A Ca²⁺-activated Cl⁻ conductance in interstitial cells of Cajal linked to slow wave currents and pacemaker activity

Abstract

Interstitial cells of Cajal (ICC) are unique cells that generate electrical pacemaker activity in gastrointestinal (GI) muscles. Many previous studies have attempted to characterize the conductances responsible for pacemaker current and slow waves in the GI tract, but the precise mechanism of electrical rhythmicity is still debated. We used a new transgenic mouse with a bright green fluorescent protein (copGFP) constitutively expressed in ICC to facilitate study of these cells in mixed cell dispersions. We found that ICC express a specialized ‘slow wave’ current. Reversal of tail current analysis showed this current was due to a Cl⁻ selective conductance. ICC express ANO1, a Ca²⁺-activated Cl⁻ channel. Slow wave currents are not voltage dependent, but a secondary voltage-dependent process underlies activation of these currents. Removal of extracellular Ca²⁺, replacement of Ca²⁺ with Ba²⁺, or extracellular Ni²⁺ (30 μm) blocked the slow wave current. Single Ca²⁺-activated Cl⁻ channels with a unitary conductance of 7.8 pS were resolved in excised patches of ICC. These are similar in conductance to ANO1 channels (8 pS) expressed in HEK293 cells. Slow wave current was blocked in a concentration-dependent manner by niflumic acid (IC₅₀= 4.8 μm). Slow wave currents are associated with transient depolarizations of ICC in current clamp, and these events were blocked by niflumic acid. These findings demonstrate a role for a Ca²⁺-activated Cl⁻ conductance in slow wave current in ICC and are consistent with the idea that ANO1 participates in pacemaker activity.

Introduction

Pacemaker activity in gastrointestinal (GI) muscles is generated by interstitial cells of Cajal (ICC; Langton et al. 1989; Ward et al. 1994; Huizinga et al. 1995; Dickens et al. 1999; see review by Sanders et al. 2006). ICC, identified by anti-Kit antibodies, retain a rhythmic phenotype in cell cultures and generate spontaneous pacemaker activity (Koh et al. 1998; Thomsen et al. 1998). These cells have been used extensively to study the mechanism of rhythmicity in ICC (Tokutomi et al. 1995; Ward et al. 2000; Huizinga et al. 2002; Kim et al. 2002; Koh et al. 2002), and models of pacemaker activity have been proposed from findings of these studies (Kim et al. 2005; Sanders et al. 2006; Faville et al. 2008). A caveat to the conclusions of experiments on cultured ICC is that depolarization of these cells fails to elicit slow waves, but it is well known that slow waves can be paced by application of current in intact muscles (Publicover & Sanders, 1986; Edwards et al. 1999; Kito et al. 2002). Thus, it is possible that an important conductance is lost from the pacemaker mechanism in cultured ICC.

Goto and colleagues (2004) studied freshly isolated ICC from the murine intestine and identified ICC by labelling with anti-Kit antibodies. Large inward currents, termed ‘autonomous’ currents, were activated by depolarization of these cells, and the authors concluded that the autonomous currents were due to a Ca²⁺-dependent, non-selective cation conductance. The ability to activate inward current with current injection is more consistent with what is known about the properties of slow waves in the GI tract. However, the preparation and identification of ICC are difficult, as described in this study, so the nature of the inward current has not been further elucidated.

Microarray analysis to characterize the expression phenotype of ICC in the small intestine was recently performed (Chen et al. 2007). Genes expressed by ICC were compared to expression levels in extracts of whole tissues, and genes highly expressed in ICC (>4 fold) were considered to be potentially important for the unique characteristics of ICC. One highly expressed gene identified by this screen was transmembrane protein 16A (Tmem16a), which encodes anoctamin 1 (ANO1; aka: TAOS2; ORAOV2 and FLJ10261; located on chromosome 7 in the mouse at 7F5). Tmem16a was about eightfold more highly expressed in ICC than in whole tissue extracts. At the time the genomic study was completed, the function of ANO1 was unknown. Gastrointestinal stromal tumours (GISTs) have also been shown to express this gene (referred to as FLJ2061 in these studies), and antibodies to the sequence of the encoded protein (called DOG1 for ‘discovered on GIST-1’) labelled up to 98% of GISTs with Kit mutations (West et al. 2004; Espinosa et al. 2008). Labelling of normal tissues with DOG1 antibody labelled cells with the same anatomical distribution as Kit-positive cells (Espinosa et al. 2008), and a recent study confirmed that ICC express ANO1 in the GI tract (Gomez-Pinilla et al. 2009). ANO1 was recently shown to function as a Ca²⁺-activated Cl⁻ channel (Caputo et al. 2008; Schroeder et al. 2008; Yang et al. 2008), and loss or block of ANO1 blocks electrical slow waves in intact GI muscles (Hwang et al. 2009).

We have recently developed transgenic mice that express a bright form of green fluorescent protein (copGFP) in ICC constitutively. Dispersed ICC can be identified by fluorescence, and voltage-clamp of these cells confirms that large inward currents can be initiated by depolarization (as in Goto et al. 2004). In the present study we explored the properties of these currents in copGFP+ ICC. We found that ICC express ANO1 protein and Cl⁻ channels with a unitary conductance consistent with that of ANO1 channels expressed in HEK293 cells (Yang et al. 2008). The reversal potential of the whole-cell inward current and blockade by low concentrations of niflumic acid are consistent with this current being due to a Cl⁻ conductance. Spontaneous pacemaker activity in single ICC was also blocked by niflumic acid. Our findings support the participation of a novel Ca²⁺-activated Cl⁻ conductance in the generation of slow waves in ICC.

