High-conductance chloride channels generate pacemaker currents in interstitial cells of Cajal

GASTROENTEROLOGY 2002;123:1627–1636

High-Conductance Chloride Channels Generate Pacemaker Currents in Interstitial Cells of Cajal JAN D. HUIZINGA,* YAOHUI ZHU,* JING YE,* and ARELES MOLLEMAN‡ *Department of Medicine, McMaster University, Hamilton, Ontario, Canada; and ‡Department of Biosciences, University of Hertfordshire, Hatfield, United Kingdom

Background & Aims: Interstitial cells of Cajal (ICCs) are responsible for slow, wave-driven, rhythmic, peristaltic motor patterns in the gastrointestinal tract. The aim was to identify and characterize the ion channels that generate the underlying pacemaker activity. Methods: Single ion channel recordings were obtained from nonenzymatically isolated ICCs and studied by using the cell attached and inside-out configurations of the patch clamp technique. Results: A high-conductance chloride channel was observed in ICCs that was spontaneously and rhythmically active at the same frequency as the rhythmic inward currents defining ICC pacemaker activity, 20 –30 cycles/min at room temperature. Main conductance levels occurred between 122–144 pS and between 185–216 pS. Periodicity in the channel opening coincided with periodicity in membrane potential change, hence, at the single channel level, chloride channels were seen to be associated with the generation of rhythmic changes in membrane potential. Conclusions: ICCs harbor high-conductance chloride channels that participate in the generation of pacemaker activity and may become a target for pharmacologic treatment of gut motor disorders.

nterstitial cells of Cajal (ICC) are responsible for the rhythmic, peristaltic, slow, wave-driven motor patterns1–5 developing in the small intestine. In fact, normal peristaltic activity in the small bowel after gastric emptying of a liquid was absent in W/Wv mutant mice, which do not have the network of ICCs that are associated with Auerbach’s plexus.6 Several gut motor disorders have been linked to abnormalities in ICCs.7–10 A major breakthrough in our understanding of gut pacemaker activity came when chemically isolated ICCs, and not smooth muscle cells, were seen to generate spontaneous rhythmic inward currents.11–13 These studies provided indirect evidence for involvement of nonselective cation channels11,12 and/or chloride channels13 based on pharmacologic manipulation of whole-cell currents. The objective of the present experiments was to identify pacemaker channels at the single-channel level. We succeeded by recording spontaneous rhythmic channel ac-

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tivity from electrodes sealed onto rhythmically contracting ICCs.

Materials and Methods Obtaining Single ICCs Single-channel currents were measured mostly from single, active ICCs, identified by vital staining with c-kit antibody (Gibco, Burlington, Ontario, Canada) coupled to Alexa-488 (Molecular Probes, Inc., Eugene, OR). Some recordings were made, as indicated, from ICCs that were close to the explant and attached to other ICCs and smooth muscle cells. The jejunum (about 1.5 cm long) was removed from newborn (i.e., 2- to 3-day-old) CD-1 mice. By using sharp dissection, the mucosa and submucosa were removed by cleaving at the deep muscular plexus, leaving most ICC associated with the deep muscular plexus (ICC-DMP) out of the preparation studied. Segments of 0.5 mm in width and length were dissected in culture medium M-199 (Gibco) and placed onto a collagencoated glass coverslip in a culture dish. The culture medium contained 10% fetal bovine serum, glutamine, and penicillin (Gibco). After 2–3 days, ICCs and smooth muscle cells grew out of the explants. The explants maintained rhythmic contraction patterns and individual ICCs even far from the explant could be seen to contract rhythmically.

Constructing Fluorescent Antibodies The conjugated ACK2 (anti-c-kit antibody; Gibco) was obtained by a labeling reaction by using the Alexa Fluor 488 Protein Labeling Kit (A-10235; Molecular Probes, Inc.). Briefly, the ACK2 protein solution (1 mg/1 mL) was mixed with a succinimidyl ester moiety of Alexa Fluor 488 for 90 minutes at room temperature to form a stable dye-protein combination. The labeled protein was purified by using column chromatography provided with the kit, resulting in separation of the conjugated protein from the free dye. A total Abbreviations used in this paper: DIDS, disodium 4,4ⴕ-diisothiocyanatostilbene-2,2ⴕdisulphonic acid; ICC, Interstitial cell(s) of Cajal; IP3 , inositol 1,4,5-triphosphate; I/V, current/voltage relationship; NMDG, N-methyl-D-glucamine; SITS, 4-acetamido-4ⴕ-isothiocyanatostilbene-2,2ⴕdisulphonic acid. © 2002 by the American Gastroenterological Association 0016-5085/02/$35.00 doi:10.1053/gast.2002.36549

