Gastric ulcers reduce A-type potassium currents in rat gastric sensory ganglion neurons

K. Dang,1 K. Bielefeldt,2 and G. F. Gebhart1

Departments of 1Pharmacology and 2Internal Medicine, The University of Iowa, Iowa City, Iowa 52242

Submitted 12 June 2003 ; accepted in final form 1 October 2003


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Voltage-dependent potassium currents are important contributors to neuron excitability and thus also to hypersensitivity after tissue insult. We hypothesized that gastric ulcers would alter K+ current properties in primary sensory neurons. The rat stomach was surgically exposed, and a retrograde tracer (1,1'-dioctadecyl-3,3,3,3'-tetramethylindocarbocyanine methanesulfonate) was injected into multiple sites in the stomach wall. Inflammation and ulcers were produced by 10 injections of 20% acetic acid (HAc) in the gastric wall. Saline (Sal) injections served as control. Nodose or T9–10 dorsal root ganglia (DRG) cells were harvested and cultured 7 days later to record whole cell K+ currents. Gastric sensory neurons expressed transient and sustained outward currents. Gastric inflammation significantly decreased the A-type K+ current density in DRG and nodose neurons (Sal vs. HAc-DRG: 82.9 ± 7.9 vs. 46.5 ± 6.1 pA/pF; nodose: 149.2 ± 10.9 vs. 71.4 ± 11.8 pA/pF), whereas the sustained current was not altered. In addition, there was a significant shift in the steady-state inactivation to more hyperpolarized potentials in nodose neurons (Sal vs. HAc: -76.3 ± 1.0 vs. -83.6 ± 2.2 mV) associated with an acceleration of inactivation kinetics. These data suggest that a reduction in K+ currents contributes, in part, to increased neuron excitability that may lead to development of dyspeptic symptoms.

stomach; visceral inflammation; hyperalgesia; dorsal root ganglia; nodose ganglia; voltage clamp; 4-aminopyridine; dendrotoxin; K+ currents


PAIN AND DISCOMFORT ARE PROMINENT symptoms of many chronic diseases of the gastrointestinal tract and are considered to reflect visceral hypersensitivity. Changes in the excitability of primary afferent neurons (peripheral sensitization) and/or altered information processing in the spinal cord or higher centers (central sensitization) are principal reasons for development of such hypersensitivity. We recently developed animal models to study the relative role of peripheral sensitization in gastric hypersensitivity (32). Because voltage-sensitive ion currents are critical for action potential generation and neuron excitability (26), we examined in these models properties of voltage-sensitive sodium currents in dissociated primary gastric sensory neurons harvested from the nodose and dorsal root ganglion (DRG) (1). We found significant changes in peak sodium current, contributed almost exclusively by the TTX-resistant current, and in activation and inactivation kinetics after gastric insult, consistent with an important role of peripheral sensitization in visceral hypersensitivity triggered by gastric inflammation.

Other currents contribute to the control of neuron excitability at rest and during stimulation. Voltage-gated K+ channels are responsible for repolarization and regulate action potential firing patterns (20). Neurons express multiple voltage-sensitive K+ currents that can be differentiated based on their biophysical and pharmacological properties (11, 17, 43). In the present study, we characterized the properties of voltage-sensitive K+ currents in vagal and spinal gastric sensory neurons and tested the hypothesis that gastric inflammation alters K+ currents, thereby contributing to sensitization of gastric afferent neurons and gastric hypersensitivity. A preliminary report of some of these data has appeared in abstract form (6).


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Male Sprague-Dawley rats (150–200 g; Harlan, Indianapolis, IN) were used for all experiments. Rats were housed under a 12:12-h light-dark cycle with free access to food and water and were handled following the guidelines of the American Physiological Society. The experimental protocol was approved by the Animal Care and Use Committee of The University of Iowa.

Cell labeling and induction of gastric ulcers. Under intraperitoneal pentobarbital sodium anesthesia (50 mg/kg), the rat stomach was surgically exposed and 1,1'-dioctadecyl-3,3,3,3'-tetramethylindocarbocyanine methanesulfonate (DiI; 100 mg in 2 ml DMSO; Molecular Probes, Eugene, OR) was injected into 10 sites in the stomach wall (4 µl/site). The wound was closed with 4.0 silk suture, and rats were allowed to recover for 7 days. Rats were again anesthetized with pentobarbital sodium (as above), the stomach was reexposed, and 20% acetic acid (HAc) or saline (Sal; control) was injected into 10 sites in the rat stomach wall in a similar manner (4 µl/site). The previously labeled areas could be easily identified due to the presence of DiI, allowing us to selectively inject HAc or Sal into areas with labeled afferents (1). Nodose and DRG neurons were harvested 7 days later.

