Induction of apoptosis using sphingolipids activates a chloride current in Xenopus laevis oocytes

R. Souktani1, A. Berdeaux1, B. Ghaleh1, J. F. Giudicelli1, L. Guize2, J. Y. Le Heuzey2, and P. Henry1,2,3

1 Laboratoire de Pharmacologie, Faculté de Médecine Paris Sud 94275; 2 Service de Cardiologie A, Hôpital Broussais 75014; and 3 Service de Cardiologie, Hôpital Lariboisière, 75475 Paris, France


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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The purpose of this study was to investigate whether the cell shrinkage that occurs during apoptosis could be explained by a change of the activity in ion transport pathways. We tested whether sphingolipids, which are potent pro-apoptotic compounds, can activate ionic currents in Xenopus laevis oocytes. Apoptosis was characterized in our model by a decrease in cell volume, a loss of cell viability, and DNA cleavage. Oocytes were studied using voltage-clamp after injection with N,N-dimethyl-D-erythrosphingosine (DMS) or D-sphingosine (DS). DMS and DS activated a fast-activating, slowly inactivating, outwardly rectifying current, similar to ICl-swell, a swelling-induced chloride current. Lowering the extracellular chloride dramatically reduced the current, and the channel was more selective for thiocyanate and iodide (thiocyanate > iodide) than for chloride. The current was blocked by 5-nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB) and lanthanum but not by niflumic acid. Oocytes injected with a pseudosubstrate inhibitor of protein kinase C (PKC), PKC-(19-31), exhibited the same current. DMS-activated current was abolished by preexposure with phorbol myristate acetate. Our results suggest that induction of apoptosis in X. laevis oocytes, using sphingolipids or PKC inhibitors, activates a current similar to swelling-induced chloride current previously described in oocytes.

cell shrinkage; swelling-induced chloride channel; protein kinase C; deoxyribonucleic acid fragmentation


    INTRODUCTION
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INTRODUCTION
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APOPTOSIS CAN BE DISTINGUISHED from accidental cell death, also known as necrosis, by specific characteristics that include loss of cell volume, formation of apoptotic bodies, nuclear condensation, and internucleosomal DNA cleavage (for review, see Ref. 29). Numerous studies have described the biochemical characteristics of apoptosis, such as the enzymes involved in internucleosomal DNA cleavage and the morphological appearance of apoptotic bodies (29). However, relatively little is known about the shrinkage of cells, which occurs in all known examples of apoptosis and is a cardinal feature of the morphological description of apoptosis (2). This loss of cell volume during apoptosis contrasts with the events that occur during necrosis, in which the cell swells, loses its membrane integrity, and eventually ruptures, causing an inflammatory response.

Numerous mechanisms that might account for the cell volume loss observed in apoptosis have been suggested. DNA fragmentation is one possibility, but this phenomenon occurs late in the time course of apoptosis events, after cell shrinkage has begun (2). Changes in Na+/H+ exchanger and Na+-K+-2Cl- cotransport activity have been proposed during both induction and suppression of apoptosis (13). However, the exact mechanism of cell shrinkage remains unexplained. The observed transient rise in the buoyant density of apoptotic thymocytes suggests that there may be changes in the activity of ion transport pathways in the plasma membrane leading to a loss of intracellular water (32).

The purpose of the present study was to investigate modulations of ionic currents during apoptosis. Sphingolipids are well recognized as playing important roles in modulation of cell growth (16). Sphingolipid derivatives have been involved in a signal transduction pathway, via the inhibition of protein kinase C (PKC), which also mediates the effects of tumor necrosis factor-alpha (TNF-alpha ) and induces loss of cell viability attributed to programmed cell death (23). We investigated whether, N,N-dimethyl-D-erythrosphingosine (DMS), a potent pro-apoptotic compound, can modulate ionic currents in Xenopus laevis oocytes. The role of PKC was investigated, and induction of apoptosis after sphingolipids treatment was characterized in oocytes.


    MATERIALS AND METHODS
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MATERIALS AND METHODS
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Oocytes culture and testing. Ovaries of adult female X. laevis were surgically removed from tricaine-anesthetized frogs. They were dissected into small tissue sections and placed in solution containing 96 mM NaCl, 2 mM KCl, 5 mM HEPES, and 1 mM MgCl2, pH adjusted to 7.5 with NaOH (ND solution). Follicle cells were removed by incubating oocytes for 1-2 h with collagenase (1 mg/ml; Sigma, St. Louis, MO) dissolved in ND solution. Following treatment, the cells were extensively washed and then incubated in ND + CaCl2 (1.8 mM). Stage V-VI oocytes were selected for experimental use.