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Figure 1. Cell superfusion set-up. Cell superfusion can be performed without disturbing the ion concentrations at the reference electrode. C identifies the patch pipette. The bath is perfused through A. The reference electrode is at position E. The outflow occurs through suction at position D. At position B, directly in front of the patch, a fast flow superfusion allows the patch to be bathed in experimental solutions without solution change at the reference electrode. The method is based on a design by Barajas-Lopez et al.14

amount of 1 mL of collected ACK2-Alexa 488 was aliquoted and stored at ⫺70°C. For preparation of single cells around an explant after 2–5 days of culture, the ACK2-dye was diluted with culture medium M-199 and added to the culture with a final concentration of 0.75–1 ␮g/mL. After 1 hour of incubation at 37°C with 95% O2 and 5% CO2, the fluorescent c-kit antibodies were bound to the extracellular part of the c-kit protein in the ICC cell membrane and c-kit–positive cells were identified under the fluorescent microscope after removal of unbound antibody.

Patch Clamp Recording Before formation of the high resistance seal between the pipette and cell membrane, junction potentials were nullified. Single-channel currents were recorded from c-kit–positive and contractile cells in the cell-attached and excised inside-out patch configurations. Pipettes were filled with solution containing (in mmol/L): 140 KCl, 5 NaCl, 1 MgCl2, 2 CaCl2, 10 HEPES, 10 glucose (pH level 7.30 –7.40 with KOH). In some experiments, KCl in the pipette solution was substituted with NaCl, CsCl, or N-methyl-D-glucamine (NMDG)-Cl. The pipette solution contained 140 mmol/L KCl unless otherwise stated. For inside-out recordings, the same solutions were used for bath perfusion except that calcium was reduced to 1 ␮mol/L. KCl was replaced as noted with Kmethanesulfonate as well as sucrose 185 mmol/L with KCl 50 mmol/L or sucrose 360 mmol/L. The osmolarity of the sucrose solutions was the same as that of the extracellular solutions. For cell-attached recordings, the extracellular solution was M199 culture medium containing (in mmol/L) NaCl 116, NaHCO3 26, NaH2PO4 1, Na-acetate 0.4, KCl 5.4, CaCl2 1.4, MgSO4 0.8, glucose 5.6. The optimal pipette tip resistance ranged from 7–10 M⍀. All experiments were conducted at a temperature of 25°C to 30°C. Data acquisition and analysis were performed by using the pClamp suite (Axon Instruments, Foster City, CA). For the inside-out patch recordings in which chloride was replaced with other ions, a fast flow system was used (Figure 1)

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designed by Barajas-Lopez et al.14 The bath was perfused with the bath solution as described earlier, in which the reference electrode was placed. The outflow suction was placed in position D. The patch pipette was placed in position C. At position B, in front of the patch, a fast flow system was placed that contained the bath solution except that KCl was partly replaced by K-methanesulfonic acid. In this way, the reference electrode did not see changes in chloride concentration and hence a liquid junction potential did not develop. The same system was also used to add drugs (i.e., the fast flow solution contained the bath solution plus disodium 4,4⬘-diisothiocyanatostilbene-2,2⬘disulphonic acid (DIDS), and so forth, as indicated in the Results section).

Data Analysis/Nomenclature By convention, upwardly deflected or positive current relates to positive current going out of the pipette or negative current entering the pipette. In the whole-cell configuration this is outward current, positive current going out of the cell or negative current going into the cell. In the cell-attached and inside-out configuration, positive current going out of the pipette actually goes from the outside to the inside of the cell membrane. To conform with standard practice, the measured currents have been reversed so that positive current reflects positive outward current (i.e., from inside to outside of the cell). In this article, positive (upward) current reflects Cl⫺ current going out of the pipette or from outside to inside of the cell. Negative (downward) current reflects Cl⫺ current going into the pipette or from inside to outside of the cell. In ICCs, Cl⫺ leaving the cell at the resting membrane potential results in a depolarizing current. In the whole-cell configuration, command potentials by the amplifier (i.e., the pipette potentials), control the membrane potential and hence are identical to the membrane potential. In the cell-attached configuration, the pipette potential does not control the cell membrane potential. The actual membrane potential is not known and in this article it is assumed to be ⫺65 mV, the average value we obtained from neonatal isolated ICCs. The membrane potential difference across the channels in the cell-attached patch (patch potential) is therefore assumed to be ⫺65 minus the pipette potential. In the insideout configuration, the membrane potential difference across the channels in the patch (patch potential) is the opposite of the pipette potential (i.e., pipette potential ⫻ ⫺1). Average values are expressed as mean ⫾ SE.

