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The Journal of Physiology logoLink to The Journal of Physiology
. 2005 Jan 13;563(Pt 2):471–482. doi: 10.1113/jphysiol.2004.080333

Inhibition of cellular responses to insulin in a rat liver cell line. A role for PKC in insulin resistance

Livia Puljak 3, Michael J Pagliassotti 1, Yuren Wei 1, Ishtiaq Qadri 2, Vinay Parameswara 3, Victoria Esser 3, J Gregory Fitz 3, Gordan Kilic 3
PMCID: PMC1665596  PMID: 15649984

Abstract

The initial response of liver cells to insulin is mediated through exocytosis of Cl− channel-containing vesicles and a subsequent opening of plasma membrane Cl− channels. Intracellular accumulation of fatty acids leads to profound defects in metabolism, and is closely associated with insulin resistance. It is not known whether the activity of Cl− channels is altered in insulin resistance and by which mechanisms. We studied the effects of fatty acid accumulation on Cl− channel opening in a model liver cell line. Overnight treatment with amiodarone increased the fat content by ∼2-fold, and the rates of gluconeogenesis by ∼5-fold. The ability of insulin to suppress gluconeogenesis was markedly reduced indicating that amiodarone treatment induces insulin resistance. Western blot analysis showed that these cells express the same number of insulin receptors as control cells. However, insulin failed to activate exocytosis and Cl− channel opening. These inhibitory effects were mimicked in control cells by exposures to arachidonic acid (15 μm). Further studies demonstrated that fatty acids stimulate the PKC activity, and inhibition of PKC partially restored exocytosis and Cl− channel opening in insulin-resistant cells. Accordingly, activation of PKC with PMA in control cells potently inhibited the insulin responses. These results suggest that stimulation of PKC activity in insulin resistance contributes to the inhibition of cellular responses to insulin in liver cells.

Insulin plays a key role in the regulation of liver metabolism. Recent studies demonstrated that the initial response of liver cells to insulin is mediated through exocytosis of a distinct population of vesicles that contain Cl− channels in their membranes (Kilic et al. 2001b). Insertion of these vesicles into the plasma membrane is directly responsible for a 15-fold increase in membrane conductance resulting from the opening of Cl− channels. Multiple lines of evidence suggest that Cl− channel activation modulates membrane potential and the intracellular pH, and these changes regulate a broad range of transport and metabolic processes in hepatocytes (Friedmann & Dambach, 1980; Boyer et al. 1992; Li & Weinman, 2002). Thus, definition of the cellular mechanisms involved in modulation of the Cl− channel activity is critical for understanding regulation of the liver metabolism. The mechanisms responsible for Cl− channel opening in response to different physiological stimuli have been described (Li & Weinman, 2002; Nilius & Droogmans, 2003). However, little is known about the cellular mechanisms that regulate opening of Cl− channels by insulin.

Numerous studies have firmly established that the intracellular accumulation of fat in the liver and muscle cells is a hallmark of insulin resistance syndrome (i.e. obesity, diabetes mellitus type II, fatty liver disease) (Shulman, 2000; Saltiel & Kahn, 2001). The excess of intracellular fat in liver is caused by the alterations in hepatocyte fatty acid transport, synthesis or metabolism (Koteish & Diehl, 2001). Notably, insulin resistance is characterized with a prominent inability of insulin to inhibit glucose production from gluconeogenesis. Based on the studies in human and animal models, two working models have been proposed. According to one model, intracellular fatty acids and their metabolites stimulate phosphorylation of the specific proteins in insulin signalling pathways, resulting in a reduced capacity of insulin to activate the downstream pathways necessary for insulin actions (Shulman, 2000; Yu et al. 2002). In another model, severe insulin resistance has been produced when the number of hepatocyte insulin receptors was decreased utilizing gene technology (Michael et al. 2000). These results imply that insulin resistance in the liver may result from inhibition of insulin signalling pathways or the decreased number of insulin receptors. While the cellular mechanisms responsible for defective regulation of gluconeogenesis have been well defined, it is not known whether the early cellular responses to insulin (exocytosis and Cl− channel opening) are altered in insulin resistance, and by which mechanisms.

The purpose of these studies was to examine the effects of fatty acid accumulation on the initial cellular response to insulin in liver cells. We found that the insulin-dependent exocytosis and Cl− channel opening are potently inhibited under these conditions. Notably, this inhibition was mediated in part by stimulation of the protein kinase C (PKC) activity, suggesting that PKC isoforms may be potential targets for modulation of the defective metabolism through effects on exocytosis and Cl− channel opening.

Methods

Cell preparation

All experiments were performed in HTC cells, a model liver cell line derived from rat hepatoma. HTC cells were obtained from Dr. Steven Lidafsky (University of Vermont) These cells have been used widely as a model of hepatocyte membrane Cl− transport because they have Cl− channels and signalling pathways analogous to those found in primary hepatocytes (Bodily et al. 1997; Roman & Fitz, 1999). Cells were plated on coverslips and maintained at 37°C in a 5% CO2 and 95% air atmosphere in culture medium containing Dulbecco's modified Eagle's medium (DMEM, Life Technologies, Inc.) supplemented with 10% fetal calf serum, 2 mml-glutamine, 100 IU ml−1 penicillin and 100 μg ml−1 streptomycin.

