1The role of the cytoskeleton in leptin-induced activation of ATP-sensitive K+ (KATP) channels was examined in rat CRI-G1 insulin-secreting cells using patch clamp and fluorescence imaging techniques.
2In whole cell recordings, dialysis with the actin filament stabiliser phalloidin (10 μm) prevented KATP channel activation by leptin.
3Application of the actin filament destabilising agents deoxyribonuclease type 1 (DNase 1; 50 μg ml−1) or cytochalasin B (10 μm) to intact cells or inside-out membrane patches also increased KATP channel activity in a phalloidin-dependent manner.
4The anti-microtubule agents nocodazole (10 μm) and colchicine (100 μm) had no effect on KATP channel activity.
5Fluorescence staining of the cells with rhodamine-conjugated phalloidin revealed rapid disassembly of actin filaments by cytochalasin B and leptin, the latter action being prevented by the phosphoinositide 3 (PI 3)-kinase inhibitor LY 294002.
6Activation of KATP channels by the PI 3-kinase product phosphatidylinositol 3,4,5-trisphosphate (PtdIns(3,4,5)P3) was also prevented by phalloidin. This is consistent with the notion that leptin activates KATP channels in these cells by an increase in PtdIns(3,4,5)P3 or a similar 3-phosphorylated phosphoinositol lipid, resulting in actin filament disruption.
It is well established that many ion channels and transporters are anchored in the membrane by either direct or indirect association with the cytoskeleton. In addition there is growing evidence that altering the integrity of cytoskeletal elements, in particular actin filaments, can modulate the activity of a variety of ion channels (Janmey, 1998) and receptors (Wang et al. 1999). For example, disruption of actin filaments with cytochalasin results in activation of ATP-sensitive K+ (KATP) channels in cardiac myocytes (Terzic & Kurachi, 1996).
In pancreatic β-cells, KATP channels play a central role in the control of insulin secretion (Ashcroft & Rorsman, 1991), such that closure of these channels results in depolarisation of the cell membrane, activation of voltage-dependent calcium channels and ultimately release of insulin (Ashford, 1990). Recent evidence indicates that the ob gene product leptin activates KATP channels in pancreatic β-cells (Kieffer et al. 1997) and rat CRI-G1 insulinoma cells (Harvey et al. 1997), an action consistent with suppression of insulin secretion. The leptin receptor is a member of the class I cytokine receptor superfamily (Tartaglia et al. 1995), which signal via association with janus tyrosine kinases. Activated janus kinases can signal via insulin receptor substrate proteins (Ihle, 1995), which once phosphorylated can bind to Src-homology 2-containing enzymes like phosphoinositide 3-kinase (PI 3-kinase). Indeed, leptin stimulates glucose transport in C2C12 myotubules via a PI 3-kinase-dependent process (Berti et al. 1997). Recently we have also shown that PI 3-kinase activation is required for leptin activation of KATP channels in CRI-G1 insulinoma cells (Harvey & Ashford, 1998; Harvey et al. 2000). Since one of the primary functions of PI 3-kinase is to phosphorylate PtdIns(4,5)P2 into PtdIns(3,4,5)P3 (Shepherd et al. 1998) and PtdIns(3,4,5)P3 mimics the actions of leptin (Harvey et al. 2000), PtdIns(3,4,5)P3 or a similar 3-phosphorylated phophoinositol lipid may underlie leptin signalling to KATP channels in CRI-G1 cells.
A number of lipid kinases, including PI 3-kinase, are localised to the cytoskeleton and their activities are modulated by a variety of cytoskeletal proteins, especially those associated with actin (Janmey, 1998). Furthermore, some receptor-mediated signal transduction pathways, such as insulin-like growth factor (IGF) 1 (Casamassina & Rozengurt, 1998) and interleukin (IL) 2 (Gesbert et al. 1998) require association of, and alteration in, the activity of PI 3-kinase and cytoskeletal elements. Consequently, we have examined the possibility that activation of KATP channels in CRI-G1 insulin-secreting cells by a leptin-induced increase in PI 3-kinase activity is also dependent upon changes in the organisation of the cytoskeleton.