Results Identification of Interstitial Cells of Cajal in Culture by Spontaneous Rhythmic Electrical or Mechanical Activity and Fluorescence Explant culture allowed access to ICCs and smooth muscle cells, as isolated cells and as small networks of cells, after 2–5 days of culture (Figure 2). This alternative to chemical dissociation avoids any potential

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alteration of ion channels owing to proteolytic activity. ICCs were identified in culture within 1 or 2 days by their morphologic characteristics.11 These ICCs showed rhythmic inward currents (Figure 3) similar to those recorded from ICCs isolated from adult mice,11 at frequencies between 20 –30 cycles/min at room temperature. ICCs are contractile in situ15 and were rhythmically contractile in culture (this study and others16,17). The ICCs thus identified stained positive with ACK2 (antic-kit antibodies) that were coupled to the fluorescent

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Figure 3. Whole-cell rhythmic currents. ICCs, identified as in Figure 2, showed rhythmic inward currents, at a frequency similar to those observed with freshly isolated cells.11 The cell had been in culture 4 days and the recording was obtained at a membrane potential of ⫺70 mV. The figure shows activity at 24 cycles/min.

compound Alexa 488 (Figure 2). This created brightly fluorescent ICCs though having no effect on their spontaneous activity. Periodic Channel Activity Observed at 20 – 30 Cycles per Minute in the Cell-Attached Configuration Spontaneous rhythmic channel activity was observed using electrodes sealed onto rhythmically contracting ICCs (Figure 2). In the cell-attached patch configuration, channel activity was recorded from a very small patch of membrane that was still part of the living cell. Channel activity was observed at a conductance level of either 122–144 pS (n ⫽ 8) or 185–216 pS (n ⫽ 5) or multiples thereof (Figures 4, 5, and 6). When channel activity was observed at 122–144 pS conductance, only rarely were channel closings seen to a conductance level of 60 –70 pS. When channel activity was observed at a 185–216 pS conductance level, occasionally channel closing was seen at conductance levels of ⬃130 pS or ⬃65 pS. The lower conductance levels were better distinguishable in the inside-out configuration (see later). In some experiments, in the cell-attached configuration, Š Figure 2. Recording of rhythmic channel activity from a positively identified ICC. (A) ICCs were identified by morphology, rhythmic contractile activity, and staining by ACK2 c-kit antibody coupled to fluorescent Alexa 488. Individual ICCs grew in culture out of small clumps of cells (explants) taken from the intestine of a 2- to 3-day-old mouse. This image was taken by a confocal microscope. Note that some c-kit–positive cells were attached to the explant whereas others were isolated and relatively far away from the explant. The cell recorded from (C) is identified by a black arrow. (B) The same image obtained with a bright field objective. Note that only relatively few cells are positive using the c-kit antibody. (C) The dish was placed in the electrophysiology set-up by using a different inverted microscope. The picture was taken while the recording was obtained, the large black triangle is the pipette out of focus. By using a cell-attached patch with a high K solution in the pipette and culture medium (145 mmol/L Na) in the bath solution, periodic channel activity was revealed as shown. The applied patch potential was ⫺80 mV. See the Materials and Methods section for complete composition of solutions. Scale bars: 10 pA, 2 s.