Detection of intracellular fat

The intracellular fat content of HTC cells was increased by incubating cells overnight with amiodarone (50 μm, Sigma, St Louis, MO, USA). Amiodarone is a potent inhibitor of fatty acid oxidation and lipoprotein export, and leads to the accumulation of lipids in hepatocytes (Fromenty et al. 1990; Letteron et al. 2003). The amount of fat was measured utilizing a fluorescent lipid marker, Nile red. Nile red is not fluorescent in water but becomes fluorescent upon binding to fatty acids and triacylglycerols (Greenspan & Fowler, 1985; Fowler et al. 1987). Consequently, the fluorescence intensity of Nile red is directly proportional to the amount of cytosolic fat. For these experiments, cells were preincubated with 1 μm Nile red for 2 h. Fluorescence was excited through an excitation filter (peak at 535 nm) and collected with an emission filter (peak at 610 nm). Data from different cells were compared by acquiring images with the same duration of light exposure of 25 ms. Similar procedures have been utilized for the quantification of intracellular fat content in other cells (Greenspan & Fowler, 1985; Ostermeyer et al. 2001; Al-Saffar et al. 2002).

Adenovirus preparation and transduction

Adenoviruses used contained the cDNAs encoding Escherichia coliβ-galactosidase or rat CPT1a under the control of the early cytomegalovirus promoter. Viral stocks/cell lysates were prepared with the human embryonal kidney cell line 293A as host, as previously described (Becker et al. 1994). Adenoviral transduction of CPT1a was performed 2 h after HTC cells were attached to the coverslips. Cells were incubated at 37°C for 90 min in 0.75 ml of DMEM containing 1010 particles of virus. The medium was then aspirated, and cells were washed with phosphate-buffered saline prior to addition of the fresh DMEM. An equivalent titre of AdV–β-gal, prepared in an identical fashion, was used to transduce control cells. In all cases, cells were incubated overnight for ∼18 h to allow expression of the virally encoded proteins before commencing the procedures.

Glucose release

Dishes with HTC cells were washed twice with phosphate-buffered saline (PBS) and DMEM containing 5 mm lactate and 0.5 mm pyruvate, and no glucose. Insulin, when present, was provided at a concentration of 1 nm. Samples of medium were taken at 0, 0.5, 1, 2, 3, and 4 h. Glucose was analysed using Sigma Kit HK-20.

Western blot analysis

Cells were washed three times in ice cold PBS. Total cell lysates were prepared from ∼107 cells by adding 0.5 ml of lysis buffer containing: 10 mm Tris, 100 mm NaCl, 1 mm EDTA, 1 mm EGTA, 1 mm NaF, 20 mm Na2P2O7, 2 mm Na3VO4, 0.5% sodium deoxycholate, 1% Triton X-100, 10% glycerol, 1 mm PMSF, 60 mg ml−1 aprotinin, 10 mg ml−1 leupeptin and 1 mg ml−1 pepstatin (pH 7.4). The lysates were centrifuged at 4°C for 10 min at 21 000 g to remove nuclei and cell debris. The supernatants were saved and used for Western blot analysis. SDS-PAGE and immunoblotting were carried out using minigels. Proteins were transferred according to the procedures previously described (Towbin et al. 1979), and processed using ECL for detection (Amersham) with specific antibodies against the insulin receptor (-subunit (Biosource) using 3% BSA in Tris-buffered saline (TBS). All washes were in 0.5% Tween/TBS for 5 min. Autoradiograms were analysed by a PhosphoImager. Band detection and analysis was performed using a Kodak Image Station CF440. Protein concentrations were determined by the bicinchoninic acid method (Pierce, Rockford, IL, USA).

Measurements of exocytosis

Exocytosis was assessed by real time imaging using a fluorescent probe FM1-43 (Molecular Probes, Eugene, OR, USA). FM1-43 binds to the membranes but does not cross lipid bilayers. The dye is not fluorescent in solution, but when it binds to biological membranes its quantum yield increases ∼350 times (Betz et al. 1996). Thus, the fluorescence intensity is directly proportional to the amount of membrane exposed to FM1-43. FM1-43 was added to the external solution at a concentration of 4 μm. Initially, FM1-43 partitions into the plasma membrane exposed to external solution. Subsequently, when vesicles fuse with the plasma membrane, FM1-43 equilibrates with the new membrane, resulting in an increase in the apparent fluorescence. Consequently, the overall change in FM1-43 fluorescence provides, in real time, a measure of the sum of all exocytic events. The fluorescence of FM1-43 was excited through an excitation filter (peak at 480 nm) and collected with an emission filter (peak at 535 nm). Images were taken every 30 s using exposures of 200 ms in duration. These methods have been utilized previously to study exocytosis in different cells (Cochilla et al. 1999).

