slope factors (control 6.9 ± 0.9, TNBS 14.3 ± 1.1, P < 0.05) (Fig. 3C). However, the voltage dependence and voltage sensitivity of inactivation for I_K were not different between control and TNBS neurons (V_1/2: control −47.6 ± 2.8 mV, TNBS −50.2 ± 3.0 mV; slope: control 7.5 ± 1.5, TNBS 9.4 ± 1.2, P > 0.05; Fig. 3D).

We also employed a pharmacological approach using 4-AP (1 mm) to isolate I_A and examine its voltage dependence. Using the above two-pulse protocol, in the absence and presence of 4-AP, the currents obtained in the presence of 4-AP were subtracted from those obtained in its absence (n = 10). These experiments indicated that sensitivity to 4-AP varied significantly with the prepulse voltage thus making analysis of the voltage dependence of inactivation of the subtracted 4-AP sensitive current impractical. This finding has been reported by others (Thompson, 1982; Rasmusson et al. 1995).

Characterization of voltage-gated sodium currents

Large inward currents were evoked in all neurons (n = 60) by depolarizing +5 mV voltage steps from −80 to +40 mV from a holding potential of −90 mV (Fig. 4A). These currents were separated into a TTX-resistant (TTX-R I_Na) and TTX-sensitive (TTX-S I_Na) components using TTX (1 μm). TTX-S I_Na was obtained by subtracting the inward currents in the presence of 1 μm TTX from the currents recorded in the absence of TTX (Fig. 4A). All cells examined exhibited both the TTX-R I_Na and TTX-S I_Na currents.

Effects of TNBS ileitis on Na⁺ current density

TTX-R I_Na and TTX-S I_Na peak current densities were obtained by normalizing the current amplitude by the individual cell's capacitance. TTX-R I_Na was increased significantly in the labelled TNBS group (n = 10) compared to labelled control neurons (n = 10) (37 %, P < 0.01) or non-labelled control neurons (n = 10) (32 %, P < 0.05) (Fig. 4B). TTX-S I_Na was not significantly different between inflamed labelled neurons (282.0 ± 12.7 pA pF⁻¹, n = 10), labelled control neurons (312.2 ± 22.4 pA pF⁻¹, n = 10, P > 0.05) or TNBS non-labelled neurons (282.3 ± 17.0 pA pF⁻¹, n = 10, P > 0.05) (Fig. 4C).

Activation and inactivation properties of TTX-R I_Na

We examined the voltage dependence of activation and inactivation of TTX-R I_Na to determine if changes in these properties could contribute to the increase in current density induced by inflammation. Activation curves were generated using depolarizing voltage steps in the presence of TTX (1 μm). Normalized conductance (G/G_max) was plotted against test pulse voltage and the data were fitted using a Boltzmann function of the form:

Voltage of half-activation and slope were calculated from the average of the fitted activation curves. Inflammation did not alter the slope (control 4.0 ± 0.50, TNBS 5.2 ± 0.32, n = 7, P > 0.05) or the voltage of half-activation of TTX-R I_Na (control −12.8 ± 2.0, TNBS −16.9 ± 0.38, n = 7, P > 0.05) (Fig. 5A).

Figure 5. Activation and inactivation curves for TTX-R I_Na.

Activation curves in A were generated using voltage steps from −80 to +20 mV. Normalized conductance (G/G_max) was plotted against test pulse voltage and fitted to a Boltzmann function. Lines represent average of the individual curve fits. TNBS ileitis did not affect the activation properties of the TTX-R I_Na. (P > 0.05, n = 7) B, inactivation curves for TTX-R I_Na. Inactivation curves were generated using a two-pulse protocol, with a 1 s prepulse between −120 and 0 mV, followed by a 30 ms test pulse to 0 mV. Normalized current (I/I_max) was plotted against prepulse voltage and fitted to a negative Boltzmann function. Line represents the average of individual curve fits. Inactivation properties were not significantly affected by TNBS ileitis (n = 7, P > 0.05).

Steady-state inactivation curves were generated using a 1 s prepulse from −120 to 0 mV, followed by a 30 ms test pulse to 0 mV. The resultant normalized peak current (I/I_max) was plotted against the prepulse voltage, and the data were fitted to a negative Boltzmann function of the form:

Mean voltage of half-inactivation and slope were calculated from the average of the individual fitted curves (Fig. 5B). Inflammation had no effect on the voltage of half-inactivation of the TTX-R I_Na (control 26.0 ±1.6 mV, TNBS 25.6 ± 2.3 mV, n = 7, P > 0.05), or the slope (Control −2.03 ± 0.45 mV⁻¹, TNBS −1.7 ± 0.42 mV⁻¹).