Cell dissociation and culture. Rats were anesthetized and decapitated, and the nodose ganglia or bilateral T9-T10 DRG was rapidly removed. The ganglia were minced and incubated at 37°C, 5% CO2 for 45 min in low glucose DMEM (GIBCO, Invitrogen, Grand Island, NY) containing collagenase (type 4; 2 mg/ml; Worthington Biochemical, Lakewood, NJ), trypsin (1 mg/ml), and DNAse (0.1 mg/ml; both from Worthington). Tissue fragments were gently triturated to encourage cell dissociation. The enzymatic digestion was terminated by adding soybean trypsin inhibitor (2 mg/ml; Sigma-Aldrich, St. Louis, MO), bovine serum albumin (2 mg/ml; Amresco, Solon, OH), and 5% rat serum (Atlanta Biologicals, Norcross, GA). Cells were collected by 5-min centrifugation at 150 g and resuspended in DMEM containing 5% rat serum and 2% chick embryo extract (GIBCO). The cells were plated on poly-D-lysine-coated coverslips and incubated at 37°C, 5% CO2 for 2–3 h before electrophysiological studies. Acutely dissociated neurons were round and devoid of any processes, thus reducing space-clamp errors.

Solutions and electrophysiological recordings. Cells were rinsed and transferred to a recording chamber (1 ml) filled with external solution (in mM): 150 N-methyl-D-glucamine (NMDG), 5 KCl, 4 MgCl2, 0.03 CdCl2, 10 HEPES, 10 glucose, and 0.1 TTX. pH was adjusted to 7.4 with HCl with an osmolarity of 295 mosM. Neurons that innervated the stomach were identified by DiI content using a rhodamine filter (excitation wavelength ~546 nm, barrier filter at 580 nm). Consistent with our previous studies, only one to five cells per coverslip (<2% of the total cells) were unambiguously labeled (1). Fire-polished micropipettes with tip resistances of 2–3 M{Omega} were used for voltage-clamp recordings. The pipette was filled with internal solution (in mM): 100 NMDG, 40 KCl, 1 CaCl2, 2 MgCl2, 10 EGTA, 4 Na2ATP, and 0.5 Tris-GTP. pH was adjusted to 7.2 using HCl with osmolarity of 295 mosM. After gigaseal formation, the membrane patch was ruptured by slight suction. The voltage was clamped at -70 mV by an Axopatch 200B amplifier (Axon Instruments, Union City, CA), digitized at 10 kHz (Digidata 1350, Axon Instruments), and controlled by Clampex software (Axon Instruments). Series resistance was compensated by >60%. Considering the peak transient outward current of <4 nA, the maximal voltage error after electronic correction would be <9 mV, assuming a doubling of the access resistance after establishing the whole cell configuration. Cells that had <60% series resistance compensation were discarded from the study. The leak current and residual capacitative transients were digitally subtracted using a P/n protocol, with P being the test pulse and n being the number of hyperpolarizing voltage steps used for digital subtraction (n = 6 unless stated otherwise). To isolate transient outward currents, we employed a previously described electrophysiological protocol (11, 27). Cells were held for 500 ms at -110 mV before stepping them to various test potentials between -30 and 50 mV. To inactivate the transient outward current, the sequence was repeated from a prepotential of -30 mV. The digital subtraction of current these traces from those triggered from a holding potential of -110 mV isolated the transient component of the outward current. To ensure that slow inactivation of the delayed rectifier did not significantly confound the results, we inhibited the transient outward current and fitted the decay of the sustained current with a single exponential. The time constant of 2.7 ± 0.1 s at a test potential of 50 mV (n = 3) is consistent with prior data on the C-type inactivation and argues against a distortion of our results due to slow inactivation of the delayed rectifier. All experiments were performed at room temperature (21–23°C).

Drugs were dissolved in water and added to the recording chamber and allowed to equilibrate for 5 min before recordings began. Unless stated otherwise, all chemicals were purchased from Sigma-Aldrich.

Statistical analysis. All data are given as means ± SE. Comparisons were made using Student's unpaired t-test. Results were considered to be statistically significant when P < 0.05.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Electrophysiological characterization of K+ currents. Consistent with prior reports, depolarization from -110 mV to various test potentials triggered rapidly activating and partially inactivating outward currents in both nodose and DRG neurons (Fig. 1). Changes in the extracellular potassium concentration shifted the reversal potential as predicted by the Nernst equation, confirming that potassium was the primary charge carrier (reversal potential at [K+]e = 5 mM: -48 mV; reversal potential at [K+]e = 20 mM: -17 mV).



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 1. Representative outward currents obtained in gastric sensory neurons evoked in dorsal root ganglia (DRG; A) and nodose ganglia (NG) neurons (B). DRG and nodose neurons had a cell capacitance of ~75 and 50 pF, respectively. Outward currents elicited by a 300-ms depolarization to +50 mV from a holding potential (Vh) of -110 or -30 mV are superimposed. The A current (IA; asterisk) was isolated by digitally subtracting the current obtained after depolarization from -30 mV from the current triggered after depolarization from -110 mV. Gastric ulceration decreased the A current in DRG (C) and NG (D) neurons. HAc, acetic acid.