Measurements of [Ca2+]i

Intracellular [Ca2+] was measured utilizing a Ca2+-sensitive fluorescent dye Fluo-3 AM (Molecular Probes) as described (Diaz et al. 2001). Fluo-3 AM was dissolved in DMSO with equal amounts of Pluronic F-127 (10% in water). Cells were incubated with 20 μm Fluo-3 AM for 30 min. After washing the dye from extracellular solution, Fluo-3 fluorescence was excited through an excitation filter (peak at 480 nm) and collected with an emission filter (peak at 535 nm). Images were taken every 10 s using exposures of 50 ms in duration. [Ca2+]i was determined using the expression:

graphic file with name tjp0563-0471-m1.jpg

where Kd was taken as 400 nm (Auld et al. 2000). F is a cellular Fluo-3 fluorescence, and Fmax is the fluorescence at saturating [Ca2+]i levels, obtained at the end of each experiments by exposing cells to 5 μm ionomycin.

Detection of PKC activity

Stimulation of PKC activity leads to kinase translocation from cytosolic compartments to the plasma membrane (Newton, 2001). The levels of PKC at the plasma membrane in HTC cells were assessed utilizing a fluorescent dye fim-1 AM. Fim-1 binds to a catalytic binding site on various PKC isoforms, and kinase activation increases the plasma membrane fluorescence (rim fluorescence) (Chen & Poenie, 1993; Ueda et al. 1996; Dupont et al. 2000). Fim-1 AM was dissolved in dimethyl sulphoxide (DMSO) and Pluronic F-127. Cells were preincubated with 20 μm fim-1 AM for 30 min. To quantify PKC activity, the fluorescence intensity of a 1 μm wide rim around the plasma membrane was measured. Preliminary measurements of the total cellular fim-1 fluorescence and quantitative Western blots of different PKC isoforms (authors' unpublished data) suggested that the expression of PKC isoforms varies substantially under different experimental conditions. Thus, to eliminate these variations, the rim fluorescence was normalized to total cellular fluorescence. Furthermore, similar to fibroblasts, fim-1 stains mitochondria in HTC cells (Chen & Poenie, 1993), and these regions were excluded from measurements of the rim fluorescence. Fim-1 fluorescence was excited through an excitation filter (peak at 480 nm) and collected with an emission filter (peak at 535 nm). Images were obtained utilizing exposures of 1.5 s in duration.

Imaging and analysis

Coverslips with HTC cells were perfused in a chamber at a rate that allowed complete exchange of chamber volume in ∼1 min. Cells were viewed through an Olympus objective (60×, oil immersion, NA = 1.4). Fluorescent images were acquired with a 12-bit SensicamQE camera controlled by SlideBook 3.0 software (Intelligent Imaging Innovations, Denver, CO, USA). Quantitative analyses of fluorescent images were performed on a Macintosh computer using NIH Image (National Institutes of Health, Bethesda, MD, USA) and IgorPro3 (WaveMetrics, Lake Oswego, OR, USA) software. Total cellular fluorescence of Nile red, FM1-43 and Fluo-3 was measured from the region containing the cell. For background subtraction, the fluorescence was measured in the same way from regions containing no cells. After background subtraction, the fluorescence intensity of FM1-43 was normalized to the values obtained immediately after staining plasma membrane with FM1-43. For Nile red studies, the fluorescence intensity was expressed relative to the values obtained in control cells.

Conductance and current measurements

Plasma membrane conductance was measured in the whole-cell configuration utilizing patch clamp techniques. Cells were voltage-clamped at 40 mV, and membrane conductance was determined every 3 s by applying 4 ms voltage pulses (pulse amplitude, −40 mV). The current response was used to determine the conductance and the capacitance as previously described (Lindau & Neher, 1988). To compare insulin-evoked conductance responses from different cells, the conductance was normalized to the initial capacitance (measure of cell surface area) and expressed in pS pF−1.

Whole-cell currents were measured in response to insulin, after membrane capacitance and access resistance were compensated. The whole-cell currents in response to voltage pulses (0.5 s in duration) were filtered with an 8-pole Bessel filter at a 1 kHz cut-off frequency and sampled every 0.5 ms. To determine reversal potential, a voltage ramp from −90 mV to 90 mV (duration, 0.2 s) was applied.

Solutions

All experiments were performed after washing the culture media with an extracellular solution that contained no amidarone. This solution contained: 142 mm NaCl, 4 mm KCl, 1 mm KH2PO4, 2 mm MgCl2, 2 mm CaCl2, 10 mmd-glucose, 10 mm Hepes/NaOH and no amiodarone. For patch clamp recordings, cells were dialysed with a pipette solution that contained: 130 mm KCl, 10 mm NaCl, 1 mm EGTA, 0.5 mm CaCl2, 2 mm MgCl2, and 10 mm Hepes/NaOH (free [Ca2+]∼0.1 μm). To increase the intracellular concentration of fatty acids, in some experiments 15 μm arachidonic acid was added to the pipette solution. The pH of all solutions was 7.25. Osmolality of the extracellular solution was 300 mosmol kg−1, and the osmolality of the pipette solution was 270–275 mosmol kg−1. All compounds were purchased from Sigma.