Cells from the rat insulin-secreting cell line CRI-G1 were grown in Dulbecco's modified Eagle's medium with sodium pyruvate and glucose (Life Technologies), supplemented with 10 % fetal calf serum (Sigma) and 1 % penicillin-streptomycin at 37°C in a humidified atmosphere of 95 % air and 5 % CO2. Cells were passaged every 2–5 days as described previously (Carrington et al. 1986), plated onto 3.5 cm Petri dishes (Falcon 3001) and used 1–4 days after plating.
Electrophysiological recording and analysis
Experiments were performed using whole cell current clamp recordings to monitor membrane potential with excursions to voltage clamp mode to examine macroscopic currents and cell-attached and inside-out configurations to examine single channel responses, as described previously (Harvey et al. 1997). During voltage clamp recordings, the membrane potential was clamped at −50 mV and 10 mV voltage steps of 100 ms duration were applied every 200 ms in the voltage range −120 to −30 mV. Current and voltage were measured using the Axopatch 200B (Axon Instruments) and List EPC-7 amplifiers. Currents evoked in response to the voltage step protocol were analysed using pCLAMP 6.0 software (Axon Instruments) whereas current clamp data were recorded onto digital audiotapes and replayed for illustration on a Gould TA 240 chart recorder. Single channel data were analysed for current amplitude and channel activity (NfPo; where Nf is the number of functional channels and Po is the open probability) as described previously (Lee et al. 1995). All data are expressed as means ±s.e.m. and statistical analyses were performed using Student's unpaired t test (unless otherwise stated). P < 0.05 was considered significant.
Recording electrodes were pulled from borosilicate glass and had resistances of 1–5 MΩ for whole cell recordings and 8–12 MΩ for single channel recordings when filled with electrolyte solution. The pipette solution for whole cell recordings comprised (mm): 140 KCl, 0.6 MgCl2, 2.73 CaCl2, 5.0 Mg-ATP, 10 EGTA, 10 Hepes (pH 7.2; free [Ca2+] of 100 nm), whereas for single channel recordings the pipette solution contained (mm): 140 KCl, 1 CaCl2, 1 MgCl2, 10 Hepes (pH 7.2). For whole cell and cell-attached recordings the bath solution was normal saline (mm): 135 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 10 Hepes (pH 7.4), whereas for inside-out membrane patches the solution comprised (mm): 140 KCl, 1 MgCl2, 2 CaCl2, 10 EGTA, 10 Hepes (pH 7.2; free [Ca2+] of 30 nm). The free Ca2+ concentration was calculated using the ‘METLIG’ program (P. England & R. Denton, University of Bristol, UK). All solution changes were achieved by superfusing the bath using a gravity feed system at a rate of 10 ml min−1, which allowed complete bath exchange within 2 min. All experiments were performed at room temperature (22–25°C).
Initially cells were incubated with either leptin (10 nm for 40 min) or the cytoskeleton disrupter cytochalasin B (10 μm for 30 min) prior to washing the cells rapidly in cold saline. Cells were then fixed and permeabilised simultaneously by immersion in 100 % methanol at −20°C for 5 min followed by two washes in normal saline (maintained at room temperature). For staining of the actin cytoskeleton, cells were incubated with rhodamine-conjugated phalloidin (50 nm in saline) for 30 min and then rinsed with saline before the labelling was visualised using a confocal imaging system (Bio-Rad Microradiance). Rhodamine-conjugated phalloidin was excited using a 542 nm line, whereas fluorescent emission was detected above 570 nm. In control experiments, rhodamine-conjugated phalloidin binding was completely inhibited by pre-treatment with phalloidin (10 μm).
Analysis of imaging
The intensity of rhodamine-conjugated phalloidin staining in the plasma membrane was determined using Bio-Rad Lasersharp software. Analysis lines were drawn along randomly selected regions of the plasma membrane and the mean fluorescence intensity calculated. Data were obtained from five cells selected at random for each condition. Within a given experimental series all conditions for capturing images, including illumination intensity and photomultiplier gains, were constant. In order to allow for quantification of experimental data obtained on separate days, the data were normalised relative to the mean plasma membrane fluorescence measured in the control cells for each day. Comparisons between the normalised experimental data sets were performed using Student's unpaired t test.