DISCUSSION

Voltage-gated Na⁺ and K⁺ channels play a fundamental role in controlling neuronal excitability (Hille, 1992) and exhibit significant diversity in sensory neurons, depending upon their innervation and functional properties. The present study characterized K⁺ and Na⁺ currents in small-diameter sensory neurons and tested the hypothesis that alteration of one or more currents underlies the hyperexcitability seen in these neurons during inflammation. DRG neurons were functionally identified by retrograde labelling from the intestine, and only small-sized cells were selected for study, as we had previously shown that small cell size correlates with properties such as TTX-resistant action potentials, capsaicin sensitivity, and inflections on the repolarizing phase of the action potential (Moore et al. 2002). Sensory neurons innervating the intestine exhibit transient I_A and sustained outward I_K K⁺ currents and both TTX-resistant and TTX-sensitive inward Na⁺ currents. The major finding was that TNBS ileitis, a model of inflammatory bowel disease caused a marked reduction in the voltage-gated potassium currents, I_A and I_K, whereas TTX-R I_Na currents were significantly increased. These changes were not observed in unlabelled neurons from animals with ileitis, suggesting that signalling occurred at the nerve terminal rather than as result of circulating inflammatory mediators. Moreover, the opposing effects on K⁺ and Na⁺ currents would suggest that multiple signalling pathways are involved.

Voltage-gated K⁺ currents in neurons innervating control and inflamed ileum

The I_A current plays an important role in setting high thresholds for spike activation and suppression of repetitive firing (Rudy, 1988; Hille, 1992; Yoshimura & de Groat, 1999). Several types of I_A current have been described in DRG sensory neurons (Gold et al. 1996; Yoshimura & de Groat, 1999) and can be separated into those with relatively fast and slow inactivation rates based on their biophysical and pharmacological properties (Gold et al. 1996; Yoshimura & de Groat, 1999). Several studies have shown that the fast inactivating I_A currents are confined to the large-sized neurons with TTX-sensitive spikes (Yoshimura et al. 1996; Gold et al. 1996). The slowly inactivating I_A current has an inactivation time constant of between 150 and 300 ms and the voltage of half-maximal inactivation is displaced more positively than the fast inactivating I_A (Akins & McCleskey, 1993; Fjell et al. 1999). In the present study the transient I_A inactivation was fitted by a single monoexponential function with a time constant of 171 ms, typical of that described for the slowly inactivating I_A. These slowly inactivating I_A currents, in contrast to the fast inactivating currents, have been found to be selectively expressed in the small-sized neurons exhibiting TTX-resistant action potentials (Yoshimura et al. 1996). Taken together, the data suggest that slowly inactivating I_A current is the dominant form of I_A in sensory afferent DRG neurons innervating the small intestine.

In addition to the transient I_A current, studies of DRG sensory neurons have identified a sustained delayed rectifier current in all cells (Akins & McCleskey, 1993; Gold et al. 1996). In the present study, these currents could be separated from the I_A currents by prepulse inactivation or by 1 mm 4-AP which blocked the I_A currents but had little effect on the sustained current, as reported by others (Gold et al. 1996). Some studies have suggested that this sustained current may represent several different currents, based largely on their steady-state inactivation properties (Akins & McCleskey, 1993; Gold et al. 1996). These sustained currents, however, often could not be separated by voltage protocols or pharmacological agents (Gold et al. 1996) making the study of changes in individual sustained currents impractical.

TNBS ileitis caused a significant reduction in both I_A and sustained I_K currents identified in this study and we have previously shown that pharmacological suppression of K⁺ currents such as I_A can significantly increase the excitability of sensory neurons innervating the intestine (Moore et al. 2002). It is unclear whether these changes in K⁺ currents occur elsewhere in the GI tract because there has been relatively little work done in this area and in studies conducted in a gastritis model of inflammation similar changes were not observed (Bielefeldt et al. 2002b). It is possible that this difference can be explained by variations in the degree of inflammation or experimental protocols. Our finding that visceral inflammation suppresses I_A and correspondingly increases excitability of sensory neurons is similar to previous studies examining the inflamed urinary bladder (Yoshimura & de Groat, 1999). This reduction in peak current density was associated with a hyperpolarizing shift in the inactivation curve, as described in the current study, suggesting there was an associated change in the biophysical properties of the channels. The shift of the inactivation curve in a hyperpolarizing direction would make fewer I_A channels available, at or near resting membrane potentials, and lead to a further increase in excitability of the cell, leading to repetitive firing. In contrast to the study in the rat urinary bladder, we also observed a marked reduction in the current density of I_K. Whether these differences also reflect degrees of inflammation in these organs or are unique to the sensory neurons and the organs they innervate remains to be established. Given that the role of these voltage-gated potassium currents is to raise the action potential threshold and limit firing (Hille, 1992) suppression of these currents would be expected to increase excitability.