Methods

Animals

A Kit targeting vector was prepared from the PAC library constructed from a female 129S6/SvEvTac mouse spleen genomic DNA (CHORI: Children's Hospital Oakland Research Institute, CA, USA). A copGFP gene amplified from a pFIV-copGFP reporter vector (System Biosciences, Mountain View, CA, USA) by PCR was inserted into the first exon of Kit along with a selection PGK-HPRT cassette (see online Supplemental Material, Supplementary Fig. 1). Targeted knock-in (KI) allele was confirmed by Southern blotting using two external probes. Since copGFP KI inactivates Kit allele, heterozygotes (Kit^+/copGFP mice), 7–12 days of age, were used for studies of ICC. Mice were anaesthetized with isoflurane (Vetone, Meridian, ID, USA) prior to cervical dislocation. The institutional Animal Use and Care Committee at the University of Nevada approved all procedures used.

Preparation of dispersed cells and cell culture

Small strips of intestinal muscle were prepared and equilibrated in Ca²⁺-free Hanks’ solution for 20 min. Cells were dispersed from these strips, as described previously (Koh et al. 1998), with an enzyme solution containing (per ml): collagenase (Worthington Type II, 1.3 mg), bovine serum albumin (Sigma, St Louis, MO, USA, 2 mg), trypsin inhibitor (Sigma, 2 mg) and ATP (0.27 mg). Cells were plated onto sterile glass coverslips coated with murine collagen (2.5 mg ml⁻¹, BD Falcon, Franklin Lakes, NJ, USA) in 35 mm culture dishes. Giga seals were difficult to obtain and maintain in freshly dispersed copGFP+ cells with the dispersion techniques we used. Thus, the cells were allowed to stabilize overnight before experiments in culture medium at 37°C in a 95% O₂–5% CO₂ incubator in smooth muscle growth medium (Clonetics, San Diego, CA, USA) supplemented with 2% antibiotic–antimycotic (Gibco, Grand Island, NY, USA) and stem cell factor (5 ng ml⁻¹, Sigma).

Immunohistochemical identification of copGFP+ cells as ICC

The tunica muscularis was opened along the mesenteric border, pinned to the Sylgard floor of a dissecting dish, and stretched to 110% of the resting length and width. The mucosa was removed by sharp dissection fixing the tissue with paraformaledehyde (4% w/v for 30 min). Tissues were subsequently washed with 0.01 m phosphate buffered saline (PBS, pH 7.4) and incubated in bovine serum albumin (1% for 1 h) to reduce non-specific antibody binding. Intestines were then incubated with an antibody raised against Kit protein (goat anti-SCF (Stem cell factor) receptor/c-kit antiserum; 1 : 500 in PBS, R&D systems Minneapolis, MN, USA) at 4°C overnight, washed in PBS and incubated in Alexa fluor 594-coupled donkey anti-goat secondary antibody (1 : 1000 in PBS; 1 h at room temperature, Invitrogen, Carlsbad, CA, USA). Control tissues were prepared by omitting either the primary or secondary antibodies from the incubation solution.

Double labelling of Kit and ANO1

Double labelling immunohistochemistry was performed on whole mount preparations to determine whether Kit immunopositive ICC express ANO1 in the murine intestine. Whole mounts were prepared as described above but fixed in acetone (10 min, 4°C). Following fixation intestinal tissues were washed in PBS and double labelled with Kit and ANO1. Briefly, tissues were incubated with anti-cKit antibody (ACK2, 5 μg ml⁻¹, Invitrogen) for 24 h, washed in PBS, and then incubated with anti-ANO1 (SP31, 1 : 1000; Abcam, Cambridge, MA, USA) for 24 h. After wash, immunoreactivity was detected via sequential incubation in Alexa fluor 488-coupled goat anti-rabbit and Alexa fluor 594-coupled donkey anti-goat secondary antibodies (Molecular Probes, Eugene, OR, USA; 1 : 500 in PBS; 1 h, room temperature). Control tissues were prepared by omitting either primary or secondary antibodies from the incubation solution.

Cells isolated from the small intestine that were copGFP+ (ICC) were also tested to be sure these specific cells express ANO1. After enzymatic dispersion cells were plated onto glass coverslips and fixed in paraformaldehyde (4% w/v for 10 min). After fixation, the cells were incubated with the anti-ANO1 antibody (1 : 1000) at 4°C overnight, washed in PBS and incubated in Alexa fluor 594-coupled goat anti-rabbit secondary antibody (Invitrogen, Carlsbad, CA, USA; 1 : 1000 in PBS; 1 h at room temperature). Control cells were prepared by omitting either the primary or secondary antibodies from the incubation solution.

After incubation in secondary antibodies, tissues and cells were washed with PBS (overnight at 4°C) and mounted on glass slides using Aqua-Mount (Lerner Laboratories, Pittsburgh, PA, USA). Tissues and cells were examined with a Zeiss LSM 510 Meta confocal microscope (Zeiss, Germany) with an excitation wavelength appropriate for Alexa fluor 488 and Alexa fluor 594. Confocal micrographs are digital composites of Z-series scans of 10–40 optical sections through a depth of 5–40 μm. Final images were constructed using Zeiss LSM 5 Image Examiner software and converted to Tiff files for final processing in Adobe Photoshop 7.0 (Adobe Systems Inc., San Jose, CA, USA) and Corel Draw 12.0 (Corel Corp. Ontario, Canada).