Phalloidin, paclitaxel (taxol), deoxyribonuclease type 1 (DNase 1), cytochalasin B, nocodazole, colchicine, tolbutamide and Mg-ATP were obtained from Sigma. Human recombinant leptin was obtained from Calbiochem. Leptin was prepared as a stock solution in normal saline and further diluted in normal saline containing 0.2 % bovine serum albumin as a carrier. Tolbutamide was made up as a 100 mm stock solution in DMSO, whereas ATP was stored as a 100 mm solution in 10 mm Hepes (pH 7.2) and kept at −4°C until required. Phalloidin was made up as a 1 mm stock solution in DMSO, whereas cytochalasin B and nocodazole were stored as 10 mm solutions in DMSO. Taxol was made up as a 2.5 mm solution in 1 % ethanol and DNase 1 was stored as a 10 μg ml−1 solution in normal saline. Colchicine was stored as a 10 mm solution in 1 % ethanol. Rhodamine-conjugated phalloidin (obtained from Molecular Probes) was stored as a 6.6 mm stock solution in 1 % methanol.
Phalloidin blocks leptin activation of KATP channels
Current and voltage clamp recordings from CRI-G1 insulin-secreting cells, with 5 mm Mg-ATP present in the electrode solution to maintain KATP channels in the closed state, resulted in a mean resting membrane potential of −38.0 ± 3.8 mV and a slope conductance of 0.40 ± 0.06 nS (n= 4). Application of the ob gene product leptin (10 nm) hyperpolarised CRI-G1 cells to −70.0 ± 1.5 mV and increased the slope conductance to 2.09 ± 0.70 nS (n= 4; Fig. 1A and B). The reversal potential (obtained from the point of intersection of the current-voltage relationships) associated with the leptin-induced increase in conductance was −79.0 ± 2.1 mV (n= 4), which is close to the calculated value for the K+ equilibrium potential (EK) of −84 mV under these recording conditions, indicating an increased K+ conductance. Application of the sulphonylurea tolbutamide (100 μm) completely reversed the leptin-induced hyperpolarisation and increase in conductance to −37.6 ± 2.9 mV and 0.46 ± 0.05 nS (n= 4), respectively. These data show that leptin activates KATP channels and are in agreement with previous studies using pancreatic β-cells (Keiffer et al. 1997) and this cell line (Harvey et al. 1997; Harvey & Ashford, 1998).
In order to explore whether leptin-induced activation of KATP channels in CRI-G1 cells involves alterations in the structural state of the actin cytoskeleton, the effects of the heptapeptide mushroom toxin phalloidin (Weiland & Faulstich, 1978) on the responses to leptin were examined. Phalloidin binds to filamentous F-actin with high affinity and, by lowering the critical concentration for polymerisation, shifts the equilibrium between F-actin and the monomeric G-actin towards the polymerised form (Cooper, 1987). As phalloidin is membrane-impermeant, cells were dialysed with an electrode solution containing 5 mm Mg-ATP and 10 μm phalloidin. Under these conditions and following dialysis for at least 10 min to allow the slow-diffusing phalloidin access to the cell interior, the mean resting membrane potential and slope conductance of CRI-G1 cells were −33.0 ± 1.6 mV and 0.90 ± 0.08 nS, respectively (n= 4). Subsequent application of leptin (10 nm) after this period of dialysis failed to hyperpolarise CRI-G1 cells (n= 4; Fig. 1C). The corresponding membrane potential and slope conductance values obtained 20 min after the addition of leptin were −35.0 ± 2.4 mV and 0.83 ± 0.07 nS, respectively (n= 4; P > 0.05; Fig. 1D). Thus, stabilisation of actin filaments in the polymerised form prevented the ability of leptin to activate KATP channels. This indicated that the likely mechanism by which leptin activates KATP channels in this cell line is by disruption of the filamentous structure of the actin cytoskeleton.