Statistics

Data were expressed as mean ± s.e.m. Results were compared using Student's t test on paired and on unpaired data.

Results

Insulin resistance in liver is closely associated with the accumulation of fat in hepatocytes. To increase the intracellular fat content, HTC cells were treated overnight with amiodarone, which is known to induce fatty liver in mice (Letteron et al. 2003). After washing with extracellular solution that contained no amiodarone, the lipid content was assessed by measuring the fluorescence of Nile red. Representative images of control and amiodarone-treated HTC cells are shown in Fig. 1A, indicating that amiodarone treatment results in a ∼2-fold increase in intracellular lipid content (Fig. 1B). To evaluate the contribution of fatty acids in this increase, carnitine palmitoyltransferase 1a (CPT1a) was overexpressed in HTC cells utilizing adenovirus technology (Perdomo et al. 2004). CPT1a plays a key role in fatty acid metabolism in liver cells by transporting fatty acids from the cytosol into mitochondria for further degradation. Overexpression of CPT1a significantly reduced Nile red fluorescence in amiodarone-treated cells (Fig. 1B), confirming that amiodarone increases the concentration of fatty acids in HTC cells.

Figure 1. Intracellular accumulation of fatty acids.

Figure 1

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Prior to imaging, HTC cells were incubated with a lipid marker Nile red (1 μm) for 2 h. A, representative fluorescent images of control (left panel) and amiodarone-treated cells (50 μm overnight exposure, right panel) are shown. Note that amiodarone treatment increases Nile red fluorescence. Scale bar 12 μm. B, Nile red fluorescence was measured in control (24 cells), and cells that were transduced with adenovirus containing β-galactosidase (Adv, 19 cells) or CPT1a (42 cells) as described in Methods. The fluorescence of Adv- and CPT1a-treated cells was not different from control (P > 0.38 for both). After treatment with amiodarone, Nile red fluorescence was significantly smaller in CPT1a-overexpressing cells (32 cells) than in control cells (48 cells, P < 0.003).

Hepatic insulin resistance is characterized by higher rates of constitutive glucose production, and a reduced capacity of insulin to suppress gluconeogenesis (DeFronzo, 1999). To assess whether excess of fatty acids alters glucose metabolism, glucose production from lactate and pyruvate was measured in control and amiodarone-treated HTC cells. Data summarized in Fig. 2 show that amiodarone-treated cells produced ∼6 times more glucose than control cells. Notably, the relative ability of insulin to suppress gluconeogenesis was markedly reduced. These findings indicate that amiodarone-treated HTC cells develop a phenotype characteristic of insulin resistance, and represent a cell model with a prominent defect in the regulation of gluconeogenesis.

Figure 2. Insulin and gluconeogenesis.

Figure 2

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HTC cells were incubated for 4 h in the presence of gluconeogenic substrates (5 mm lactate, 0.5 mm pyruvate), and no glucose. Control and amiodarone-treated cells were exposed to insulin (1 nm). The experiments were done in triplicate. Glucose release is expressed in nmol h−1 (mg of protein)−1. Insulin inhibited gluconeogenesis in HTC cells by 70% (P < 0.03), and only by 25% in amiodarone-treated HTC cells (P < 0.02).

Binding of insulin to the α-subunit of the insulin receptor is necessary for activation of insulin signalling pathways (Saltiel & Kahn, 2001). Consequently, we examined whether insulin resistance is caused by a decrease in the number of insulin receptors. In Fig. 3 Western blot analysis, obtained utilizing specific antibodies against the insulin receptor α-subunit, demonstrates that amiodarone-treated HTC cells express the same number of insulin receptors as control cells.

Figure 3. Expression of insulin receptors.

Figure 3

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Representative Western blots of insulin receptor α-subunit immunoprecipitates obtained from control and amiodarone-treated HTC cells. Bottom graph shows relative band densities from 4 experiments. The densities were expressed relative to the values obtained in control cells.