Electrophysiological experiments

Interstitial cells of Cajal (ICC) were identified as green fluorescent protein positive cells under an inverted microscope with attached fluorescence set-up (see Fig. 1). The standard whole-cell patch clamp configuration was also employed to record membrane currents (voltage clamp) and potentials (current clamp, I= 0). For single channel recordings, the excised patch configuration was employed. Currents or potentials were amplified with an Axopatch 200B patch-clamp amplifier (Axon Instruments, Union City, CA, USA) and digitized with a 16-bit analog to digital converter (Digidata 1322A, Axon Instruments) and stored directly on-line using pCLAMP software (version 9.2, Axon Instruments). Data were sampled at 4 kHz and filtered at 2 kHz using an eight-pole Bessel filter for whole-cell experiments and 10 kHz sampling with 2 kHz filtration for single channel experiments. All data were analysed using clampfit (pCLAMP version, 9.2, Axon Instruments) and Graphpad Prism (version 3.0, Graphpad Software Inc., San Diego, CA, USA) software.

Figure 1. Expression of copGFP and ANO1 in ICC.

A, copGFP+ cells in the small intestine of Kit^+/copGFP mice (green, arrows). copGFP fluorescence was primarily located within the perinuclear region of cells, cellular processes were labelled but to a lesser extent. Double labelling with Kit antibody (red, B) confirmed the copGFP+ cells were ICC (C, merged files). D, Kit labelling of ICC in the small intestine. E, labelling of cells with antibody to ANO1. ICC were Kit- and ANO1-immunopositive (F, merged file). Enzymatic dispersions of small intestinal muscles of Kit^+/copGFP mice yielded ICC that were identified by copGFP (green cell; G). These cells were ANO1 immunopositive (red; H). I, a merged file of G and H. J, an insert in panel I, is a copGFP+ cell visualized with DIC. copGFP-negative cells are also present in nearly every field, and these cells were ANO1-negative (arrows). Scale bar in panel F applies to A–F. Scale bar in panel I applies to G–I. Scale bar in panel J is 10 μm.

The pipette tip resistance ranged between 3 and 6 MΩ for whole cell recordings and 5 and 10 MΩ for single channel recordings. Whole-cell and single channel experiments were conducted at 30°C with the use of a CL-100 bath heater (Warner Instruments, Hamden, CT, USA) and room temperature (20°C), respectively.

Solutions for patch clamp experiments

External solution for whole-cell recordings was a Ca²⁺-containing physiological salt solution (CaPSS) containing (mm): 5 KCl, 135 NaCl, 2 CaCl₂, 10 glucose, 1.2 MgCl₂, and 10 Hepes adjusted to pH 7.4 with Tris. In some experiments cells were perfused in Ca²⁺-free solution (omitting 2 mm CaCl₂ from CaPSS). The pipette solutions used for these experiments are described in Table 1. The junction potential was digitally corrected during experiments. For single channel recordings, pipette solution and external solution was high N-methyl-d-glucamine (NMDG) solution containing (mm); 140 NMDGCl, 1 EGTA, 5 Hepes adjusted to pH7.2. Free Ca²⁺ concentrations were calculated by Maxchelator software (http://maxchelator.stanford.edu). The effects of niflumic acid (Sigma) and various concentration of cytosolic Ca²⁺ were tested by a fast bath perfusion system (Automate Scientific, Inc., Berkeley, CA, USA). Within 1 min perfusion, full changes of bath solution were successful.

Table 1.

The composition of pipette solutions

Solutions (mm)	I	II	III	IV
KCl	135	30	0	0
CsCl	0	0	135	30
Potassium aspartate	0	110	0	0
Caesium aspartate	0	0	0	110
MgATP	3	3	3	3
NaGTP	0.1	0.1	0.1	0.1
Creatine phosphate disodium	2.5	2.5	2.5	2.5
EGTA	0.1	0.1	0.1	0.1
Hepes	10	10	10	10
Junction potential (mV)	5.3	14.6	5.3	14.6

All pipette solutions were adjusted pH 7.2 with Tris.

Statistical analysis

Data were tabulated and presented as means ±s.e.m. The n values given represent the number of cells on which specific protocols were performed. Differences between data sets were determined with Student's paired t test and considered significant when P < 0.05.

Results

Expression of copGFP and ANO1 in ICC

Tunica muscularis of small intestine from Kit^+/copGFP mice was labelled with Kit antibodies to verify that copGFP is expressed in ICC. Previous studies have shown that Kit-like immunoreactivity (Kit-LI) is a specific label for interstitial cells of Cajal (ICC) in the murine intestine (Ward et al. 1994). Examination of whole mount preparations revealed that all cells that expressed copGFP also expressed Kit-LI (Fig. 1A–C). Double labelling immunohistochemistry was also performed using anti-Kit and anti-ANO1 antibodies. We found that cells with Kit-LI also expressed ANO1-LI, as others have reported (Fig. 1D–F; Gomez-Pinilla et al. 2009; accompanying paper: Hwang et al. 2009). Dispersed copGFP+ cells (ICC) were also labelled with anti-ANO1 antibodies (Fig. 1G–I). Other cells in the mixed cell populations obtained by enzymatic digestion of intestinal muscles were ANO1-negative (Fig. 1J). Thus, expression of ANO1 protein persisted in copGFP+ cells after enzymatic dispersion.

Isolation of chloride-permeable ionic conductance in ICC

The conventional configuration of the patch clamp technique with cell dialysis was used for whole-cell experiments. External and pipette solutions were CaPSS and pipette solution I, respectively (E_Cl= 0 mV). The average capacitance of single ICC was 5.5 ± 0.5 pF under these conditions. Cells were held at −80 mV and depolarization steps from −80 to +30 mV in 10 mV increments yielded large transient membrane currents (Fig. 2A) averaging −442 ± 60 pA at maximum (−50 mV; n= 4). The currents evoked in ICC had the general characteristics of ‘autonomous currents’ observed in studies of ICC from the murine small intestine previously (Goto et al. 2004); however, as shown below, the conductance responsible for the large amplitude inward currents in ICC displayed different properties than previously reported. The evoked currents were inward at negative potentials and reversed at about −20 mV (Fig. 2B). We also examined the current–voltage (I–V) relationship using pipette solution II (E_Cl=−40 mV). Small net inward currents (average =−23 ± 52 pA; n= 4) were elicited by stepping voltage to −40 mV under these conditions, but outward currents were generated at all potentials positive to −40 mV (Fig. 2C and D). Large amplitude inward tail currents were observed upon returning to −80 mV at either value of E_Cl. These data suggest that a major component of whole-cell currents elicited by depolarization in ICC under physiological ionic gradients is carried by chloride ions. However, K⁺ conductances are likely to have contributed to the whole cell currents using pipette solutions I and II.