Intracellular injection. DMS and D-sphingosine (DS) were obtained from Sigma and were diluted in dimethyl sulfoxide (DMSO), and PKC-(19-31) was diluted in sterile water. The final concentration of DMSO was <0.1%. Control oocytes were injected with the same volume and concentration of DMSO diluted with sterile water. For intracellular injection, a micropipette was used, connected to an automated nitrogen-driven microinjector (Picospritzer II; General Valve). The micropipette was calibrated volumetrically by counting the number of injections needed to expel a known volume of solution. It was then introduced into the cell, and a 50-nl volume was administered corresponding to 17 ng of lipids or proteins per oocyte. The adequacy of injection was verified by observing the slight increase in cell size on injection.

Electrophysiological study. Three to 24 h after injection, a single oocyte was placed in a recording chamber filled with 3 ml of ND solution. Microelectrodes were pulled in one stage from 1.0-mm capillary glass (Clark, Pangbourne, UK) on a pipette puller (Bioscience). They were filled with 3 M KCl, and tip resistance was usually 1-3 MOmega . The cell was voltage-clamped using a two-microelectrode voltage-clamp amplifier (Geneclamp 500; Axon Instruments, Foster City, CA), connected to a homemade data acquisition system running on an 80386-based microcomputer. All experiments were conducted at room temperature. Current was measured as current flowing to ground through a low-resistance electrode containing 2% agarose in 3 M KCl. For reversal potential determination, oocytes were perfused at a constant rate (3-4 ml/min) with ND solution. The perfusate was then changed to a solution containing 98 mM KCl, 5 mM HEPES, and 1 mM MgCl2, pH adjusted to 7.5 with KOH (KD solution). One thousand milliseconds of hyperpolarizing and depolarizing ramps from -140 mV to +60 mV from a holding potential at 0 mV and 200-ms steps from -100 mV to +60 mV every 20 mV were administered in ND or KD solutions.

Pharmacological agents. 5-Nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB; 0.1 mM), niflumic acid (0.05 mM), and phorbol myristate acetate (PMA; 1 µM) were dissolved in DMSO. The final concentration of DMSO was always <0.1%. BaCl2 (1 mM), CaCl2 (1.8 mM), lanthanum chloride (1 mM), and EGTA (0.2 mM) were dissolved in ND solution. To investigate ionic selectivity of the channel, 96 mM NaCl was replaced by sodium gluconate, sodium iodide, and sodium thiocyanate in ND solution. These pharmacological agents were obtained from Sigma and added directly to the bath from freshly prepared stock solutions.

Assessment of cell shrinkage. Cell shrinkage was evaluated using a capillary glass pipette (internal diameter, 1 mm; external diameter, 1.5 mm). Twenty-four hours after injection, 30 oocytes were placed in the capillary glass and centrifuged for 2 min at 100 g. The level of each column was compared. The diameter of oocytes was also optically measured, and the volume was calculated assuming that each oocyte was a sphere (volume = 4/3pi R3, where R = radius). DMSO-, DMS-, and PKC-(19-31)-injected oocytes were analyzed with these methods.

DNA isolation and analysis of DNA fragmentation. Thirty oocytes were incubated in 1 ml of digestion buffer (10 mM NaCl, 10 mM Tris · HCl, 1 mM EDTA, 1.5 mM MgCl2, and 1% SDS, pH 8.0) containing 1 mg/ml proteinase K at 37°C for 30 min. After incubation, samples were cooled on ice and centrifuged at 5,000 g for 10 min at 4°C. The supernatant was then extracted with phenol:chloroform:isoamyl (25:24:1), precipitated in ethanol, and resuspended in TE buffer (10 mM Tris · HCl, 1 mM EDTA, pH 8.0). After RNase treatment (10 µg/ml), samples were reextracted with phenol:chloroform:isoamyl (25:24:1), precipitated in ethanol, and resuspended in water. DNA content was quantitated by reading the absorbance at 260 nm before analyzing by electrophoresis on a 2% agarose gel. A Hae III digest of phi X174 DNA (Genaxis Biotechnology) was applied to each gel to provide size markers of 1,353, 1,078, 872, 603, 310, 281, 271, 234, 194, and 118 base pairs, respectively. The DNA in gels was visualized under ultraviolet light after staining with ethidium bromide (Sigma).