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Figure 4. Spontaneous rhythmic channel activity recorded with an electrode attached to a rhythmically contracting ICC. (A) Spontaneous inward currents recorded from an ICC in the cell-attached configuration. C, closed state; O, open state. The channel opened in a rhythmic manner at hyperpolarized patch potentials including the resting membrane potential of the cell. The frequency of the periods of activation was 18 cycles/min, similar to that of observed rhythmic whole-cell currents. Assuming a resting membrane potential of ⫺65 mV, the reversal potential was ⫺25 mV. Channels opened to a conductance level of 144 pS. Lower conductance levels were observed rarely. The second tracing shows a conductance level at half the dominant level, observed at a patch potential of ⫺5 mV. The graph shows the relationship between current amplitude and voltage, the latter expressed as membrane potential across the patch (patch potential). (B) In another ICC, 5 dominant conductance levels were observed in a cell-attached patch; at a patch potential of ⫺100 mV, unitary current amplitudes of 11 pA were obtained as shown in the current amplitude histogram. C, closed state; O, open state. The histogram indicates the relative time the channels were in the different open states. The closed state was determined by other sections of this recording (not shown). From current/voltage relationship (I/V) curves obtained at all conductance levels, the dominant conductance levels were 122, 244, 360, and 480 pS. Occasional current transitions of 5.5 pA (see inset at arrow) suggest a subconductance or existence of cochannels at a conductance of 67 pS (see Discussion section). The amplitude histogram was obtained from a 26-s recording and the major ticks represent 1000 events. The I/V curve expresses current amplitudes based on the first major conductance level obtained at the different holding potentials (expressed as membrane potential across the patch).

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Figure 5. Channel activity is not influenced by substitution of Na⫹ by Cs⫹ or NMDG⫹. A ramp protocol was applied to inside-out patches in which the patch potential was stepped from 0 to ⫺60 mV for 100 ms followed by a ramp to ⫹90 mV over a time period of 300 ms. Each figure shows two 300 ms sweeps. Currents are shown without additional filtering at the voltages between ⫺60 and 90 mV. The extracellular (pipette) solution contained 140 mmol/L KCl. The intracellular (bath) solution contained 140 mmol/L NaCl (top figure). In the other graphs (2– 4), 140 mmol/L NaCl was replaced by 140 mmol/L CsCl, 140 mmol/L NMDGCl, or 385 mmol/L sucrose. See the Methods section for complete description of solutions. When the patch potential was changed from 0 to ⫺60 mV, channel activity was seen immediately at a conductance level of 140 pS (Na⫹), 140 pS (Cs⫹), and 134 pS (NMDG⫹). After 100 ms, the patch potential changed gradually from ⫺60 to ⫹90 mV as shown. In many sweeps, the channel(s) remained open for the duration of the ramp protocol. In the sweeps shown, cochannel closings can be seen at patch potentials between ⫹50 mV and ⫹90 mV. This type of experiment shows that the main conductance of the channel is 140 pS, maintained over a large voltage range. At more depolarized potentials a subconductance is revealed, or evidence of the existence of cochannels (see Discussion section). The reversal potentials were 2 mV (Na), 2 mV (Cs), and ⫺1 mV (NMDG). In 5 such experiments, neither the conductances nor the reversal potentials were significantly different between results obtained with different cations. When NaCl was replaced by sucrose, no current went into the pipette. However, the open probability of the channel dramatically reduced such that currents observed leaving the pipette were rare.

prolonged quiescent periods (1–5 s) alternated with periods of channel activity. The period of high open probability occurred between 20 –30 times/min determined from recordings as in Figures 2C and 4A in which periodic activity was clearly separated by long periods of quiescence. This frequency is similar to that of the rhythmic inward currents observed in the whole-cell configuration.18 The I/V curve of the earlier-described

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Figure 6. Revealing subconductances or cochannels by ramp protocols. Channel activity was observed with NMDG-Cl on both sides of an inside-out patch. The tracing in the middle section of the figure shows channel activity obtained with a pulse protocol in which a holding potential corresponding to a patch potential of ⫹20 mV was held for 13 seconds. Large current transitions were observed to conductance levels of 184 pS and 360 pS. The I/V curve shown represents the first conductance level. Three seconds after this experiment was completed, under identical conditions, a ramp protocol (see legend of Figure 5) was applied (bottom panel). Currents shown indicate a maximum channel conductance of 185 pS. At depolarizing potentials above 40 mV, currents decrease to subconductance levels of 126 and 62 pS, or alternatively it can be interpreted as showing single cochannel activity of 62 pS. Because most current transitions are to a level of 182 pS, the individual channels or cochannels show a high degree of cooperativity. The figure shows a composite of 3 sweeps.