Figure 2. Whole cell currents of ICC dialysed with pipette solution I (K⁺-rich solution).

A and C, ICC was depolarized from −80 to +30 mV in 10 mV increments from a holding potential of −80 mV. When E_Cl of internal solution set at 0 mV (pipette solution I), inward currents were evoked at potentials negative to −20 mV (A). Current reversed at approximately −20 mV. Depolarization to −50 mV resulted in activation of inward current, but development of the current response (*) was delayed by 330 ms. Repolarization to −80 mV at the end of the depolarization steps yielded long-lasting tail currents. When E_Cl was adjusted to −40 mV (pipette solution II), little inward current was noted, and outward currents were observed at test potentials to −40 mV (C). B and D, summary current–voltage (I–V) relationships from experiments with pipette solutions I (n= 4) and II (n= 4), respectively.

To eliminate contamination from K⁺ conductances, additional I–V protocols were performed with cells dialysed with pipette solution III (K⁺ concentration of pipette solution I was replaced with equimolar Cs⁺, E_Cl= 0 mV) to inhibit K⁺ currents. Under these conditions, inward currents were elicited at −60 mV, peaked at −50 mV (−1474 ± 382 pA, n= 4, Fig. 3A and B) and reversed near 0 mV (Fig. 3B). When ICC were dialysed with pipette solution IV (E_Cl=−40 mV; Cs⁺ replacement of intracellular K⁺), small inward currents were evoked at −50 mV but outward currents were elicited at potentials positive to −40 mV (Fig. 3C). The evoked currents reversed at about −40 mV (Fig. 3D). To confirm the reversal potential obtained during the I–V plots, additional experiments were performed to analyse the I–V relationship of tail currents.

Figure 3. Whole cell currents in ICC dialysed with pipette solution III (Cs⁺-rich solution).

A and C, ICC was depolarized from −80 to +20 mV in 10 mV increments from a holding potential of −80 mV. When E_Cl was adjusted to 0 mV (pipette solution III) and outward currents were blocked by replacement of intracellular K⁺ with Cs⁺, inward currents were evoked at negative potentials. Current responses reversed at 0 mV (A). Depolarization to −60 mV resulted in activation of inward current, but development of the current response (*) was delayed by 95 ms. When E_Cl was adjusted to −40 mV (pipette solution IV), outward currents were elicited by potentials positive to −40 mV (C). B and D, summary current–voltage (I–V) relationships using pipette solutions III (n= 6) and IV (n= 4), respectively.

Tail currents were analysed by applying a pre-test step from −80 mV to −40 mV to maximally activate the inward current. The evoked current displayed inactivation, so short-duration (100 ms) pre-test steps were used. Instantaneous tail currents were recorded by stepping from −40 mV to potentials ranging from −80 mV to +40 mV (Fig. 4A). With pipette solution III (E_Cl= 0 mV), tail currents reversed at 0 mV (n= 4, Fig. 4B). Outward currents were evoked from a holding potential of −80 mV by a pre-test depolarization to 0 mV when cells were dialysed with pipette solution IV (E_Cl=−40 mV, Fig. 4C). The calculated reversal potential was −39 ± 4 mV with a linear regression plot (n= 4, Fig. 4D). These data suggest that the inward current evoked in ICC by depolarization is due to a Cl⁻ selective conductance.

Figure 4. Reversal of tail currents in ICC.

A and C, reversal potentials were measured by analysis of instantaneous tail currents using CaPSS in the bath and pipette solution III and IV. In A the cell was stepped to −40 mV to activate inward current from a holding potential of −80 mV (pipette solution III, E_Cl= 0 mV). After full activation of inward currents (100 ms), the cell was stepped to various potentials from −80 to +40 mV in 10 mV increments. In C the cell was dialysed with pipette solution IV and stepped to 0 mV to activate outward current from a holding potential at −80 mV (E_Cl=−40 mV). After full activation of outward currents (200 ms), cells were stepped to various potentials from −80 to +40 mV in 10 mV increments. B and D, summaries of tail current I–V relationships from pipette solution III (n= 4) and IV (n= 4), respectively. The measured reversal potentials were 0 mV (B) and −39 mV (D) under these conditions.

Characteristics of chloride-permeable ionic conductance

We further examined the kinetics, and voltage dependence of activation and inactivation of the chloride-permeable inward current using pipette solution III (E_Cl= 0 mV). In experiments like the one shown in Fig. 3A, we noted a very sharp increase in maximal current between −70 and −50 mV. At more positive potentials the amplitude of the current fell linearly with the driving force on Cl⁻. We also noted an odd delay in the activation of currents by steps to −60 mV (see trace denoted by asterisk in Fig. 3A), although peak current amplitude was activated at this potential. Thus, we explored the activation threshold for current and the voltage dependence and kinetics of the current using depolarization step intervals of 5 mV and 2 mV from a holding potential of −80 mV using pipette solution III (Fig. 5A and C).

Figure 5. Threshold and voltage dependence or activation and inactivation of inward currents.