Actin filament structure is controlled by reversible polymerisation of G-actin which forms F-actin, and this process is under the dynamic control of various actin binding proteins (Pollard et al. 1994). This actin-based structure can be disrupted by specific reagents such as deoxyribonuclease type 1 (DNase 1) and the cytochalasins (Cooper, 1987; Pollard et al. 1994). It has previously been reported that these microfilament disrupters enhance the activity of cardiac muscle KATP channels when applied directly to the cytoplasmic domain of isolated membrane patches, indicating that these channels are functionally coupled to the actin cytoskeleton in myocytes (Terzic & Kurachi, 1996). Consequently, in order to examine whether such a linkage occurs in pancreatic β-cells and to make a comparison with the effects of leptin described above, we have determined the actions of these actin disrupters on the activity of KATP channels in CRI G1 insulin-secreting cells. Initially we examined the actions of DNase 1, which is known to form complexes with G-actin and prevent microfilament polymerisation (Kabsch et al. 1990). In a separate series of control experiments, current and voltage clamp recordings were obtained from CRI-G1 cells, with 5 mm Mg-ATP in the pipette solution. The mean resting membrane potential and slope conductance were −33.0 ± 1.8 mV and 0.50 ± 0.03 nS (n= 6), respectively, and these values were stable for at least 30 min of recording. In contrast, dialysis of cells with a pipette solution containing 5 mm Mg-ATP and DNase 1 (50 μg ml−1) caused, following a delay of 7–12 min, a slow hyperpolarisation, associated with an increased cellular conductance (Fig. 2A) which was maintained for the duration of the recordings (up to 45 min). The membrane potential increased, from a mean value (obtained within 1–2 min of formation of the whole cell configuration) of −32.3 ± 2.1 mV to −67.4 ± 2.9 mV (after 15–18 min), an action associated with a concomitant change in the slope conductance from an initial value of 0.53 ± 0.02 nS to 3.12 ± 0.58 nS (n= 5; Fig. 2B). The reversal potential associated with this increase in conductance was −79.1 ± 1.6 mV (n= 5), which is close to the calculated value for EK (−84 mV) under these recording conditions, indicating an increase in K+ conductance.
Unequivocal identification of KATP channels as the molecular target for these actions of DNase 1 was obtained from cell-attached and inside-out recordings. Application of DNase 1 (50 μg ml−1) to the bath during cell-attached recordings induced an increase in the activity of single potassium channels (Fig. 2C and D). There was a delay of approximately 5–10 min following application of DNase 1 before the potassium channel activation occurred. Analysis of total channel current (over a 120 s period), 1–3 min after addition of DNase 1, resulted in a mean channel activity (NfPo) of 0.04 ± 0.01, which increased to 0.39 ± 0.21 (n= 4; P < 0.05), 15–20 min later. Identical control experiments in the absence of DNase 1 (Fig. 2C and D) resulted in no increase in potassium channel activity over the same time periods, with mean values of 0.06 ± 0.02 and 0.08 ± 0.03 (n= 12; P > 0.1), respectively. The effects of DNase 1 on membrane potential, macroscopic and single channel currents were reversibly antagonised (n= 4) by application of the sulphonylurea tolbutamide (100 μm; Fig. 2A and C). Inside-out recordings were made in symmetrical K+-containing solution at a membrane potential of −40 mV. With 0.1 mm Mg-ATP present in the bathing solution KATP channel activity was reduced, relative to control, by 72.3 ± 5.5 % (n= 8). Subsequent exposure to DNase 1 (50 μg ml−1), resulted in a rapid (2–4 min) increase (by 661.7 ± 32.6 %; n= 8) in KATP channel activity (Fig. 2E), which was essentially irreversible since washout of this agent over a 10 min period failed to attenuate channel activity. Furthermore, following washout of DNase 1, KATP channel activity was relatively resistant to inhibition by 1 mm Mg-ATP (Fig. 2E), a mean decrease of 13.1 ± 6.7 % being observed (n= 3). The reduced sensitivity towards intracellular ATP following disruption of the actin microfilament network is in agreement with previous reports (Brady et al. 1996).
The effect of a structurally distinct actin filament disrupter, cytochalasin B (Fukada et al. 1981), was also examined on KATP macroscopic and single channel currents. In whole cell recordings, the presence of 10 μm cytochalasin B in the electrode solution (with 5 mm Mg-ATP) hyperpolarised CRI-G1 cells from a mean resting membrane potential of −35.1 ± 2.7 mV (determined 1–2 min after formation of whole cell configuration) to −65.8 ± 5.6 mV (after 15–18 min) and this was accompanied by an increase in mean slope conductance from 0.64 ± 0.09 to 1.37 ± 0.22 nS (n= 5; Fig. 3A and B). The reversal potential associated with the increased conductance was −78.4 ± 0.8 mV (n= 5), indicating opening of potassium channels. Cell-attached recordings (not illustrated) also show that bath application of cytochalasin B (10 μm), after a delay of 5–10 min, increased mean potassium channel activity from a control value of 0.03 ± 0.01 to 0.14 ± 0.06 (n= 3; P < 0.05). These effects of cytochalasin B were also antagonised reversibly (n= 4) by application of tolbutamide (100 μm). Furthermore, addition of 10 μm cytochalasin B to inside-out patches in the presence of 0.1 mm Mg-ATP rapidly increased (by 709.1 ± 44.1 %) channel activity (n= 6; Fig. 3C). Following washout of this agent, channel activity did not return to control values and consistent with the action of DNase 1, addition of 1 mm Mg-ATP had little effect on channel activity (Fig. 3C).