channel was linear from ⫺130 to ⫹ 40 mV patch membrane potential, assuming a cell membrane potential of ⫺65 mV, which was the average value of the resting membrane potential of neonatal isolated ICCs. Periodic, rhythmic channel activity was seen with high Na⫹ or high K⫹ solution in the pipette and culture medium solution in the bath (Figures 2C and 4). Under these conditions, the reversal potential was ⫺34 ⫾ 5 mV (n ⫽ 5). This is consistent with a chloride equilibrium potential; the intracellular chloride concentration in smooth muscle is estimated to be high (42–70 mmol/L) and the reversal potential in smooth muscle is estimated to be between ⫺24 and ⫺40 mV.19,20 Studies of the Channel in the Inside-Out Configuration After switching to the inside-out patch configuration, the bursting nature of the channel activity was

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lost and the channel(s) remained open for prolonged periods of time. In the inside-out configuration, a patch of membrane is excised from the cell and the amplifier controls the voltage over the patch of membrane attached to the recording electrode. Ionic composition at both sides of the patch can be controlled. The most dominant conductance levels were 139 ⫾ 10 pS (n ⫽ 15), and 195 ⫾ 12 pS (n ⫽ 8). With KCl or NaCl in the pipette, and KCl, NaCl, or CsCl in the bath, the reversal potential was ⫺3.5 ⫾ 1.0 mV, and no significant difference in reversal potential or channel conductance was noted between the different permeant cations (n ⫽ 25) (Figure 4). Lower conductance levels were observed with ramp protocols in which the patch potential was stepped from 0 to ⫺60 mV for 100 ms followed by a ramp to ⫹90 mV over a time period of 300 ms. Between patch potentials of ⫹60 and ⫹90 mV, channel conductance dropped occasionally to the 60 –70 pS level or a multiple thereof or to the closed state (Figures 5, 6, 7, and 8). When the main cation was exchanged in the bath for NMDG⫹, no shift in reversal potential occurred (n ⫽ 10) (Figure 4). This result is most consistent with the channel primarily carrying Cl⫺. Substitution of Na⫹ or K⫹ with NMDG⫹, which has a molecular weight of 195, did not affect open probability as judged from ramp protocol experiments (Figures 5 and 6). When KCl or NaCl at the intracellular side of the channel was replaced by sucrose, no currents were flowing into the pipette, but the open probability was dramatically reduced such that reversal potentials could not be established reliably (Figure 5). Further evidence for designating the channel as a chloride channel was obtained by changing the chloride gradient across the patch. When the pipette contained 140 mmol/L KCl and the superfusion was changed from one containing 140 mmol/L KCl to one containing 50 mmol/L KCl plus 185 mmol/L sucrose, the reversal potential shifted from 0 –3 mV to ⫺20 ⫾ 5 mV (n ⫽ 5) (Figure 7). A similar shift of ⬃⫺20 mV was observed when NaCl in the bath was replaced by Na methane sulfonic acid or isotheonate (not shown). Replacement of Cl⫺ ions at the intracellular site by sucrose or methane sulfonic acid markedly reduced open probability of the channel. As a consequence, current-voltage relationships in reduced intracellular Cl⫺ were often studied with reduced channel activity as in Figure 7. A third strong indication that Cl⫺ was the main current carrier was that the chloride channel blocker DIDS (100 ␮mol/L) in a time-dependent manner blocked channel activity (n ⫽ 3) (Figure 8). Inhibition was also observed with 4-acetamido-4⬘-isothiocyanatos-

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Figure 7. Reduction in chloride concentration causes shift in reversal potential. In the inside-out configuration (similar to experiments shown in Figure 5), the bath solution contained 140 mmol/L KCl. Then the patch was superfused through a fast flow system with 140 mmol/L KCl replaced by 50 mmol/L KCl and 185 mmol/L sucrose (see Figure 1). In 140 mmol/L KCl, the dominant conductance level was 217 pS and the reversal potential was 4 mV. When part of the KCl was replaced by sucrose, the open probability decreased and the dominant conductance level was 64 pS, and the reversal potential shifted from ⫹4 to ⫺20 mV.

tilbene-2,2⬘disulphonic acid (SITS) (100 ␮mol/L) (not shown).