The threshold for activation of inward currents was investigated using 5 mV and 2 mV increments. Cells were bathed in CaPSS, and pipette solution III (E_Cl= 0 mV) was used for these experiments. In A currents were elicited by voltage steps from −80 to +10 mV with 5 mV increments from a holding potential of −80 mV. A summary I–V relationship from 4 cells shows a sharp threshold for activation of current and a linear relationship between current and voltage positive to −65 mV (B). Note the delay, but eventual activation of full magnitude currents, with steps to −65 mV (a) and −60 mV (b). In C current were elicited in the same cell as in A using 2 mV increments between steps from −80 to −60 mV to measure times to peak current of each potential tested. Note the delays, but eventual activation of currents of full magnitude, with steps to −72 mV (a), −70 mV (b) and −68 mV (c). D, summary of the I–V relationship for 4 cells. Again, there was a sharp threshold at −72 mV and a linear relationship between current and voltage positive to −72 mV. Times-to-peak current responses are plotted in E from an experiment using 2 mV increments. The time-to-peak current slows at negative test potentials, suggesting a voltage-dependent mechanism may underlie activation of the inward currents. Voltage vs. activation and inactivation are plotted in F (data analysed from currents in C). Activation (normalized to maximum conductance, G/G_max) and inactivation (normalized to maximum current, I/I_max) could not be fitted with Boltzmann equations suggesting the inward currents are not voltage dependent.

Using 5 mV step increments, the threshold potential for activation of the inward current was −70 mV (Fig. 5B). This was refined to −72 mV using 2 mV increments (Fig. 5D). The time to peak current (measured as the time from the application of the depolarization step to the peak of the evoked inward current) was dramatically dependent upon voltage over a very narrow range of potentials. For example, with 2 mV steps, the time to peak current was 651 ms at −72 mV and 90 ms at −64 mV (Fig. 5E). We also noted that the current amplitude was not affected by voltage other than linear dependence upon the driving force for Cl⁻. Once the conductance was activated, currents of maximal amplitude developed (and the maximal current amplitude was only affected by the change in driving force for Cl⁻ ions; Fig. 5B). The same sharp activation threshold was noted in cells with low current density.

We also analysed the voltage dependence of activation and inactivation of chloride-permeable inward currents using 2 mV voltage-step increments. The activation and inactivation properties were poorly fitted with Boltzmann equations, suggesting that the Cl⁻-permeable conductance responsible for the inward current activated by depolarization is not itself voltage dependent (Fig. 5F). A secondary voltage-dependent process (e.g. Ca²⁺ entry or Ca²⁺-induced Ca²⁺ release) may be responsible for activating the inward current (see Fig. 5C).

Ca²⁺ dependence of chloride-permeable conductance and channels

We next examined the Ca²⁺ dependence of the chloride-permeable currents in ICC. For these experiments we used Ca²⁺-free solution (i.e. Ca²⁺ removed from the external buffer). The chloride-permeable inward currents were completely abolished by Ca²⁺-free solution (Fig. 6A and B) and reversed upon restoration of [Ca²⁺]_o in the CaPSS (see protocol Fig. 6B inset). We also examined the effects of replacing [Ca²⁺]_o with Ba²⁺ (i.e. BaPSS in which 2 mm Ca²⁺ was replaced with equimolar Ba²⁺) on chloride-permeable inward currents. BaPSS also inhibited inward currents (Fig. 6C and D), and the currents were restored by restoration of [Ca²⁺]_o. These data suggest that the chloride-permeable currents are Ca²⁺ dependent or may depend upon the entry of Ca²⁺ for initiation of the current. We also tested of Ni²⁺, a blocker of low-threshold voltage-dependent Ca²⁺ channels. The Cl⁻ current was blocked by Ni²⁺ (30 μm; n= 5; Fig. 6E and F).

Figure 6. Effects of external Ca²⁺ on inward currents in ICC.

In A peak currents were recorded repetitively by stepping cells from −100 mV to −60 mV every 30 s (pipette solution III; E_Cl= 0 mV). External CaPSS was replaced with Ca²⁺-free (0-Ca²⁺) solution (black bar). Reduction in external Ca²⁺ caused inhibition of inward current, and this effect was reversible upon restoration of CaPSS. B, representative traces (a–d) before, during and after removal of Ca²⁺ at time points indicated in A. C, peak currents were recorded by stepping repetitively from −80 mV to −50 mV every 30 s. External Ca²⁺ (CaPSS) was replaced with equimolar Ba²⁺ (2 mm; BaPSS; black bar). Peak currents were inhibited by BaPSS, and this effect was reversible when CaPSS was restored. Representative traces from the time points indicated in C are shown in D (a–d). E, peak currents were recorded by stepping repetitively from −80 mV to −35 mV every 30 s. Ni²⁺ (30 μm, black bar) added to the CaPSS inhibited the inward current. The effects of Ni²⁺ were slow to wash out. Representative traces from the time points indicated in E are shown in F (a–d).

We performed experiments using excised patches from ICC and symmetrical gradients of NMDGCl (140/140 mm) to determine whether Ca²⁺-dependent Cl⁻ channels are expressed by ICC (as immunohistochemical studies implied). Under the conditions of these experiments only Cl⁻ channels could be detected. Channel openings with a single channel conductance of 7.8 ± 0.2 pS (n= 4) were observed, and currents reversed at 0 mV (Fig. 7A and B). Ramp depolarization of patches (i.e. −80 mV to +80 mV) elicited currents that were greatly increased in amplitude when the cytoplasmic surface of the patches was exposed to high Ca²⁺ (10⁻⁶m) vs. low Ca²⁺ (10⁻⁸m). The low concentration of Ca²⁺ (10⁻⁸m) decreased the open probability of 7.8 pS channels (Fig. 7C and D). Reversal potential (0 mV) was unchanged by differences in [Ca²⁺]_i. These experiments demonstrate that the 7.8 pS Cl⁻ channels in ICC are activated by Ca²⁺. These channels are potentially responsible for the Ca²⁺-activated Cl⁻ currents identified in whole-cell experiments.