 

To further characterize the transient current, we used the electrophysiological approach to separate inactivating from sustained currents described above (see MATERIALS AND METHODS). Gastric nodose neurons expressed inactivating outward currents that could be differentiated based on their voltage dependence and kinetics. A rapidly activating and inactivating current was present in 14 of 20 cells examined. The current reached its peak amplitude within 3.9 ± 0.4 ms and decayed with a time constant of 5.6 ± 0.8 ms after a depolarization to +50 mV. As shown in Fig. 2, A and B, the time course of activation and inactivation was voltage dependent. A second inactivating component became obvious at more depolarized potentials in the same 14 of these 20 cells. At steps to +50 mV, the current reached its peak within 28 ± 7.4 ms and inactivated with a time constant of 136 ± 8 ms. In contrast to the rapidly decaying current, the time course of inactivation did not depend on voltage within the voltage range tested (Fig. 2). Similarly, we identified rapidly and slowly activating and inactivating outward currents in 14 and 19 of 30 DRG neurons, respectively. The time constants at 50 mV were 6.1 ± 1 and 125.1 ± 10.9 ms, respectively (Fig. 2, C and D).



View larger version (26K):
[in this window]
[in a new window]
 
Fig. 2. Activation and inactivation kinetics of outward currents in control NG (A and C) and DRG (B and D) neurons. The fast- and slow-inactivating IA components were obtained by digital subtraction of test pulses to 50 mV (see text for details). A: the fits obtained over a range of voltages and plotted as a function of the test potential for the rapidly (filled symbols; n = 14) and slowly decaying component (open symbols; n = 14), demonstrating strong voltage-dependent inactivation kinetics for the rapidly inactivating component in NG neurons. The time to peak is plotted as a function of the test potentials in B for the fast (filled symbols)- and slow (open symbols)-inactivating currents, showing voltage-dependent activation kinetics for the fast-activating component. Similarly, C shows strong voltage-dependent inactivation kinetics for the rapidly (filled symbols; n = 14), but not for the slowly decaying component (open symbols; n = 19) in DRG neurons. The time to peak (D) for the fast (filled symbols)- and slow (open symbols)-inactivating currents shows voltage-dependent activation kinetics for the fast-activating component.

 

To further characterize the transient outward currents, we determined the voltage dependence of inactivation by changing the voltage from prepotentials between -110 and 0 mV for 500 ms to a test potential of +10 mV. All results were normalized to the peak current at -110 mV. As shown in Fig. 3, the inactivating current in both nodose and DRG neurons could be dissected into two components based on its voltage dependence of inactivation. Approximately 35% of the peak current inactivated at hyperpolarized potentials, with an additional 15–20% inactivating at steps to -20 mV or more depolarized potentials. We fitted the decrease in peak currents at voltages negative to -40 mV to a Boltzmann equation. The voltage of half-inactivation was -78.9 ± 1.2 mV (slope factor: -7.7 ± 0.5) for DRG neurons (n = 11) and -76.3 ± 1.0 mV (slope factor: -9.6 ± 0.6) for nodose neurons (n = 12; not significant compared with DRG neurons). The second inactivating component could not be properly fit to a Boltzmann equation due to its relatively small amplitude.



View larger version (12K):
[in this window]
[in a new window]
 
Fig. 3. Steady-state inactivation kinetics in control gastric sensory neurons. Current (I) availability was determined by stepping the neurons to a test potential of +10 mV from 500-ms prepotentials between -110 and 0 mV. Current was normalized relative to that at -110 mV. Gastric ulceration (filled symbols) shifted the steady-state inactivation kinetics to more hyperpolarized potentials compared with control (open symbols) DRG (A) and NG (B) neurons.

 

Effects of 4-aminopyridine and {alpha}-dendrotoxin. To determine the pharmacological properties of outward currents, we used 4-aminopyridine (4-AP) and {alpha}-dendrotoxin (DTX). 4-AP inhibited the fast and slow transient outward currents in both nodose and DRG neurons (Fig. 4). In the presence of 5 mM 4-AP, the peak inactivating component decreased by 65 ± 7% in nodose neurons (n = 6) and 50 ± 6% in DRG neurons (n = 5). In cells with slowly inactivating outward currents, 4-AP significantly decreased the current at the end of the depolarizing pulse, which may be at least partly due to the slow inactivation kinetics. However, as shown in Fig. 4A, 4-AP did not only block the transient outward current but also affected the sustained component, albeit to a lesser degree. These results are consistent with prior reports, demonstrating that 4-AP can inhibit the delayed rectifier in some cells (24, 35).