The initial response of HTC cells to insulin is mediated through exocytosis of a distinct population of vesicles that contain Cl− channels in their membranes (Kilic et al. 2001b). To determine whether insulin resistance modulates this response, exocytosis and the conductance were measured under control conditions and after increasing fatty acid concentrations. The amount of exocytosis was measured utilizing a fluorescent membrane marker FM1-43 as previously described (Smith & Betz, 1996; Kilic et al. 2001b). In the absence of insulin, FM1-43 fluorescence gradually increased due to constitutive exocytosis at a rate of 2.3 ± 0.2% min−1 (8 cells) in control, 1.2 ± 0.2% min−1 (10 cells) in amiodarone-treated cells, and 5.3 ± 0.4% min−1 (8 cells) after exposure to arachidonic acid. Representative measurements in Fig. 4A and B show that insulin rapidly increased FM1-43 fluorescence in control cells by ∼10% within a few minutes of exposure. We have previously shown that this insulin-dependent exocytosis is distinct from constitutive exocytosis. Membrane capacitance of HTC cells was 21.8 ± 2.6 pF (9 cells) as measured in the whole-cell configuration with patch clamp techniques. Given that the cell capacitance is directly proportional to the plasma membrane area (∼10 fF μm−2) (Lindau & Neher, 1988), and assuming that a one-to-one relationship exists between the capacitance and FM1-43 fluorescence in HTC cells as found in other cells (Smith & Betz, 1996; Kilic et al. 2001a), these results imply that insulin stimulates insertion of ∼7000 distinct vesicles (0.1 μm in diameter) into the plasma membrane within minutes of exposure. Interestingly, the insulin-dependent exocytosis was markedly inhibited in amiodarone-treated HTC cells (Fig. 4B and C). Similar results were obtained in control cells by exposures to arachidonic acid (15 μm, Fig. 4C). Thus, insulin resistance in liver cells is associated with almost complete inhibition of the exocytic response to insulin.

Figure 4. Inhibition of insulin-dependent exocytosis.

Figure 4

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A, representative fluorescent images of a cell before (left panel) and 3 min after insulin exposure (10 μm, right panel) in the presence of FM1-43. Scale bar 5 μm. B, the time course of changes in FM1-43 fluorescence measured from control and amiodarone-treated HTC cells in response to insulin (at arrow). Prior to imaging, amiodarone was removed by washing with extracellular solution. FM1-43 fluorescence was expressed relative to the values obtained after initial staining the plasma membrane with the dye (100%). Note that insulin stimulates exocytosis in control but not in amiodarone-treated cells. C, exocytosis in response to insulin was measured as a change in cellular FM1-43 fluorescence before and 3 min after the insulin exposure in control (29 cells), amiodarone-treated cells (34 cells), or in the presence of 15 μm arachidonic acid (AA, 15 cells). For these experiments, cells were exposed to arachidonic acid 5 min before stimulation with insulin. The changes in FM1-43 fluorescence due to constitutive exocytosis were subtracted form the measurements. Note that insulin-dependent exocytosis is almost completely inhibited after amiodarone treatment (P < 10−5), or exposure to arachidonic acid (P < 10−4).

To assess whether the conductance response to insulin is also inhibited in insulin resistance, membrane conductance was measured utilizing patch-clamp techniques. A representative recording in Fig. 5A shows that insulin activates the conductance that gradually reaches a peak ∼10 min after exposure. Similar to exocytosis, insulin activated the conductance in a dose-dependent manner (1 nm to 10 μm, Fig. 5B), providing further support for the hypothesis that exocytosis of channel-containing vesicles may contribute to the conductance response (Kilic et al. 2001b). Notably, the insulin-dependent conductance was inhibited in amiodarone-treated cells (P < 0.02), or after intracellular dialysis with arachidonic acid (P < 0.05) (Fig. 5A and B). Thus, the conductance response of liver cells to insulin is potently inhibited in insulin resistance.

Figure 5. Inhibition of insulin-dependent Cl− conductance.

Figure 5

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A, representative patch-clamp recordings of plasma membrane conductance in response to insulin (10 μm, arrow) from control, amiodarone-treated cells, and after dialysis of 15 μm arachidonic acid through a patch pipette. The conductance was normalized to the cell capacitance. Dashed lines indicate basal conductance levels. Holding potential −40 mV. B, normalized peak conductance (pS pF−1) was measured in response to different insulin concentrations (open bars). The number of cells was 2–9. The conductance response to 10 μm insulin was also measured in amiodarone-treated HTC cells (7 cells), and after intracellular dialysis with arachidonic acid (AA, 5 cells). C, whole-cell currents were measured after exposure to insulin in response to the voltage pulses of varying amplitudes (0.5 s in duration). Pulse potentials (in mV) are shown on the right.

To further examine the insulin-dependent conductance, the reversal potential (Er) of the whole-cell currents evoked by insulin was measured. The Er was 8.4 ± 0.8 mV (9 cells). This value was close to the Cl− equilibrium potential of −1 mV which was determined according to the Nernst equation, and was indicative of the activation of Cl− channels as previously described (Kilic et al. 2001b). Moreover, the whole-cell currents evoked by insulin exhibited an outward rectification and a slow activation at positive holding potentials (Fig. 5C). To assess whether insulin activates Cl− channels, cells were exposed to a non-selective Cl− channel blocker, 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB). In the presence of 25 μm NPPB, conductance response to insulin decreased from 148 ± 36 pS pF−1 (9 cells) to 41.0 ± 4.3 pS pF−1 (5 cells). These biophysical properties are consistent with the previous work (Kilic et al. 2001b), suggesting that insulin stimulates opening of a distinct population of Cl− channels in liver cells.