Figure 7. Single channel currents in ICC.

Patches were excised from ICC to study single channel conductances that might contribute to inward currents. A, the external and internal solutions were symmetrical NMDGCl (140/140 mm). Under these conditions, only Cl⁻ could be recorded. The calculated free Ca²⁺ concentration was 10⁻⁶m. The representative traces at various holding potentials show channel openings (dotted lines to denote unitary current levels; the 0 current level is denoted by the continuous line in each trace). An all-points histogram describing currents at −40 mV is shown as an inset in B. The single channel currents reversed at 0 mV (continuous line). B, a plot of the average current–voltage relationships from 5 patches (error bars contained within symbols). The calculated single channel conductance by linear regression was 7.8 pS. C, cytoplasmic [Ca²⁺] was changed from 10⁻⁶ to 10⁻⁸m. The low [Ca²⁺] reduced or abolished channel openings. Ramp protocol from −80 mV to +80 mV from a holding potential of −80 mV were applied to the patches. Low [Ca²⁺] (10⁻⁸m) completely abolished currents. These experiments were repeatable. The 0 current level is denoted by the dotted line. D, representative current traces elicited by ramp protocols applied at the points noted in C ([Ca²⁺]= 10⁻⁶m, a; intermediate value during exchange, b; and Ca²⁺= 10⁻⁸m, c).

Niflumic acid sensitivity of Ca²⁺-activated chloride currents and its functional role on pacemaker potentials

We tested the effect of niflumic acid on inward current elicited in ICC by depolarization. Using pipette solution III, cells were stepped to −35 mV every 30 s from a holding potential of −80 mV. The peak current was plotted as a function of time. Niflumic acid (10 μm) decreased the inward current from −761 ± 75 pA to −162 ± 33 pA (n= 5; P < 0.01, Fig. 8A and B). Various concentrations of niflumic acid (1–100 μm) were also tested to determine the concentration–response relationship for block of the inward current. An IC₅₀ of 4.8 μm (Hill slope = 0.34) was calculated from these experiments. It should be noted that although niflumic acid reduced the amplitude of the inward current, the kinetics of deactivation of the current were slowed by this compound (Fig. 8B).

Figure 8. Inhibition of inward currents by niflumic acid.

A, currents were elicited by stepping from −80 to −35 mV every 30 s. Peak currents are plotted as a function of time. Niflumic acid (NFA, 10 μm) decreased the amplitude of the inward currents. Representative traces taken at the time points noted in A are shown in B (a–d). Note the slowing of the inactivation kinetics in response to NFA. The concentration-response relationship for the inhibitory effects of niflumic acid (1–100 μm) are shown in C and fit with a sigmoidal function (Hill equation). The calculated IC₅₀ was 5 μm (Hill slope = 1.38, n= 5). Representative currents elicited by voltage steps from −80 to 0 mV show the effects of 1 μm (a), 5 μm (b), 10 μm (c), 30 μm (d) and 100 μm (e) niflumic acid in one experiment (Panel D).

Current clamp experiments were also performed. We have previously shown that ICC discharge small amplitude, unitary potentials in current clamp conditions (Kim et al. 2002). Thus, a spontaneous pacemaker mechanism exists in ICC that can depolarize cells to the threshold for the generation of the large inward current measured under voltage-clamp. Switching to current clamp (I= 0), we observed large-amplitude, spontaneous oscillations in membrane potential in ICC dialysed with pipette solution I (Fig. 9). The resting membrane potentials (RMP) of cells (maximum potential between voltage oscillations) averaged −51 ± 3 mV (n= 6). The amplitude of the spontaneous oscillatory membrane potentials was 36 ± 2 mV and these events occurred at a frequency of 11.8 ± 1.4 cycles min⁻¹. The half-duration of the oscillations was (1478 ± 477 ms) and maximal upstroke velocity was (0.42 ± 0.17 mV ms⁻¹; measured from 10% of RMP to 90% of peak amplitude). Niflumic acid (5 μm) did not significantly affect RMP (−47 ± 2 mV) and the amplitude of spontaneous oscillatory membrane potentials (32 ± 2 mV), but increased the half-duration of the transient depolarizations to 2029 ± 489 ms (P < 0.05) and decreased maximal rising slope (0.21 ± 0.10 mV ms⁻¹, P < 0.05) and frequency (6.67 ± 0.61 cycles min⁻¹, P < 0.01) significantly (n= 6). The increase in half-duration of the membrane potential oscillations is consistent with the observation that niflumic acid slowed the kinetics of the deactivation of the inward current. Higher concentrations of niflumic acid (>10 μm) blocked the large amplitude spontaneous membrane potential oscillations, but did not block the underlying small unitary potential activity. These data suggest that Ca²⁺-activated Cl⁻ currents are involved in spontaneous slow waves in ICC.

Figure 9. Effects of niflumic acid on membrane potential in ICC under current clamp.

Spontaneous membrane potential oscillations were recorded in single ICC under current clamp (I= 0). CaPSS is the external solution and pipette solution I was used for the data recorded in A. Application of niflumic acid (NFA, 5 μm) decreased the frequency of spontaneous oscillations without affecting resting membrane potentials (most negative potential between oscillations). When regular oscillations were inhibited by niflumic acid, smaller amplitude noisy transient depolarization potentials were still present. The inhibitory effects of niflumic acid were reversible upon washout. B, data during the periods denoted in A by the boxes before and during NFA are shown at higher sweep speed. C–F, the effects of niflumic acid on amplitude (C), half-width (D), maximum slope (E) and frequency (F) from 6 cells. *P < 0.05, **P < 0.01.