In order to be assured that the actions of DNase 1 and cytochalasin B on KATP channel activity are specifically via actin filament disruption, their responses in the presence of phalloidin were determined. Cells dialysed with an electrode solution containing 5 mm Mg-ATP, 10 μm phalloidin and 50 μg ml−1 DNase 1 (Fig. 4A) had resting membrane potential and slope conductance values following dialysis for at least 20 min of −40.2 ± 1.7 mV and 0.51 ± 0.08 nS, respectively, which were not significantly (P > 0.1) different from control values obtained immediately after formation of the whole cell recording configuration (−40.7 ± 0.92 mV and 0.53 ± 0.09 nS; n= 4). Similar data were obtained with 10 μm cytochalasin B in the electrode solution in place of DNase 1 (data not shown), where the equivalent membrane potential and slope conductance values were −33.6 ± 3.4 mV and 0.69 ± 0.11 nS, respectively, after 20 min compared to control values of −36.0 ± 1.3 mV and 0.75 ± 0.02 nS (n= 4; P > 0.1). The presence of phalloidin (10 μm) in the bath solution also prevented activation of KATP channels by the actin disrupters in the inside-out patch configuration (Fig. 4B). Application of 0.1 mm Mg-ATP and 10 μm phalloidin to the cytoplasmic aspect of inside-out patches resulted in a 78.3 ± 7.4 % inhibition of channel activity (n= 8). Subsequent addition of 50 μg ml−1 DNase 1 failed to activate KATP channels in all eight patches; the mean channel activity was 0.11 ± 0.04 compared to a mean value of 0.23 ± 0.09 (P > 0.1) before DNase 1. Similar data were obtained on addition of 0.1 mm Mg-ATP and 10 μm cytochalasin B in the presence of 10 μm phalloidin (data not shown), the mean KATP channel activities being 0.71 ± 0.36 and 0.54 ± 0.40 (n= 5; P > 0.1) in the absence and presence, respectively, of the actin disrupter.
Antimicrotubule agents do not affect KATP channel activity
As changes in microtubule structure have also been implicated in the control of ion channel activity (Hamm-Alvarez & Sheetz, 1998; Morris et al. 1998; Unno et al. 1999) we also examined the actions of agents that specifically depolymerise (colchicine or nocodazole) or stabilise (taxol) microtubules. In whole cell recordings, with taxol (30 μm) added to the electrode solution (in the presence of 5 mm Mg-ATP) the initial (within 1–2 min) resting membrane potential and slope conductance of CRI-G1 cells were −34.5 ± 2.2 mV and 0.60 ± 0.07 nS (n= 4), respectively. After 15–20 min dialysis there was no significant change (P > 0.1) in the membrane potential (−35.0 ± 2.3 mV) or slope conductance (0.65 ± 0.08 nS) of the cells (Fig. 5A). Taxol (30 μm) also had no effect on KATP channel activity in cell-attached recordings (n= 3; data not illustrated) or when applied in the bath solution to inside-out patches in the presence of 0.1 mm Mg-ATP (n= 7; data not illustrated). Furthermore, disruption of the microtubule component of the cytoskeleton with nocodazole or colchicine also failed to increase KATP channel activity in this cell line, in agreement with previous studies on KATP channels in cardiac myocytes (Terzic & Kurachi, 1996). Thus, application of either nocodazole (10 μm) or colchicine (100 μm) to the bath during cell-attached recordings (Fig. 5B) failed to increase the activity of single potassium channels. Analysis of total channel current (over a 120 s period), 4–6 min after obtaining the cell-attached configuration, resulted in a mean channel activity (NfPo) of 0.09 ± 0.07 (n= 4; nocodazole) and 0.04 ± 0.01 (n= 3; colchicine), which did not increase following application of either agent for at least 40 min (nocodazole: 0.08 ± 0.02, n= 3,P > 0.05; colchicine: 0.05 ± 0.02, n= 3,P > 0.05). Identical control experiments in the absence of either antimicrotubule agent resulted in no increase in potassium channel activity over the same time periods, with mean values of 0.04 ± 0.02 and 0.04 ± 0.02 (n= 3; P > 0.05), respectively.