View larger version (16K):
[in this window]
[in a new window]
 
Fig. 4. Effects of 4-aminopyridine (4-AP) and {alpha}-dendrotoxin (DTX) on the outward currents. Outward currents were evoked by depolarization to test potential of +50 mV from a holding potential of -110 mV (500 ms) in the presence and 5 min after the application of the 4-AP (A and B) and DTX (C and D), respectively. To demonstrate the effect of these agents on outward currents, the trace obtained after drug administration was digitally subtracted from the control trace (*).

 

Application of DTX (1 µM, 5 min) reduced the peak and the sustained currents in both groups of neurons studied (Fig. 5). DTX inhibited 35 ± 8 and 20 ± 4% of the A current in nodose (n = 4) and DRG (n = 5) neurons, respectively. Similarly, the sustained current at the end of a 300-ms step to 50 mV was decreased by 25 ± 7 and 11 ± 4% in nodose and DRG neurons, respectively.



View larger version (28K):
[in this window]
[in a new window]
 
Fig. 5. Effects of gastric ulcers on outward current density in sensory neurons. Total outward current was reduced in DRG (A) and NG (B) of rats with gastric ulcers compared with controls. Digital isolation of the IA showed ~40 and 60% reductions of IA in DRG and NG neurons of HAc-treated rats compared with controls. C and D: normalized conductance of the IA for DRG and NG neurons, respectively. In NG neurons, the voltage dependence of activation was shifted significantly to more positive potentials following gastric ulceration (filled symbols) compared with saline-treated controls (open symbols) but remained unchanged in both groups of DRG neurons.

 

Effects of gastric ulceration on K+ currents. HAc-induced gastric ulcers were confirmed macroscopically by the presence of ulcerations or wall thickening due to inflammation, consistent with previous report (2). Gastric ulceration significantly increased nodose (n = 18) cell capacitance to 57.1 ± 3.1 pF compared with 43.1 ± 2.8 pF from Sal-treated controls (P < 0.05). In contrast, DRG (n = 18) cell capacitance remained unchanged (62.3 ± 2.5 vs. 68.1 ± 3.5 pF; not significant). To determine the effect of gastric injury on K+ currents, we measured the outward currents triggered by a depolarization to 50 mV. To control for changes in cell size, we divided the results by cell capacitance (current density). As shown in Fig. 5, gastric ulceration caused a significant decrease in density of the peak current from 276 ± 11 to 205 ± 11 pA/pF (n = 18; P < 0.001, Sal vs. HAc) in nodose neurons and from 266 ± 9 pA/pF to 209 ± 11 pA/pF in DRG neurons (n = 24; P < 0.01, Sal vs. HAc). In contrast, the density of the sustained outward current did not change significantly (nodose neurons: 154 ± 10 vs. 144 ± 13 pA/pF; DRG neurons: 192 ± 12 vs. 215 ± 14 pA/pF; Sal vs. HAc, respectively). Consistent with these results, digital subtraction revealed that there was a significant decrease in the peak A current from 149 ± 11 to 71 ± 12 pA/pF (n = 18; P < 0.001) in nodose neurons and from 83 ± 8 to 47 ± 6 pA/pF in DRG neurons (n = 24; P < 0.01).

To examine whether gastric ulceration altered properties of the transient outward current, we determined kinetics and the voltage dependence of activation and inactivation as described above. Because we noted a decrease in the peak current, we analyzed the voltage dependence of activation of the electrophysiologically isolated A current. We expressed the results as normalized conductance determined by dividing the current (test voltages used in the experiments ranging from -30 to +50 mV) by the electrochemical driving force potassium reversal potential at 22°C (Vtest - Ek) and fitted the results with a Boltzmann equation. In nodose neurons, the voltage of half-activation was significantly shifted from -26.6 ± 1.8 to -14.3 ± 4.0 mV (P < 0.05; Fig. 5). The significant decrease in transient outward current only allowed a detailed analysis in some cells. In contrast to the results obtained in nodose neurons, we did not see a significant change in the voltage dependence of activation after induction of gastric ulcerations (Fig. 5; -4.5 ± 3.5 vs. -12.7 ± 2.7 mV; Sal vs. HAc; not significant). Thirteen nodose neurons exhibited sufficient current density to allow determination of kinetics; eight primarily expressed rapidly inactivating currents, and five predominantly expressed slowly inactivating currents. Whereas the fast inactivation was not changed, the time constants of the more slowly decaying component were significantly shorter in cells from animals with gastric ulcers (Fig. 6). Of 13 DRG neurons examined, a rapidly inactivating current sufficient in amplitude to allow determination of current kinetics was present in only one cell. Two neurons expressed slowly decaying currents with time-to-peak and inactivation time constants within the range recorded under control conditions.