Recent studies have demonstrated that the whole-cell currents activated by insulin or increases in intracellular [Ca2+] have similar properties (Kilic et al. 2001b; Kilic & Fitz, 2002). Consequently, to assess whether insulin increases [Ca2+]i levels, [Ca2+]i was monitored utilizing Fluo-3 fluorescence. A representative recording in Fig. 6A shows that insulin had no effect on [Ca2+]i. In contrast to insulin, thapsigargin, known to activate similar Cl− channels in HTC cells (Kilic & Fitz, 2002), produced a large transient increase in [Ca2+]i (Fig. 6B). Thus, activation of the Cl− channels by insulin does not require increases in [Ca2+]i.

Figure 6. Insulin and [Ca2+]i.

Figure 6

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Cells were preincubated with 20 μm Fluo-3 AM (30 min). After washing the dye, [Ca2+]i was determined by measuring Fluo-3 fluorescence. Representative measurements of [Ca2+]i in response to 10 μm insulin (A) and 20 μm thapsigargin (B) are shown. Dashed lines indicate [Ca2+]i of 100 nm. Similar results were obtained with insulin and thapsigargin in 11 and 12 cells, respectively.

Insulin resistance in muscle cells is associated with activation of the PKC activity (Shulman, 2000). Thus, we measured PKC activity after treatments with amiodarone or arachidonic acid utilizing fim-1 as previously described (Chen & Poenie, 1993). Fim-1 binds to various PKC isoforms, and when PKC translocates to the plasma membrane upon activation, plasma membrane fluorescence (rim fluorescence) increases (Chen & Poenie, 1993; Dupont et al. 2000). Representative images in Fig. 7A show that regions around the plasma membrane (arrows) are brighter in amiodarone-treated than in control cells. Initially, to ascertain that fim-1 stains PKC, HTC cells were stimulated with phorbol 12-myristate 13-acetate (PMA). PMA increased the rim fluorescence (Fig. 7B), suggesting that fim-1 is useful probe for monitoring PKC activity. Similar results were obtained with amiodarone or arachidonic acid treatments (Fig. 7B), indicating that excess of fatty acids in liver cells potently stimulates the PKC activity.

Figure 7. Fatty acids and PKC activity.

Figure 7

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A, cells were preincubated with 20 μm fim-1 AM for 30 min. Fluorescent images of a control cell and amiodarone-treated cell are shown. The plasma membrane is marked with arrows. Scale bar 5 μm. B, normalized rim fluorescence was measured under different conditions as indicated. Rim fluorescence was measured from a 1 μm wide rim around the plasma membrane, and then normalized to the total cellular fluorescence (see Methods). To stimulate PKC in control cells, PMA was added at 1 μm for 10 min in the incubation media. In other experiments, cells were treated overnight with amiodarone (50 μm) or arachidonic acid (15 μm). The number of cells was 8–38. Note that all treatments increased normalized rim fluorescence (P < 0.02, for all). C, [Ca2+]i was measured in response to 50 μm amiodarone at arrow. Dashed line indicates [Ca2+]i of 100 nm. Note that amiodarone produced only a small change in [Ca2+]i.

Previous studies in HTC cells demonstrated that activation of PKC activity is closely coupled with the increases in [Ca2+]i (Roman et al. 1998). Thus, in addition to fatty acids, amiodarone treatment could also stimulate the PKC activity by increasing [Ca2+]i. To assess this possibility, [Ca2+]i was measured after acute exposure to amiodarone. A representative recording in Fig. 7C and the data presented in Table 1 show that amiodarone stimulates only a very small increase in [Ca2+]i (∼10 nm). Accordingly, basal levels of [Ca2+]i in amiodarone-treated cells were not significantly different than in control cells (Tables 1, P > 0.25). Thus, activation of PKC in amiodarone-treated cells is not associated with increases in [Ca2+]i.

Table 1.

Effect of amiodarone on [Ca2+]i. Basal [Ca2+]i was measured in HTC cells

Basal [Ca2+]i (nm) Δ[Ca2+]i (nm)
Control 91.9 ± 4.9 (n = 19) 10.7 ± 0.9 (n = 16)
Amiodarone treatment 94.4 ± 5.3 (n = 13) —
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The Δ[Ca2+]i is the change 5 min after acute exposure to 50 μm amiodarone. Basal [Ca2+]i was also measured after overnight treatment with amiodarone (50 μm). n is the number of cells.

To assess whether PKC activity contributes to the inhibition of insulin responses, exocytosis and conductance were measured after manipulation of the PKC activity. In a first set of experiments, PKC was stimulated acutely with PMA in control cells. Under basal conditions, PMA increased the rate of constitutive exocytosis to 3.5 ± 0.2% min−1 (44 cells), but had no effect on the conductance (17.9 ± 2.0 pS pF−1, 5 cells). Thus, PKC activation per se does not stimulate ion channel activity in HTC cells. Interestingly, exposure to PMA markedly inhibited the insulin response (Fig. 8), suggesting that PKC functions as an inhibitor of these responses. If PKC activation is responsible for defective insulin responses, then inhibition of PKC in amiodarone-treated cells would be expected to restore the insulin responses. To test this hypothesis, in a second set of experiments, insulin responses were measured in amiodarone-treated cells after treatment with the PKC inhibitors chelerythrine and calphostin C. The data summarized in Fig. 8 show that inhibition of the PKC activity led to a partial or full recovery of the exocytic responses, but had variable effects on the conductance responses. Specifically, chelerythrine recovered exocytosis to ∼75% and conductance to ∼130% of the control values. Calphostin C almost doubled the exocytic response but had no effect on the conductance response. These results suggest that increased PKC activity in insulin resistance may contribute to inhibition of the exocytic and conductance responses to insulin.