These studies, therefore, show that the actions of DNase 1 and cytochalasin B are specific for actin filaments, and clearly implicate the process of actin filament depolymerisation in the activation of KATP channels.
Leptin induces PI 3-kinase-dependent disassembly of actin filaments
The effects of the actin depolymerising agents and of leptin on KATP channel activity in this cell line have many features in common; i.e. there is a 5–10 min delay before the initiation of KATP channel activation, whether monitored as membrane hyperpolarisation and increased conductance in whole-cell recordings, or as a change in single channel activity, and these responses are blocked by phalloidin. Consequently, it appears likely that leptin-induced KATP channel activation in CRI-G1 cells is through the destabilisation of actin filaments. To determine directly if leptin receptor activation induces reorganisation of the actin cytoskeleton, F-actin was visualised by staining with rhodamine-conjugated phalloidin, and the effects of leptin and cytochalasin B were compared (Fig. 6). In control cells there was pronounced phalloidin-positive labelling of the cell membrane and a more diffuse, granular staining within the cytoplasmic compartment (Fig. 6Aa and c). In contrast, cells treated with either 10 nm leptin or 10 μm cytochalasin B for 30–40 min displayed a much reduced intensity of phalloidin labelling, at the level of both the cell membrane (which exhibited a more punctate signal) and the cytosol (Fig. 6Ab and d). A quantitative assessment of the actions of leptin and cytochalasin B on rhodamine-conjugated phalloidin staining at the cell membrane was made. Cytochalasin B and leptin induced a significant reduction, by 63.4 ± 3.2 % (n= 7; 76 cells) and 66.2 ± 2.1 % (n= 5; 42 cells), respectively, of the intensity of phalloidin labelling (Fig. 6B). Previous studies have demonstrated that leptin activation of KATP channels in CRI-G1 cells requires activation of PI 3-kinase (Harvey & Ashford, 1998). Prior incubation of cells with the PI 3-kinase inhibitor LY 294002 (10 μm; for 10 min; n= 5) in the absence and presence of leptin caused a small (by 17 ± 2 and 14 ± 4 %, respectively) but significant (P < 0.05) reduction in rhodamine-conjugated phalloidin staining compared with control (Fig. 6Ae). However, the presence of 10 μm LY 294002 did prevent the leptin-induced reduction in staining (P > 0.05 when compared with LY 294002 alone; n= 5; Fig. 6A and B). These data indicate that leptin destabilises F-actin by a mechanism dependent upon the activity of PI 3-kinase.
PtdIns(3,4,5)P3-induced activation of KATP channels is prevented by phalloidin
There is an increasing amount of literature implicating a significant role for phosphatidylinositol phosphate kinase isoforms and phosphoinositol lipid products in the regulation of the assembly and disassembly of actin filaments in vitro (Zigmond, 1996; Jamney, 1998). The majority of studies have implicated phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2) as an important modulator of actin structural re-arrangements (e.g. Fukami et al. 1992; Sakisaka et al. 1997). Furthermore, a number of recent studies have shown that PtdIns(4,5)P2 can activate native and cloned KATP channels when applied to the cytoplasmic aspect of isolated inside-out membrane patches (Hilgeman & Ball, 1996; Shyng & Nichols, 1998; Baukrowitz et al. 1998). However, we have shown (Harvey et al. 2000) that PtdIns(4,5)P2-mediated KATP channel activation, in CRI-G1 cells, is prevented by PI 3-kinase inhibitors and that leptin-induced KATP channel activity probably involves an increase in a PI 3-kinase lipid product, possibly phosphatidylinositol 3,4,5-trisphosphate (PtdIns(3,4,5)P3). Thus, we have investigated the effects of phalloidin, on the ability of PtdIns(3,4,5)P3 to activate KATP channels.