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219 pS) at the different amplitudes. It follows that the differences in amplitude should not be attributed to changes in channel conductance, but to a rhythmically changing cell membrane potential giving a rhythmically changing driving force. From 2 corresponding points from the 2 I/V curves, the difference in cell membrane potential between low and high amplitude currents can be calculated. For example, at 0 mV holding potential, the highest and lowest current amplitudes were ⫺9.4 pA and ⫺13.6 pA; with a channel conductance of 216 pS, the difference in membrane potential was ⬃19 mV. Membrane potential oscillations of this magnitude corresponded to those measured in whole-cell recordings from ICCs.11 At 0 mV patch potential, when Cl⫺ current flowed from the pipette into the cell, the driving force was highest when the membrane potential was in its depolarized phase (⬃⫺46 mV), hence, the area of highest current amplitudes, and was lowest when the mem-

Rhythmic Changes in Open Probability Consistent With Rhythmic Generation of Depolarizing Current Previously we observed transient rhythmic depolarizations (slow waves) in ICCs by using whole-cell current clamp recording.11 Now we report direct evidence that the high-conductance chloride channel is associated with these rhythmic membrane potential changes in ICCs. This was accomplished by recording single-channel activity from ICCs, close to the explants, that were undergoing rhythmic membrane potential changes. Note that in the cell-attached configuration, the amplifier cannot control the cell membrane potential, although a perfect seal is obtained and channel activity can be recorded very accurately. Figure 9 shows chloride channel activity at 216 pS. With 140 mmol/L KCl in the recording pipette, and 140 mmol/L NaCl in the bath surrounding the cell, the I/V relationship revealed a reversal potential of ⫺23 mV. Two critical observations connected rhythmic membrane potential changes with rhythmic chloride channel activity. First, during the open state of the channels, a rhythmically fluctuating current amplitude is seen (Figure 9). The closed state did not oscillate, excluding the possibility that the oscillation could be artifactual. We obtained I/V curves by averaging values from 500-ms sections of the current amplitudes taken from at least 5 regions of highest amplitude and 5 regions of lowest amplitude at each of the different holding potentials. From these I/V curves (Figure 9A), it can be concluded that the channel conductance was the same (⬃216 and

Figure 8. Effect of DIDS on channel activity. A ramp protocol was applied in which the patch potential was gradually changed from ⫺60 mV to ⫹90 mV over a time period of 300 ms. Inside-out configuration with high K solution in bath and pipette. Top: control conditions. The figure shows a composite of 3 sweeps. Twenty sweeps showed channel activity at 182 pS, 130 pS, and 75 pS. In all sweeps, the 182 and 130 conductance levels were dominant. Middle: Two minutes after addition of DIDS (100 ␮mol/L) in the bath solution. The figure shows a composite of 3 sweeps. In 20 sweeps, the 130 pS and the 75 pS conductance levels were dominant. The channel was closed part of the time at hyperpolarizing and depolarizing potentials as shown. Bottom: Four minutes after addition of DIDS. The figure shows a composite of 2 sweeps. In all 20 sweeps, the channel was closed most of the time.

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brane potential was at its hyperpolarized phase (assumed resting membrane potential of ⫺65 mV). Second, when the open probability of the channel was assessed in 500-ms time spans at the most depolarized and the most hyperpolarized portion of the slow wave, the open probability was consistently higher in the depolarized phase (Figure 9B). When open probability was assessed at various holding potentials (Figure 9A), there was no relationship between patch potential and open probability. Therefore, open probability changes were not evoked by voltage changes but by intracellular events such as changing levels of calcium. Hence, periodic increases in chloride channel activity coincided with periodic depolarizations of the cell, directly linking the chloride channel activity to the rhythmic depolarizations of ICCs.

Discussion

Figure 9. Periodic increase in amplitude and open probability reflect channel activity generating slow wave in membrane potential. Cellattached configuration with 140 mmol/L K⫹ in the pipette and culture medium in the bath. The period frequency of the rhythmic changes was 19 cycles/min, similar to that of the rhythmic whole-cell inward currents described previously. The ICC that was patched in this experiment was connected to other ICCs close to the explant that was rhythmically contracting at 19 cycles/min, hence the ICC, but not smooth muscle cells, will generate rhythmic depolarizations as shown previously.11 (A) At a patch potential of 0 mV (⫺65 mV pipette potential), hence, at the expected resting membrane potential, the single-channel current was outward, corresponding to chloride entering the cell. The single-channel amplitude was seen to change in a rhythmic fashion but only when the channel was open. The bottom of the recording represents the closed state, no oscillation occurred here. In the expanded trace, the left part shows high amplitude channel activity whereas the right part shows relatively low amplitudes. The hypothesis is that the current amplitude fluctuates because of changes in cell membrane potential caused by slow wave activity, hence, cyclic changes in driving force for the ions occurs (see text). At 0 mV, when Cl⫺ current flowed from the pipette into the cell, the driving force was highest when the membrane potential was in its depolarized phase (⬃⫺46 mV), hence, the area of highest current amplitudes, and it was lowest when the membrane potential was in its hyperpolarized phase (assumed resting membrane potential of ⫺65 mV). It can be seen that the open probability is highest during the depolarized phase of the slow wave. The I/V curve (patch potential