Figure 8. Insulin responses and PKC activity.

Figure 8

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Insulin-dependent exocytosis and the peak conductance were measured in control and amiodarone-treated HTC cells under different conditions, as indicated. Results were normalized to the values obtained in control cells. In control cells, PKC was stimulated by preincubating cells with 1 μm PMA for 10 min. The PKC activity in amiodarone-treated cells was inhibited by preincubation with 25 μm chelerythrine (Chel) or 0.5 μm calphostin C (Calph) for 15 min. Chelerythrine and calphostin C increased the rate of constitutive exocytosis in amidarone-treated cells to 3.8 ± 0.3% min−1 (10 cells) and 7.2 ± 0.9% min−1 (46 cells), respectively. Because PMA and PKC inhibitors increased the rates of constitutive exocytosis, these changes were subtracted from the measurements of insulin-dependent exocytosis. The number of cells was 5–46. Note that inhibition of the PKC with chelerythrine in amiodarone-treated cells partially recovers the insulin responses.

To examine the role of PKC in regulation of glucose release, the rates of gluconeogenesis were measured in control and amiodarone-treated cells after inhibition of PKC activity. Interestingly, in both control and insulin resistant cells, chelerythrine (25 μm, 30 min) alone increased the rates of glucose release by ∼200 nmol mg−1 h−1 (P < 0.003). Furthermore, chelerythrine had no effect on the suppression of glucose release by insulin (not shown), indicating that different mechanisms may be involved in the regulation of gluconeogenesis and Cl− channel opening by PKC.

Discussion

These studies provide support for the concept that accumulation of fatty acids in liver cells leads to the development of insulin resistance, and suggest that fatty acids per se are able to inhibit the insulin-dependent exocytosis and Cl− channel opening through stimulation of the PKC activity.

Overnight treatment with amiodarone markedly increased fatty acid content, and led to a potent inhibition of the insulin responses in liver cells. These effects were mimicked in control cells by exposures to high concentrations of the naturally occurring fatty acid arachidonic acid. Because amiodarone was not present during measurements, these results suggest that the effects of amiodarone may be mediated by fatty acids. Notably, amiodarone treatment produced a cellular phenotype of insulin resistance. The suppression of gluconeogenesis by insulin was markedly reduced. These findings are consistent with the previous studies, suggesting that accumulation of fatty acids may be an early event responsible for the development of insulin resistance in liver cells (Pagliassotti et al. 1996; Michael et al. 2000; Cho et al. 2001; Roden et al. 2001). It should be noted that amiodarone treatment has an advantage of allowing fat accumulation in isolated cells in the absence of changes in hormones, nutrients or other regulatory factors. Because it has been difficult to study insulin resistance in intact animals due to the complex systemic environment and associated changes in the levels of circulating hormones, amiodarone-treated liver cells may be useful for studying insulin resistance in single cells.

Recent studies demonstrated that insulin stimulates Cl− channel opening through exocytosis of a distinct pool of vesicles that contain Cl− channels in their membranes (Kilic et al. 2001b). Multiple lines of evidence suggest that opening of Cl− channels is directly involved in the regulation of cell metabolism and growth (Roman et al. 2001; Wondergem et al. 2001; Li & Weinman, 2002). Thus, it is attractive to speculate that defective exocytosis and associated Cl− channel opening may be critical events contributing to the development of insulin resistance in liver cells. These findings are intriguing in view of recent studies demonstrating that the activation of glucose uptake in muscle cells by insulin is mediated through exocytosis of a distinct pool of vesicles that contain glucose transporters GLUT4 in their membranes (Saltiel & Pessin, 2002). Notably, defective exocytosis of GLUT4-containing vesicles contributes to the inhibition of glucose uptake, and leads to insulin resistance in muscle cells (Minokoshi et al. 2003). While muscle and liver cells have different metabolic functions, exocytosis may be a common mechanism for insertion of the transport proteins into the plasma membrane in response to insulin, to meet physiological demands of these tissues.

Utilizing fim-1 as a probe, we demonstrated that the PKC activity is potently stimulated after treatments with amiodarone or arachidonic acid. These findings are consistent with other studies that demonstrate a link between fatty acids and PKC in different models of insulin resistance. For example, several PKC isoforms were stimulated in livers of the patients with type II diabetes mellitus (Considine et al. 1995). Similar results were found in primary hepatocytes obtained from insulin-resistant rats (Karasik et al. 1990; Griffin et al. 1999; Lam et al. 2002). Indeed, recent biochemical studies showed that fatty acids and their metabolites are capable of stimulating the PKC activity (Verkest et al. 1988; Khan et al. 1992).