Discussion

In the present study we measured large amplitude inward currents in ICC, identified by constitutive expression of copGFP, with some characteristics similar to the ‘autonomous’ currents previously reported by Goto and coworkers (2004). The inward currents in ICC were activated by depolarization, but the conductance underlying these currents does not appear to be intrinsically voltage dependent, as a very sharp threshold voltage was detected for activation and inactivation, and, once exceeded, full activation and inactivation processes occurred. Furthermore, the activation and inactivation properties of the inward current were poorly fit by Boltzmann functions. It appears that another voltage-dependent process underlies depolarization-induced initiation of the large inward current in ICC. At potentials close to the threshold for activation, we also noted dramatic and atypical time-dependent characteristics in which the activation of the current was delayed for periods of time that decreased as a function of voltage over a narrow range of potentials near threshold. When whole cell currents were free from contamination by K⁺ currents, the reversal potential for the current closely followed E_Cl. Reversal of tail currents demonstrated that the large inward currents in ICC were due to activation of a Cl⁻ conductance, in contrast to the conclusions of Goto and coworkers (2004). We found that the inward current was blocked by low concentrations of niflumic acid, and this compound blocked the spontaneous, slow wave-like depolarizations in ICC under conditions of current clamp. Slow waves were also blocked by niflumic acid in intact GI muscles (Hwang et al. 2009); however the block of spontaneous depolarizations in single ICC occurred at lower concentrations than block of slow waves in intact muscles. In single channel experiments we found Ca²⁺-activated Cl⁻ channels with a unitary conductance of 7.8 pS in excised membranes of ICC. This conductance is equivalent to the single channel conductance of ANO1 channels expressed in HEK cells (i.e. 8 pS; Yang et al. 2008). Ca²⁺-activated Cl⁻ channels of this magnitude have also been recorded in endothelial cells (Nilius et al. 1997) and hepatocytes (Koumi et al. 1994); however the molecular identity of these channels has not yet be determined.

Discovery and initial characterization of an inward current activated by depolarization in freshly dispersed Kit-positive cells of the murine small intestine was an important contribution to understanding the pacemaker activity of ICC (Goto et al. 2004). The currents recorded previously (Goto et al. 2004) were termed ‘autonomous’ because once activated, the current persists in spite of repolarization, as evidenced by persistence of long-duration tail currents upon repolarization (as in Fig. 4). The inward current we observed in ICC is not ‘autonomous’ in the strict sense of the word, and its kinetics are more likely a function of the dynamics of intracellular Ca²⁺. The contribution of this conductance to slow waves in ICC and in intact muscles (Hwang et al. 2009) suggests that ‘slow wave current’ is a better descriptive term for this current. The channels responsible for and/or the mechanism of voltage-dependent activation of inward current are lost in cultured cells within a few days (as are other features fundamental to the ICC phenotype, such as Kit expression; K. M. Sanders and S. Ro, unpublished observation). We failed to resolve this conductance in previous characterizations of pacemaker activity in cultured ICC (e.g. Koh et al. 2002). Thus, models that we, and others, have proposed in the past to describe pacemaker activity, which are based on periodic activation of non-selective cation currents, are incomplete (e.g. Sanders et al. 2006; Youm et al. 2006; Faville et al. 2008; Faville et al. 2009). Non-selective cation conductances may still be responsible for the spontaneous transient inward currents that underlie unitary potentials in ICC, because Cl⁻ channel blocking drugs did not inhibit these events in intact muscles (Hwang et al. 2009) and in isolated ICC (this study). Constitutive labelling of ICC by expression of copGFP greatly facilitates identification of ICC in mixed cell populations. This animal model will enable future studies of excitability and the pacemaker mechanism in ICC.

The current study identified several important new properties of the large inward current in ICC. Of primary importance, the conductance responsible for depolarization-activated inward currents is Cl⁻ selective, as determined by the reversal of currents and tail currents at E_Cl. The properties of the current suggested that the Cl⁻ conductance itself is not voltage dependent, but rather it is activated by an underlying voltage-dependent process. Threshold for activation of the inward current under voltage clamp was quite negative (approximately −70 mV), and near-threshold depolarization resulted in delayed activation of full-amplitude currents. Significant delays between application of a stimulus and activation of a slow wave (termed by the authors –‘regenerative potentials’) have also been observed in response to depolarizations of small muscle bundles that slightly exceeded threshold (Hirst et al. 2002). The latency of activation decreased as a function of the intensity of the depolarizing current. Although latencies were longer in intact muscles than in isolated cells, in both cases the latencies between depolarization and activation exceeded the normal time constants for voltage-dependent activation of ion channels (millisecond range) by orders of magnitude. Causes for this latency have been suggested to include activation of an underlying voltage-dependent current (possibly a Ca²⁺ conductance; Kim et al. 2002) that might result in (i) entry of Ca²⁺ and Ca²⁺-induced release of Ca²⁺ from intracellular stores or (ii) voltage-dependent activation of phospholipase C, increased IP₃ production and release of Ca²⁺ from intracellular stores (Ganitkevich & Isenberg, 1993; Hirst et al. 2002). Either process might be capable of entraining Ca²⁺ release and activation of a Ca²⁺-activated Cl⁻ conductance. Our data favour the former mechanism of activation, however, because low concentrations of Ni²⁺ also blocked depolarization-induced activation of inward current.