Voltage-gated Na⁺ currents in neurons innervating control and inflamed ileum

Three types of sodium channels predominate in DRG neurons; Na_V1.7 channels, which are responsible for the fast, rapidly inactivating TTX-S I_Na current, Na_V1.8 channels, which result in a more slowly inactivating TTX-R I_Na and are relatively selectively expressed in nociceptors, and the recently described Na_V1.9 which is responsible for a TTX-R persistent current that is ultra-slowly inactivating (Dib-Hajj et al. 2002).The activation and inactivation properties of our TTX-R I_Na currents fit the known properties of the Na_V1.8 channel, although a detailed molecular or immunocytochemical characterization of guinea pig DRG sodium channels has yet to be performed.

We observed a significant increase in TTX-R I_Na in sensory neurons innervating the inflamed ileum, as has been implicated in other viscera (Yoshimura et al. 2001; Bielefeldt et al. 2002a,b) and the somatic nervous system. In the somatic nervous system (Khasar et al. 1998; Tanaka et al. 1998) these changes were also accompanied by neuronal hyperexcitability and molecular evidence of increased expression of sodium channels (Gould et al. 1998,1999), specifically the TTX-R channel subtype, Na_V1.8 (Tanaka et al. 1998). Moreover, mice deficient in Na_v1.8 exhibit decreased neuronal excitability (Renganathan et al. 2001) and decreased visceral pain (Laird et al. 2002). Antisense Na_V1.8 oligodeoxynucleotide can also prevent inflammation-induced mechanical hyperalgesia (Khasar et al. 1998) and cyclophosphamide cystitis-induced bladder hyperreflexia (Yoshimura et al. 2001). Taken together, these studies suggest that inflammation-induced increased expression of TTX-R I_Na in nociceptive neurons is common to inflammation in both the somatic and visceral organs, including the GI tract, and is an important mechanism underlying neuronal hyperexcitability and visceral hyperalgesia.

Potential mechanisms mediating inflammation-induced changes in ion channels

The inflammatory milieu surrounding the nerve terminal contains numerous inflammatory mediators, such as PGE₂, and 5-HT, which have been shown to augment TTX-R Na⁺ currents in vitro, in both visceral (Gold et al. 2002) and somatic afferents (Cardenas et al. 2001; Gold et al. 2002). PGE₂ has also been shown to decrease I_K potassium currents through the same cyclic AMP-protein kinase C-dependent mechanism which alters Na⁺ currents, in unidentified DRG neurons (Nicol et al. 1997; Evans et al. 1999). These changes occur within minutes and are dependent on the activation of intracellular second messenger systems, and probably subsequent phosphorylation of ion channels or other regulatory cell signalling proteins. These actions cannot account for the findings in the present study because recordings were obtained from cell bodies of DRG neurons located in ganglia outside the inflammatory milieu of the intestine and recordings were made at least 6–8 h after removal of the neurons from the animal at a time when unlabelled neurons were unaffected. Consequently, changes in ionic currents appear to result from longer term alterations such as transcriptional events altering the number of channels, their subunits or other biophysical properties of the membrane channels themselves.

Both activity-dependent and growth factor signalling pathways have been proposed to activate transcription factors which can alter ion channel expression. In particular there is considerable evidence that the neurotrophin nerve growth factor (NGF) can increase expression of TTX-R I_Na (Fjell et al. 1999; Bielefeldt et al. 2003). In addition NGF and other neuotrophins have been shown to modulate various potassium channels, via activation of tyrosine kinases (Yang et al. 2001) or via activation of ceramide (Zhang et al. 2002). Thus NGF appears to be an important candidate for modulating ion channel activity in visceral inflammation. However, experiments specifically designed to test this hypothesis are needed. It is also unknown whether the diverging effects on these currents are due to signalling through a single pathway or whether multiple pathways are involved.

Conclusions

This study demonstrates that an animal model of inflammatory bowel disease is associated with a suppression of transient and sustained K⁺ currents and augmentation of TTX-R I_Na currents in intestinal sensory neurons. The implications are that these mechanisms are important in the increased neuronal excitability seen in these neurons as well as in the increased nociceptive trafficking which occurs during intestinal inflammation, such as in Crohn's disease. Furthermore, there is increasing evidence that visceral hyperalgesia underlies many functional bowel disorders, such as post-infectious irritable bowel syndrome. Whether the findings observed in the current study are present following the resolution of macroscopic inflammation, or whether low levels of microscopic inflammation, which are suggested to persist in these conditions (Collins et al. 2001; Chadwick et al. 2002; Tornblom et al. 2002), are sufficient to sustain these changes remains to be established.