These studies provide evidence that stimulation of the PKC activity is directly responsible for inhibition of the initial cellular responses to insulin. Activation of PKC by acute exposure to PMA, arachidonic acid or amiodarone treatment, all resulted in a potent inhibition of exocytosis and Cl− channel opening. Accordingly, inhibition of the PKC activity with chelerythrine led to a partial or full recovery of the insulin responses in amiodarone-treated cells. Interestingly, another PKC inhibitor, calphostin C doubled the exocytic response, but had no effect on the conductance response. This was a surprising result. In contrast to chelerythrine, which binds to a catalytic site on PKC and prevents the transfer of phosphate groups to target proteins, calphostin C binds to a regulatory site and prevents PKC translocation to the plasma membrane. Because PKC was already present at high concentrations at the membrane in these cells, we expected that calphostin C would have no effect. The dissociation between exocytic and conductance responses indicated that these processes may be regulated through different mechanisms. However, many biochemically unrelated manipulations inhibited both exocytosis and Cl− channel opening. These include PKC activation (PMA, these studies), PI 3-kinase inhibitors (wortmannin, LY294002), disruption of microtubule stability (nocodazole) (Kilic et al. 2001b); and tyrosine kinase inhibitors (genistein, authors' unpublished observations). These results suggested another explanation. Calphostin C may act through a PKC-independent mechanism (Zheng et al. 2004), and lead to recruitment of another pool of vesicles that can undergo exocytosis in response to insulin. These vesicles may be distinct from the vesicles containing Cl− channels, producing no net increase in the conductance, as observed in these studies.

The potential targets of PKC in liver cells are not known. Recent studies have demonstrated that Munc18 and SNAP25 are involved in modulation of exocytosis by the PKC in neuroendocrine cells (Nagy et al. 2002; Barclay et al. 2003). Munc18 and other proteins essential for exocytosis have been also found in hepatocytes (Fujita et al. 1998). However, in contrast to liver cells, PKC potentiates Ca2+-dependent exocytosis in neuroendocrine cells (Gillis et al. 1996; Zhu et al. 2002). Thus, it is not clear whether and how these proteins are involved in the insulin-dependent exocytosis. PKC may also inhibit insulin actions by inactivating insulin receptor β-subunit (IR-β) and insulin receptor substrate proteins (IRS), as previously described (Takayama et al. 1988; Yu et al. 2002). Activation of these proteins by insulin is an early event necessary for stimulation of insulin signalling pathways. Given that PKC isoforms can in principle interact with many different proteins to regulate a large variety of important cellular functions (Mochly Rosen, 1995), cellular mechanisms responsible for inhibition of the insulin-dependent exocytosis by PKC still need to be defined.

Assuming that these findings are relevant for the response of liver cells to insulin, two additional considerations merit emphasis. First, HTC cells express at least 10 different PKC isoforms (authors' unpublished observations). Thus, it is acknowledged that the specific PKC isoforms responsible for inhibition of the insulin responses are not known. Second, amiodarone was used here as a pharmacological tool to increase the intracellular fatty acid content. However, it is not known how and by which mechanisms amiodarone alters fatty acid metabolism. Recent studies in mouse hepatocytes demonstrated that amiodarone stimulates expression of the genes that are directly involved in fat metabolism (McCarthy et al. 2004). Other studies have shown that amiodarone induces fatty liver by inhibiting fatty acid oxidation and lipoprotein secretion (Letteron et al. 2003). Thus, multiple mechanisms may be involved in the amiodarone action. Interestingly, amiodarone is also widely used as an antiarrhythmic drug, and cellular mechanisms of its action in cardiac cells have been well documented. For example, amiodarone potently inhibits voltage-dependent ion channels in ventricular myocytes (Kodama et al. 1999). In endothelial cells, amiodarone induces vasodilatation through a sustained increase in [Ca2+]i levels (Grossmann et al. 2000). Because there is no evidence for the presence of voltage-dependent channels in liver cells (Graf & Haussinger, 1996), and amiodarone does not disrupt [Ca2+]i homeostasis (Fig. 7C, Table 1), it is unlikely that these mechanisms contribute to the effect of amiodarone in liver cells.

In summary, these studies demonstrate that the intracellular accumulation of fatty acids leads to the development of insulin resistance in liver cells. Notably, insulin-dependent exocytosis and Cl− channel opening are potently inhibited under these conditions. We also found that stimulation of the PKC activity is directly responsible for this inhibition. Thus, PKC isoforms may be potential targets for pharmacological modulation of the defective metabolism in insulin resistance through effects on exocytosis and Cl− channel opening.

Acknowledgments

This work was supported in part by National Institutes of Health grants (DK43278 and DK46082 to J.G.F., DK44716 and DK55386 to M.J.P), and the Waterman Foundation (J.G.F). G.K. is a recipient of American Liver Scholar Award.

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