The large inward current activated by depolarization in ICC appears to be Ca²⁺ dependent. We found that the current was reduced within 1 min by removal of extracellular Ca²⁺ or its replacement with Ba²⁺. Thus, entry of Ca²⁺ and possibly recruitment of Ca²⁺ release from intracellular stores might be the signal that activates the large inward current. Previous studies have suggested that voltage-dependent activation of low-threshold Ca²⁺ channels is required for entraining slow waves (Kim et al. 2002). In single channel studies in excised patches, we resolved Ca²⁺-dependent Cl⁻ channels in ICC, and these channels had the same unitary conductance (7.8 pS) as ANO1 channels expressed in HEK cells (Yang et al. 2008). Genomic and immunohistochemical evidence of high levels of ANO1 expression in ICC and loss of slow waves in muscles of mice with inactivated Tmem16a (Hwang et al. 2009) support the idea that this class of Ca²⁺-activated Cl⁻ channels is responsible for the Ca²⁺-dependent inward currents in ICC. Experiments on ICC from Tmem16a^−/− animals would tighten this correlation further, but Tmem16a^−/− mice, which have limited post-natal viability, with copGFP-labelled ICC have not yet been produced.

By superficial comparison, the time course and kinetics of inward current in ICC and the current–voltage relationship for this current do not closely resemble Ca²⁺-activated Cl⁻ currents carried by ANO1 expressed in HEK293 cells (see Hartzell et al. 2009). However, the differences in characteristics are likely to be due to the dynamics of cytoplasmic Ca²⁺ concentration ([Ca²⁺]_c), which may change in ICC during the initiation and time course of the inward current. Subcompartments in ICC may also influence the [Ca²⁺] available to interact with channels. Pure ANO1 currents are outwardly rectifying and carry little inward current at submaximal [Ca²⁺]_c, but as [Ca²⁺]_c rises the current–voltage relationship becomes linear (Schroeder et al. 2008). The conductance responsible for inward current in ICC carries significant inward current at negative potentials and is relatively linear (see Figs 4 and 5). Thus, activation of inward current may be due to entry and/or liberation of Ca²⁺ in ICC, raising [Ca²⁺] near the plasma membrane to levels that facilitate inward current (i.e. >1 μm). The time course and kinetics of the inward current in ICC are also likely to be influenced by the dynamics of [Ca²⁺]. In experiments on expressed ANO1 in axolotl oocytes, brief laser pulses (250 ms) to uncage IP₃ resulted in currents and membrane potential changes lasting for several seconds (Schroeder et al. 2008). The kinetics of these currents presumably followed the time course of [Ca²⁺]_c that was elevated in oocytes for a period that far exceeded the duration of the stimulus. Thus, ANO1 currents can be initiated by brief stimuli and then follow the time course of intracellular Ca²⁺ dynamics and possibly other regulatory mechanisms. This is analogous to the inward current we measured in ICC, which can be initiated by depolarization, but once threshold is achieved, intracellular regulatory mechanisms (presumably changes in [Ca²⁺]) determine the magnitude and time course of the current.

Previous studies have reported a variety of conductances in ICC, including (i) voltage-dependent Ca²⁺ channels (Kim et al. 2002); (ii) ether-à-go-go-related K⁺ channels (Zhu et al. 2003; McKay et al. 2006); (iii) ATP-sensitive K⁺ channels (Choi et al. 2006), (iv) Ca²⁺-activated K⁺ channels (Fujita et al. 2001; Zhu & Huizinga, 2008); (v) delayed rectifier (K_V1.1) channels (Hatton et al. 2001); and (vi) inwardly rectifying chloride channels (Zhu et al. 2005). From these studies a variety of mechanisms have been proposed to explain the spontaneous voltage transients in GI muscles, known as slow waves, but no previous studies have described the Ca²⁺-activated Cl⁻ conductance activated by depolarization that we observed in individually indentified ICC. Others have proposed a role for the Cl⁻ conductances in ICC pacemaking (Huizinga et al. 2002; Zhu et al. 2005); however the conductances described were not Ca²⁺ sensitive and no mechanism was proposed for spontaneous and periodic activation of these conductances as must occur in slow wave activity. It must be recognized that in previous studies each cell tested was not definitively identified as an ICC. Instead, cells were selected because they displayed morphology similar to the cells labelled by Kit antibody (e.g. Koh et al. 2002). There are numerous cell phenotypes present in dispersions of GI muscle tissues, and it is abundantly clear that the phenotypes and morphologies of cells change dramatically in cell cultures and explants. Compounding this problem, ICC, smooth muscle cells and fibroblast-like cells form gap junctions with each other, and therefore, unless a cell is physically isolated from others, it is not possible to be certain that conductances measured are the exclusive properties of the cell with which the giga seal is formed. In cell dispersions from the mouse intestine, cells with copGFP fluorescence were small (5 pF cell capacitance) and round, some displayed short processes (see Fig. 1). Enzymatic dispersion may break off or limit the extent of cell processes. These cells were clearly isolated from other cells as demonstrated by the low capacitance and cell access. Our data and initial observations from studies of ICC that are constitutively labelled suggest that many of the published properties of ICC need to be carefully re-examined in light of the problems of identification of ICC and phenotypic changes that develop in various culture conditions (e.g. single cell cultures, explants, and influence of substrate).

Glossary

Abbreviations

ANO1

anoctamin 1

BaPSS

2 mm Ca²⁺ in CaPSS replaced with equimolar Ba²⁺

CaPSS

Ca²⁺-containing physiological salt solution

E_Cl

equilibrium potential of chloride

GFP

green fluorescent protein

GI

gastrointestinal

GISTs

gastrointestinal stromal tumours

ICC

interstitial cells of Cajal

KI

knock-in

Kit-LI

Kit-like immunoreactivity

NMDG

N-methyl-d-glucamine

RMP

resting membrane potentials

Tmem16a

transmembrane protein 16A

Author contributions

M.H.Z., T.W.K. and S.D.K. performed patch-clamp experiments. S.R. and W.Y. generated the copGFP knock-in mice used to isolate ICC. S.M.W. performed immunohistochemistry to verify that copGFP-expressing cells were ICC. S.D.K. and K.M.S. planned experiments, analysed data and wrote the manuscript.