View larger version (15K):
[in this window]
[in a new window]
 
Fig. 6. Effects of gastric ulcers on inactivation kinetics in nodose neurons. The fast (A)- and slow-inactivating (B) IA components were obtained by digital subtraction of test pulses to 50 mV (see text for details) and fitted with a single exponential. The fast-inactivating component of the IA did not differ between the 2 groups (A). The slowly inactivating component (B) was significantly faster in HAc-treated rats (filled symbol) compared with controls (open symbols).

 

Because the reduction in A current density may be partly due to a change in the voltage dependence of inactivation, we examined this property in sensory neurons from HAc-treated animals following the protocol described above. As shown in Fig. 3, the two distinct components could still be separated with an apparent shift of the steady-state inactivation to more hyperpolarized potentials in nodose neurons. By fitting values obtained with prepulse potentials negative of -40 mV with the Boltzmann equation (see above), we obtained voltages for half-inactivation of -83.6 ± 2.2 mV (slope factor: 9.0 ± 0.6; n = 9; P < 0.05 compared with Sal) for nodose neurons and -79.7 ± 1.2 mV (slope factor 9.2 ± 0.5; n = 13; not significant compared with Sal) for DRG neurons.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
With the use of a previously established animal model of gastric ulceration, we have recently shown behavioral changes consistent with the development of gastric hypersensitivity (32). To better understand peripheral mechanisms contributing to these behavioral effects, we examined the properties of primary afferent neurons innervating the stomach. The key results of the present study are 1) gastric afferent neurons in the nodose and DRG ganglion express potassium currents that can be distinguished based on their electrophysiological and pharmacological properties; 2) gastric ulceration decreases the transient outward current; and 3) gastric ulceration changes the electrophysiological properties of the transient outward current in nodose neurons. In the aggregate, these results suggest that hypersensitivity following gastric insult is contributed to, in part, by changes in K+ currents that increase excitability of gastric sensory neurons.

Consistent with previously published reports (11, 17, 43), we identified three electrophysiologically distinct components of the total outward currents in DRG and nodose ganglion neurons. In addition to the sustained potassium current, gastric sensory neurons express two inactivating outward currents that can be distinguished based on their kinetics of activation and inactivation and their voltage dependence of inactivation. The time course of activation was strongly voltage dependent for both components. Similarly, the decay of the rapidly inactivating current accelerated with stronger depolarization, a characteristic property of the rapidly inactivating potassium current in mammalian neurons (5, 11, 27). In contrast, the more slowly inactivating component decayed at rates that did not vary consistently over the voltage range tested. Similar results have been reported for brain stem neurons and the heterologously expressed human Kv4.3 channel (9, 14), whereas others (5, 11) have described a voltage-dependent acceleration of current decay in the slowly inactivating outward current as well. These apparent discrepancies may be, at least in part, due to different recording conditions, because calcium buffering can affect inactivation kinetics (7).

We used pharmacological approaches to further characterize the transient potassium currents. Prior studies (15, 17, 41) demonstrated that 4-AP inhibits both the fast and slowly inactivating potassium currents with increasing concentrations over a range from 0.1 to 5 mM, thus limiting its use to separate these components. The onset and recovery of this block are time dependent, with prolonged inhibition even at hyperpolarized voltages, thereby compromising cell viability (39). Therefore, a single high concentration of 4-AP was selected to test whether the pharmacological characteristics are consistent with prior reports. The reduction of both the fast and slow components of the transient outward current by 4-AP confirms that this current has the typical pharmacological properties of an A current (11, 12, 27, 43). Whereas some investigators (11, 41) have reported an inhibition of transient outward currents by DTX, we and others (16) did not observe such a selective effect on A currents in visceral sensory neurons. This difference between neurons innervating the skin (11) and the generally unmyelinated visceral sensory neurons may be due to differential expression of potassium channel subunits (13, 19). Such a differential expression of potassium channel subunits has been documented for DRG neurons (34).

Gastric ulceration altered the properties of potassium currents in both DRG and nodose ganglia by decreasing the fast and slow components of the transient outward current, whereas the sustained potassium current remained unchanged. We also noted a change in the voltage dependence of activation and inactivation of the rapidly inactivating current, with a shift to more hyperpolarized potentials in nodose neurons, which will further decrease the number of channels that can be activated on depolarization from the resting membrane potential. Similar results have previously been described in visceral sensory neurons studied during inflammation of the bladder (42), whereas axotomy decreased potassium channel expression (22) or current density without altering channel properties (10). Such differential effects of organ inflammation compared with nerve injury have also been reported for sodium currents (1, 8).

Ishikawa et al. (22) recently reported selective downregulation of Kv1.1–3 and Kv2.1 proteins after axotomy and suggested that the reduction in expression of these K+ channel proteins may be responsible for the reduction in A current observed in the same model of injury (11). Similarly, changes in the expression and distribution of sodium channels have been described in response to inflammation within the target organs or axotomy (3, 18). The sparse innervation of the gastrointestinal tract with <1% of DRG neurons projecting to the stomach did not allow us to examine the molecular changes that underlie the decrease in A current density in more detail. However, others (10) have attributed the reduction of A current in cutaneous neurons after axotomy to selective downregulation of potassium channel subunits. Furthermore, potassium channel Kv1.1–2 and 1.6 proteins have been reported to influence the excitability of rat visceral sensory neurons (16).