Our data show that single ICCs, obtained without chemical dispersion from the mouse small intestine, display periodic activation of high-conductance chloride channels associated with spontaneous rhythmic membrane potential changes. The dominant conductance levels were between 122–144 pS and between 185–216 pS or multiples thereof. There is significant variability in the main conductance level of high-conductance chloride channels, the main conductance level of this channel in lymphocytes was 139 ⫾ 50 pS (n ⫽ 8, mean ⫾ SD),21 and similar variability was observed in other studies.22,23 Because the conductance levels in experiments in which the dominant conductance levels were between 122–144 pS and/or between 185–216 pS occasionally dropped to ⬃60 –70 pS (see Figures 4, 5, 6, and 8), it may be suggested that the high-conductance channel is comŠ vs. current amplitude) was drawn from data at both the depolarized phase (E) and the hyperpolarized phase (‚). The reversal potential at the resting membrane potential was ⫺20 mV, assuming a cell membrane potential of ⫺65 mV. The graph at bottom right shows channel open probability vs. patch potentials. At all holding potentials, the open probability in the depolarized phase was higher compared with the open probability in the hyperpolarized phase. However, there was no relationship between open probability and holding potential, indicating that not depolarization but an intracellular event was driving the open probability and hence the generation of outward current. (B) At the resting membrane potential (with a holding potential of 0 mV pipette potential), the current was inward, corresponding to chloride leaving the cell. In this situation, the largest driving force was experienced during the most hyperpolarized phase, in which, hence, the current amplitude was highest. It is again seen that the open probability was the highest during the depolarized phase, that is, when the lowest amplitude currents were seen. The graphs show open probability of 53% during the hyperpolarized phase and 92% during the depolarized phase.

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posed of 2 or 3 highly cooperative cochannels of 60 –70 pS conductance. In pulmonary alveolar epithelial cells where the main conductance level of a chloride channel was 404 pS, the conductance of the cochannels was thought to be 60 –70 pS.24 Interestingly, in these cells the channel openings also exhibited a bursting behavior. In rabbit longitudinal smooth muscle cells a high-conductance chloride channel was observed with a main conductance level of 309 pS, assumed to be obtained through cooperation of channels of ⬃80 pS conductance.25 Cooperative channel behavior was also suspected with high-conductance chloride channels observed in lymphocytes.21 Interestingly, whereas we observed these channels to be spontaneously active in ICCs, they were only activated in rabbit longitudinal smooth muscle cells by NK1 agonists. In lymphocytes, spontaneous activity of high-conductance chloride channels did occur and it was strongly temperature sensitive. We were able to record high-conductance chloride channel activity in the cell-attached mode from ICCs that were undergoing rhythmic membrane potential changes. In a rhythmic manner, single-channel open probability increased over a time period of ⬃500 ms and concomitantly the cell depolarized as reflected by a decrease in channel amplitude (driving force becomes smaller on depolarization) at a holding potential of 0 mV (patch at resting membrane potential). Clearly, increased ion channel activity contributed to the depolarization. Whether or not it was the only channel contributing to this depolarization is not known. There was likely little effect of depolarization on channel activity because we did not observe a relationship between open probability and patch potential as studied by changing the holding potential. These data are consistent with a high-conductance chloride channel being a pacemaker channel activated by intracellular factors. A role for chloride channels in gut pacemaker activity was suggested previously based on pharmacologic manipulation of rhythmic whole-cell currents from ICCs,13 and chloride channels also were found to be associated with pacemaker activity in rabbit urethra.26 The identification of a role for high-conductance chloride channels in slow-wave generation in tissue experiments using ion replacements is difficult because under certain conditions these channels may be quite permeable to cations.25 Therefore, under certain conditions these ion channels may be rather nonselective. Chloride channels likely are not the only channel that is involved in pacemaker activity. Other studies on rhythmic whole-cell currents on isolated ICCs suggested a role for nonselective cation channels.11,12