In control experiments, CRI-G1 cells were dialysed with an electrode solution containing 5 mm ATP and 1 μm PtdIns(3,4,5)P3. Immediately after obtaining the whole cell configuration (1–2 min) the mean resting membrane potential and slope conductance were −36.2 ± 3.1 mV and 0.72 ± 0.05 nS, respectively (n= 4). Within another 5 min of dialysis with PtdIns(3,4,5)P3, a slowly developing membrane hyperpolarisation to −64.3 ± 1.7 mV (n= 4) was observed, which was accompanied by an increase in the slope conductance to 1.95 ± 0.29 nS (n= 4; Fig. 7A and B). The sulphonylurea tolbutamide (100 μm) completely reversed the hyperpolarisation and increase in slope conductance induced by PtdIns(3,4,5)P3 to control levels. Thus these data indicate that PtdIns(3,4,5)P3 activates KATP channels in CRI-G1 insulin-secreting cells, in agreement with our previous study using this cell line (Harvey et al. 2000).
However, when cells were dialysed with a pipette solution containing 5 mm ATP, 10 μm phalloidin and 1 μm PtdIns(3,4,5)P3, no change in membrane potential or slope conductance was observed over the equivalent time period (Fig. 7C and D). The initial (1–2 min) mean resting membrane potential and slope conductance values were −33.3 ± 2.4 mV and 0.83 ± 0.13 nS, respectively (n= 3), and after at least 20 min dialysis were not significantly altered (P > 0.05), being −32.7 ± 4.3 mV and 0.91 ± 0.09 nS (n= 3), respectively. Thus, stabilisation of actin filaments with phalloidin prevents KATP channel activation by PtdIns(3,4,5)P3 in CRI-G1 cells, indicating that, like leptin, the PtdIns(3,4,5)P3-induced opening of KATP channels involves disruption of the actin cytoskeleton.
Previous studies have shown that altering the integrity of cytoskeletal networks modulates KATP channel function. For example, in cardiac myocytes (Terzic & Kurachi, 1996; Yokoshiki et al. 1997), disruption of the actin, rather than microtubule, component of the cytoskeleton has been shown to increase KATP channel activity. The present study also demonstrates that depolymerisation of actin filaments results in activation of KATP channels in CRI-G1 insulinoma cells, and this effect is blocked by stabilisation of polymerised actin with phalloidin. Furthermore, in parallel with cardiac KATP channels, depolymerisation or stabilisation of microtubules failed to affect KATP channel activity in CRI-G1 cells. Interestingly, the rundown of cardiac KATP channels is also regulated by the state of the actin, but not the microtubule, polymerisation-depolymerisation cycle (Furukawa et al. 1996). The modulatory actions of cytoskeleton disrupters on KATP channel function observed in the present study parallel the actions of these agents on another ATP binding cassette ion channel protein system, the cystic fibrosis transmembrane conductance regulator (CFTR) Cl− channel. Thus, in mouse adenocarcinoma cells (Prat et al. 1995), mouse 3T3 fibroblasts (Fischer et al. 1995) and rat ventricular myocytes (Cantiello, 1996), disassembly of actin filaments results in the activation of CFTR Cl− channels. Furthermore, in a manner similar to KATP channels, microtubule disruption failed to affect CFTR Cl− channel activity. Actin cytoskeletal disrupters are also reported to increase, in a phalloidin-dependent manner, the activity of other channel types, including voltage-gated K+ channels in retinal bipolar neurones (Maguire et al. 1998) and epithelial Na+ channels (Cantiello et al. 1991). In contrast, however, actin depolymerisation reduces NMDA channel activity in cultured hippocampal neurones (Rosenmund & Westbrook, 1993) and the activity of voltage-gated sodium (Mironov & Richter, 1999) and calcium channels (Johnson & Byerly, 1993; Unno et al. 1999) can be modulated by disruption of both the actin and microtubule components of the cytoskeleton. Thus, like numerous other channel types the activity of KATP channels is influenced by the dynamics of the cytoskeleton.
In the present study the sensitivity of KATP channels to ATP inhibition is attenuated after exposure to DNase 1. This finding parallels reports of reduced ATP sensitivity of cardiac KATP channels following activation by actin depolymerisation (Terzic & Kurachi, 1996). However, the ability of the sulphonylurea tolbutamide to inhibit KATP channels after treatment with actin disrupters in this cell line is unaffected. This is in contrast to the reduced sensitivity of cardiac KATP channels to sulphonylureas following disruption of the actin cytoskeleton (Brady et al. 1996; Yokoshiki et al. 1997). This differential sensitivity to ATP and tolbutamide may be related to the different forms of sulphonylurea receptor present in cardiac tissue (SUR2A) and pancreatic β-cells (SUR1; Babenko et al. 1998). The selective attenuation of ATP sensitivity in this cell line is consistent with ATP and tolbutamide acting at different sites on the KATP channel complex, as suggested in previous studies (Tucker & Ashcroft, 1998) and, in particular, indicates that the binding site for ATP inhibition is closely associated with the cytoskeleton.