Interestingly, we also noted a change in the size of nodose neurons, assessed by measuring the whole cell capacitance. Although we did not systematically study this and cannot reach firm conclusions based on our data, the small number of labeled gastric neurons argues against a significant selection bias. Moreover, similar findings have previously been reported in response to inflammation (1, 29, 42). The changes in voltage-sensitive currents and cell size suggest that mediators affect the soma of afferent neurons projecting to the inflamed area. Neurotrophins have been identified as important target-derived factors involved in injury and tissue repair mechanisms. Functional receptors for neurotrophins have been identified in the endings of afferent and efferent nerves (21, 29). These factors play a critical role in maintaining or modulating the functions of neurons. This is consistent with reports demonstrating degenerative changes in efferent neurons after axonal injury (31). Conversely, inflammation is associated with increases in production and release of growth factors, which may alter the electrophysiological and neurochemical properties of neurons (2, 38, 40). Zhang et al. (45) recently demonstrated a rapid change in potassium current due to nerve growth factor (NGF)-induced generation of ceramide. However, this fast effect is reversible and should thus not persist 2–8 h after cell dissociation and culture. Others (4) have described NGF-dependent changes in potassium channel expression. However, the exact role of NGF in regulating the expression of ion channels and altering neuron excitability remains unclear.

With their rapid activation and inactivation kinetics, transient potassium currents regulate action potential duration and firing patterns in neurons (23, 37, 42). Repeated depolarizations will progressively inactivate the transient currents, thereby prolonging individual action potentials and increasing the number of spikes generated during longer lasting stimulations (25). Consistent with previous publications on altered excitability of sensory neurons innervating the ileum and bladder (30, 42), we observed a reduction in action potential threshold and an increase in the number of spike discharges to current injections in gastric sensory neurons from a similar rat model of gastric ulcers (K. Dang, K. Bielefeldt, and G. F. Gebhart, unpublished observations). Considering the steep relationship between intracellular calcium concentrations and transmitter release (23), prolongations of the action potential will prolong calcium inflow through voltage-dependent calcium channels and thereby facilitate signal transmission at the synapse.

The stomach receives dual innervation through vagal and spinal afferent neurons. Although it is generally accepted that spinal pathways mediate nociceptive signaling (36), vagal afferents may contribute to chemonociception, discomfort, and emotive components associated with aversive stimuli (28, 44). Although we did not observe changes in outward currents after induction of mild superficial gastritis with iodoacetamide (1), the present results document that a more severe form of gastric inflammation altered the properties of both vagal and spinal afferents, consistent with a sensitization of both sensory pathways. These results confirm prior in vivo experiments demonstrating that inflammatory mediators rapidly sensitized vagal and spinal afferent fibers (33), suggesting that sensitized vagal afferents may contribute to dyspeptic symptoms.


    ACKNOWLEDGMENTS
 
We thank M. Burcham for graphics assistance.

GRANTS

This work was supported by National Institutes of Health Grants DK-01548, NS-35790, and NS-19912.