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We observed a complete inhibition of the high-conductance channel by DIDS (100 ␮mol/L). Recent studies in our laboratory showed that DIDS or SITS markedly reduced the slow wave amplitude and rate of increase.27 SITS also inhibited whole-cell currents obtained from kit-positive cells, isolated from the mouse small intestine.13 These data provide strong evidence for a role of Cl channels in the generation of the upstroke of the slow wave. The slow wave activity measured in tissue, and the rhythmic inward currents generated by ICCs, are not very sensitive to voltage changes but are highly temperature sensitive and easily modified by intracellular metabolic changes.28,29 The details of intracellular activation of the high-conductance chloride channel in ICCs remains to be investigated; it is interesting to note that indirect evidence in the small intestine points to inositol 1,4,5-triphosphate (IP3)-mediated activation of slow waves and the whole-cell rhythmic inward currents in ICCs can be blocked by IP3-receptor antagonists.28,30 These findings suggest IP3-mediated activation of chloride channels in ICCs, which is consistent with the fact that IP3 can activate the high-conductance chloride channels in colonic smooth muscle.25 Thuneberg and Peters16 showed the characteristics of spontaneously contractile ICCs grown within and out of a tissue explant. These were ICC associated with Auerbach’s plexus (ICC-AP) as judged by methylene blue accumulation (ICC-DMP do not accumulate methylene blue) and electron microscopy.17 We provided evidence for contracting ICCs in short-term culture at http://www.fhs.mcmaster.ca/huizinga/nature.htm. In the present study, c-kit–positive ICCs, grown out of explants without the use of digestive enzymes (see Figure 2), were rhythmically contractile, not owing to muscle contraction. Often only ICC processes contracted rhythmically and much more forcefully than surrounding smooth muscle cells similar to observations made by Thuneberg and Peters.16 Rhythmic contractile activity is consistent with the occurrence of calcium oscillations in ICCs at a similar frequency.31 We have no evidence whether or not the spontaneous contractile activity is always linked to slow-wave generation although rhythmic contractile activity occurs in the same frequency range as the slowwave activity. The contractile nature of ICCs is controversial because most ICCs identified by electron microscopy lack thick filaments. However, ICC-DMP in the canine small intestine showed myosin light chain– like immunoreactivity and electron microscopy showed thick filaments.32 Freshly isolated ICCs were seen to lack smooth muscle myosin messenger RNA33 but there are

November 2002

about 20 different types of myosin, most not studied in ICCs. Although ICCs are contractile cells, the exact molecular nature of the generation of contraction still is to be resolved. Pacemaker activity in the gut is not primarily controlled by voltage.11,30 The high-conductance chloride channel that we have identified fits the profile of a channel that can be cyclically activated by intracellular metabolic activity. In other systems, this type of channel is gated by NK1 receptors via G-protein activation.25 Rhythmic activation of chloride channels has been observed in secretory epithelial cells where they are activated by calcium probably released by IP3 from the sarcoplasmic reticulum that is fixed near chloride channels through microfilaments.34 An involvement of IP3induced calcium release in activating the chloride channels in ICCs seems likely. We recently showed that blockers of the IP3 receptor and blockers of the sarcoplasmic reticulum calcium pump inhibited slow-wave activity.28 Blockers of IP3 receptors also inhibited the rhythmic whole-cell inward currents of the murine ICCs.30 Hence, an important next step in the unraveling of the pacemaker activity in the gut will be the elucidation of the metabolic changes that rhythmically activate this type of channel. The chloride channel described here may become a target for pharmacologic manipulation of gut motor activity.

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Received December 21, 2001. Accepted August 1, 2002. Address requests for reprints to: Jan D. Huizinga, Ph.D., McMaster University, HSC-3N5C, 1200 Main Street West, Hamilton, Ontario, L8N 3Z5 Canada. e-mail: [email protected]; fax: (905) 522-3454. Supported by the Canadian Institutes of Health Research. The authors appreciated discussing the manuscript with Wayne Giles, Carlos Barajas-Lo ´pez, and David Andrews.