Recently, we and others have shown that, in the CRI-G1 insulin-secreting cell line (Harvey et al. 1997) and pancreatic β-cells (Kieffer et al. 1997), the ob gene product leptin activates KATP channels. This action of leptin is likely to require activation of PI 3-kinase, as the effects are prevented by inhibitors of PI 3-kinase (Harvey & Ashford, 1998; Harvey et al. 2000). The present study demonstrates that activation of KATP channels by leptin also involves depolymerisation of actin filaments since phalloidin prevented the leptin membrane hyperpolarisation and increase in K+ conductance. Furthermore, the PI 3-kinase inhibitor LY 294002 occluded the leptin-induced reduction in phalloidin staining, indicating that leptin destabilises actin by a mechanism dependent on PI 3-kinase activity. Together these data indicate that activation by leptin of KATP channels in CRI-G1 cells involves PI 3-kinase-dependent disruption of the actin cytoskeleton. The involvement of the actin cytoskeleton in leptin activation of KATP channels in this cell line parallels the role of the actin cytoskeleton proposed for the signalling pathways of other cytokines and growth factor receptors. For example, stimulation of porcine aortic endothelial cells with platelet-derived growth factor (PDGF) leads to marked reorganisation of actin microfilaments (Valgeirsdottir et al. 1998), whereas growth hormone is reported to depolymerise actin microfilaments in Chinese hamster ovary (CHO) cells (Goh et al. 1997). Furthermore, in human SH-SY5Y neuroblastoma cells (Kim & Feldman, 1998), insulin-like growth factor (IGF) 1 evokes a PI 3-kinase-dependent reorganisation of the cytoskelelton.
There is growing evidence that inositol phosphates closely associate with the cytoskeleton and can modulate the activity of a variety of proteins (Machesky & Hall, 1996; Janmey, 1998). Recent studies also indicate that PtdIns(3,4,5)P3, the lipid product of PI 3-kinase, activates both native and cloned KATP channels (Shyng & Nichols, 1998; Baukrowitz et al. 1998; Harvey et al. 2000). Thus leptin disruption of actin filaments may involve a PtdIns(3,4,5)P3-dependent process. Consistent with this possibility, phalloidin prevented the PtdIns(3,4,5)P3-driven activation of KATP channels in this cell line. This is also consonant with the hypothesis that a leptin-driven increase in PtdIns(3,4,5)P3 must occur prior to depolymerisation of actin filaments and activation of KATP channels. A consequence of this result is that actin depolymerisation per se appears to activate KATP channels, and the activation is not due to a direct interaction of phosphoinositides with channel components. It will be interesting now to determine the mechanism by which PtdIns(3,4,5)P3 acts on actin filaments in order to activate KATP channels. A recent study (Missy et al. 1998) has demonstrated that PtdIns(3,4,5)P3 can selectively associate with the Rho family of GTPases, in particular Rac1, and these proteins are key regulators of the actin filament structure (Settleman, 2000).
In conclusion, these data indicate that in CRI-G1 insulin-secreting cells, leptin activation of KATP channels involves a complex and novel signalling pathway that ultimately leads to reorganisation of the actin cytoskeleton. Since leptin also activates KATP channels in hypothalamic glucose-receptive neurones, it will be interesting to determine whether the leptin receptor transduction pathway in these neurones also involves disruption of the cytoskeleton. This may be particularly important in obese individuals who display resistance to leptin since leptin resistance is most probably due to a defect in the signalling downstream of the leptin receptor. Therefore, the involvement of the cytoskeleton in leptin activation of KATP channels may be an important factor in our understanding of the signalling pathways underlying these defects.
This work is supported by The Wellcome trust (grant nos 042726, 047368 and 055291). J.H. and A.J.I. are Wellcome Research Career Development Fellows. We thank Emma Horn for assistance with cell culturing.