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. Dang, Dept. of Pharmacology, Univ. of Iowa, Iowa City, IA 52242 (E-mail: khoa-dang{at}uiowa.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Bielefeldt K, Ozaki N, and Gebhart GF. Experimental ulcers alter voltage-sensitive sodium currents in rat gastric sensory neurons. Gastroenterology 122: 394-405, 2002.[ISI][Medline]
  2. Bielefeldt K, Ozaki N, and Gebhart GF. Role of nerve growth factor in modulation of gastric afferent neurons in the rat. Am J Physiol Gastrointest Liver Physiol 284: G499-G507, 2003.[Abstract/Free Full Text]
  3. Black JA, Cummins TR, Plumpton C, Chen YH, Hormuzdiar W, Clare JJ, and Waxman SG. Upregulation of a silent sodium channel after peripheral, but not central, nerve injury in DRG neurons. J Neurophysiol 82: 2776-2785, 1999.[Abstract/Free Full Text]
  4. Boettger MK, Till S, Chen MX, Anand U, Otto WR, Plumpton C, Trezise DJ, Tate SN, Bountra C, Coward K, Birch R, and An P. Calcium-activated potassium channel SK1- and IK1-like immunoreactivity in injured human sensory neurones and its regulation by neurotrophic factors. Brain 125: 252-263, 2002.[Abstract/Free Full Text]
  5. Cooper E and Shrier A. Inactivation of A current and A channels in rat nodose neurons. J Gen Physiol 94: 881-910, 1989.[Abstract]
  6. Dang K, Bielefeldt K, and Gebhart GF. Gastric ulcers reduce A-type potassium current in rat sensory ganglia (Abstract). Gastroenterology 124: A-344, 2003.
  7. Deschenes I, DiSilvestre D, Juang GJ, Wu RC, An WF, and Tomaselli GF. Regulation of Kv43 current by KChIP2 splice variants: a component of native cardiac I(to)? Circulation 106: 423-429, 2002.[Abstract/Free Full Text]
  8. Dib-Hajj S, Black JA, Felts P and Waxman SG. Down-regulation of transcripts for Na channel alpha-SNS in spinal sensory neurons following axotomy. Proc Natl Acad Sci USA 93: 14950-14954, 1996.[Abstract/Free Full Text]
  9. Dilks D, Ling HP, Cockett M, Sokol P, and Numann R. Cloning and expression of the human Kv4.3 potassium channel. J Neurophysiol 81: 1974-1977, 1999.[Abstract/Free Full Text]
  10. Everill B and Kocsis JD. Reduction in potassium currents in identified cutaneous afferent dorsal root ganglion neurons after axotomy. J Neurophysiol 82: 700-708, 1999.[Abstract/Free Full Text]
  11. Everill B, Rizzo MA, and Kocsis JD. Morphologically identified cutaneous afferent DRG neurons express three different potassium currents in varying proportions. J Neurophysiol 79: 1814-1824, 1998.[Abstract/Free Full Text]
  12. Fedulova SA, Vasilyev DV, and Veselovsky NS. Voltage-operated potassium currents in the somatic membrane of rat dorsal root ganglion neurons: ontogenetic aspects. Neuroscience 85: 497-508, 1998.[CrossRef][ISI][Medline]
  13. Ficker E and Heiemann U. Slow and fast transient potassium currents in cultured rat hippocampal cells. J Physiol 445: 51-63, 1992.
  14. Funahashi M, Mitoh Y, and Matsuo R. Two distinct types of transient outward currents in area postrema neurons in rat brain slices. Brain Res 942: 31-45, 2002.[CrossRef][ISI][Medline]
  15. Furukawa Y, Kandel ER, and Pfaffinger P. Three types of early transient potassium currents in Aplysia neurons. J Neurosci 12: 98-1000, 1992.
  16. Glazebrook PA, Ramirez AN, Schild JH, Shieh CC, Doan T, Wible BA, and Kunze DL. Potassium channels Kv11, Kv12 and Kv16 influence excitability of rat visceral sensory neurons. J Physiol 541: 467-482, 2002.[Abstract/Free Full Text]
  17. Gold MS, Shuster MJ, and Levine JD. Characterization of six voltagegated K+ currents in adult rat sensory neurons. J Neurophysiol 75: 2629-2646, 1996.[Abstract/Free Full Text]
  18. Gold MS, Weinreich D, Kim CS, Wang R, Treanor J, Porreca F, and Lai J. Redistribution of NaV18 in uninjured axons enables neuropathic pain. J Neurosci 23: 158-166, 2003.[Abstract/Free Full Text]
  19. Halliwell JV, Othman IB, Pelchan-Mathews A, and Dolly JO. Central action of dendrotoxin: selective reduction of transient K conductance in hippocampus and binding to localized acceptors. Proc Natl Acad Sci USA 83: 493-497, 1986.[Abstract]
  20. Harvey AL. Recent studies on dendrotoxins and potassium ion channels. Gen Pharmacol 28: 7-12, 1997.[CrossRef][Medline]
  21. Helke CJ, Adryan KM, Fedorowicz J, Zhou H, Park JS, Curtis R, Radley HE, Distefano PS. Axonal transport of neurotrophins by visceral afferent and efferent neurons of the vagus nerve of the rat. J Comp Neurol 393: 102-117, 1998.[CrossRef][ISI][Medline]
  22. Ishikawa K, Tanaka M, Black JA, and Waxman SG. Changes in expression of voltage-gated potassium channels in dorsal root ganglion neurons following axotomy. Muscle Nerve 22: 502-507, 1999.[CrossRef][ISI][Medline]
  23. Jackson MB, Konnerth A, and Augustine GJ. Action potential broadening and frequency-dependent facilitation of calcium signals in pituitary nerve terminals. Proc Natl Acad Sci USA 88: 380-394, 1991.[Abstract]
  24. Kerr PM, Clement-Chomienne O, Thorneloe KS, Chen TT, Ishii K, Sontag DP, Walsh MP, and Cole WC. Heteromultimeric Kv12-Kv15 channels underlie 4-aminopyridine-sensitive delayed rectifier K+ current of rabbit vascular myocytes. Circ Res 89: 1038-1044, 2001.[Abstract/Free Full Text]
  25. Kocsis JD, Ruiz JA, and Waxman SG. Maturation of mammalian myelinated fibers: changes in action potential characteristics following 4-aminopyridine application. J Neurophysiol 50: 449-463, 1983.[Free Full Text]
  26. McClesky EW. and Gold MS. Ion channels of nociception. Annu Rev Physiol 61: 835-856, 1999.[CrossRef][ISI][Medline]
  27. McFarlane S and Cooper E. Kinetics and voltage-dependence of A-type currents on neonatal rat sensory neurons. J Neurophysiol 66: 1380-1391, 1991.[Abstract/Free Full Text]
  28. Michl T, Jocic M, Heinemann A, Schuligoi R, and Holzer P. Vagal afferent signaling of a gastric mucosal acid insult to medullary, pontine, thalamic, hypothalamic and limbic, but not cortical, nuclei of the rat brain. Pain 92: 19-27, 2001.[CrossRef][ISI][Medline]
  29. Mo ZL, Katafuchi T, and Hori T. Effects of IL-1 on neuronal activities in the dorsal motor nucleus of the vagus in rat brain slices. Brain Res Bull 41: 249-255, 1996.[CrossRef][ISI][Medline]
  30. Moore BA, Stewart TM, Hill C, and Vanner SJ. TNBS ileitis evokes hyperexcitability and changes in ionic membrane properties of nociceptive DRG neurons. Am J Physiol Gastrointest Liver Physiol 282: G1045-G1051, 2002.[Abstract/Free Full Text]
  31. Navaratnam V, Jacques TS, and Skepper JN. Ultrastructural and cytochemical study of neurones in the rat dorsal motor nucleus of the vagus after axon crush. Microsc Res Tech 42: 334-344, 1998.[CrossRef][ISI][Medline]
  32. Ozaki N, Bielefeldt K, Sengupta JN, and Gebhart JF. Models of gastric hyperalgesia in the rat. Am J Physiol Gastrointest Liver Physiol 283: G666-G676, 2002.[Abstract/Free Full Text]
  33. Ozaki N and Gebhart GF. Characterization of mechanosensitive splanchnic nerve afferent fibers innervating the rat stomach. Am J Physiol Gastrointest Liver Physiol 281: G1449-G1459, 2001.[Abstract/Free Full Text]
  34. Rasband MN, Park EW, Vanderah TW, Lai J, Porreca F, and Trimmer JS. Distinct potassium channels on pain-sensing neurons. Proc Natl Acad Sci USA 98: 13373-13378, 2001.[Abstract/Free Full Text]
  35. Rothman JS and Manis PB. Differential expression of three distinct potassium currents in the ventral cochlear nucleus. J Neurophysiol 89: 3070-3082, 2003.[Abstract/Free Full Text]
  36. Sengupta JN and Gebhart GF. Gastrointestinal afferent fibers and sensation. In: Physiology of the Gastrointestinal Tract (3rd ed.). New York: Raven, 1994, p. 483-519.
  37. Stansfeld CE, Marsh SJ, Halliwell JV, and Brown DA. 4-Aminopyridine and dentrotoxin induce repetitive firing in rat visceral sensory neurons by blocking slowly inactivating outward current. Neurosci Lett 64: 299-304, 1986.[CrossRef][ISI][Medline]
  38. Toma H, Winston J, Micci MA, Shenoy M, and Pasricha PJ. Nerve growth factor expression is up-regulated in the rat model of L-arginine-induced acute pancreatitis. Gastroenterology 119: 1373-1381, 2000.[ISI][Medline]
  39. Wagoner PK and Oxford GS. Aminopyridines block an inactivating potassium current having slow recovery kinetics. Biophys J 58: 1481-1489, 1990.[Abstract]
  40. Winston J, Toma H, Shenoy M, and Pasricha PJ. Nerve growth factor regulates VR-1 mRNA levels in cultures of adult dorsal root ganglion neurons. Pain 89: 181-186, 2001.[CrossRef][ISI][Medline]
  41. Wu RL and Barish ME. Two pharmacologically and kinetically distinct transient potassium currents in cultured embryonic mouse hippocampal neurons. J Neurosci 12: 2235-2246, 1992.[Abstract]
  42. Yoshimura N and de Groat WC. Increased excitability of afferent neurons innervating rat urinary bladder after chronic bladder inflammation. J Neurosci 19: 4644-4653, 1999.[Abstract/Free Full Text]
  43. Yoshimura N, White J, Weight F, and de Groat WC. Different types of Na+ and A-type K+ currents in dorsal root ganglion neurones innervating the rat urinary bladder. J Physiol 494: 1-16, 1996.[Abstract]
  44. Zagon A. Does the vagus nerve mediate the sixth sense? Trends Neurosci 24: 671-673, 2001.[CrossRef][ISI][Medline]
  45. Zhang YH, Vasko MR, and Nicol GD. Ceramide, a putative second messenger for nerve growth factor, modulates the TTX-resistant Na+ current and delayed rectifier K+ current in rat sensory neurons. J Physiol 544: 385-402, 2002.[Abstract/Free Full Text]