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RESULTS
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Oocytes injected with DMS or DS exhibit a fast-activating, slowly inactivating, slightly rectifying current. The membrane potential of oocytes in ND solution determined 1 min after impalement by the measuring electrodes was -31 ± 6 mV (mean ± SE) for DMSO-injected oocytes and -23 ± 2 mV for DMS-injected oocytes. Oocytes injected with DMS exhibited a fast-activating, slowly inactivating, slightly rectifying current (Fig. 1, A and B). The peak inward current (-100 mV) was -10 ± 4 nA in DMSO-injected and -81 ± 73 nA in DMS-injected cells (mean of 10 cells, P < 0.05). The peak outward current (+60 mV) was 40 ± 16 nA in DMSO-injected and 310 ± 65 nA in DMS-injected cells (P < 0.01, Fig. 2A). In KD solution, the reversal potential was -14 mV, the peak inward current (-100 mV) was -26 ± 12 nA in DMSO and -191 ± 89 nA in DMS-injected cells (mean of 10 cells, P < 0.05), and the peak outward current (+60 mV) was 36 ± 7 nA in DMSO-injected and 435 ± 85 nA in DMS-injected cells (P < 0.05, Fig. 2A). When a ramp stimulation was repeated on the same cell during 10 min, the reversal potential gradually shifted from -21 mV to -31 mV (mean of 5 cells). The current was detectable 3 h after DMS injection and was maximal ~6 h after injection. Later on, the current decreased gradually (Fig. 2B). Forty-eight hours later, ~50% of oocytes injected with DMS, but less than 10% injected with DMSO, exhibited a vertical and straight current-voltage (I-V) relation curve in voltage clamp, suggesting leak current and death of oocytes.


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Fig. 1.   Injection of N,N-dimethyl-D-erythrosphingosine (DMS) activated a fast-activating, slowly inactivating, slightly rectifying current in Xenopus laevis oocytes. A: electrophysiological recording after stimulation of current in KD solution in dimethyl sulfoxide (DMSO, left) and DMS-injected oocytes (right) (pulsing protocol). B: average steady-state current-voltage relationship of DMSO and DMS-injected oocytes (mean value ± SD on 10 cells). DMS-injected oocytes exhibited a fast-activating, slowly inactivating, slightly rectifying current compared with DMSO-injected oocytes (KD solution). KD solution is 98 mM KCl, 5 mM HEPES, and 1 mM MgCl2, pH adjusted to 7.5 with KOH.



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Fig. 2.   DMS-induced current in ND and KD solutions and decrease of the current as a function of time. A: average steady-state current-voltage relationship of DMS-injected oocytes in 2 mM KCl-96 mM NaCl (DMS-ND) and 98 mM KCl-0 mM NaCl (DMS-KD) solution (mean value on 8 cells). B: after DMS injection, the current was maximal ~6 h after injection. After 24 and 36 h, the current decreased gradually (mean value on 6 cells for each period). ND solution is 96 mM NaCl, 2 mM KCl, 5 mM HEPES, and 1 mM MgCl2, pH adjusted to 7.5 with NaOH.

Given the shift in membrane potential and the shape of the current, it seemed likely that DMS activated a chloride-permeable conductance pathway similar to a swelling-induced chloride current, ICl-swell, previously described in oocytes (1). This was established by examining the effects of various extracellular chloride concentrations, anion permeabilities, and chloride channel blockers on the DMS-induced current.

Lowering the extracellular chloride from 100 mM to 52 and 4 mM by replacement of 48 and 96 mM NaCl with sodium gluconate in ND solution reduced the peak outward current (+60 mV) from 292 ± 71 to 192 ± 35 and 92 ± 21 nA, respectively (P < 0.05, Fig. 3A). The reversal potential was shifted by +13 mV toward 0 mV with the 52 mM extracellular chloride solution and +2 mV with the 4 mM extracellular chloride solution without any significant effect on the inward current (-65 ± 28 to -71 ± 28 nA; mean of 6 cells; Fig. 3A).


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Fig. 3.   Chloride sensitivity and anion permeability of the DMS-induced current. A: average steady-state current-voltage relationship of DMS-injected oocytes (mean value ± SD on 6 cells) before (solid circles) and after the extracellular chloride was lowered from 100 mM to 52 mM (gray circles) and 4 mM (open circles) by replacement of 48 and 96 mM NaCl with sodium gluconate in ND solution. B: average steady-state current-voltage relationship of DMS-injected oocytes (mean value on 8 cells) in ND (open circles) and after replacement of NaCl by sodium iodide (gray circles) and sodium thiocyanate (solid circles).

Replacement of NaCl (96 mM) with sodium iodide and sodium thiocyanate in ND solution increased the peak outward current (+60 mV) from 310 ± 26 to 401 ± 34 and 444 ± 31 nA (mean of 8 cells, P < 0.05) and the peak inward current (-100 mV) from -57 ± 29 to -61 ± 32 and -64 ± 34 nA, respectively (not significant, Fig. 3B). The reversal potential shifted from -23 mV in sodium chloride to -28 mV in sodium iodide and -29.5 mV in sodium thiocyanate.

DMS current was reversibly blocked by the carboxylate analog chloride channel blocker NPPB in a voltage-independent manner; 0.1 mM NPPB decreased the peak outward current from 465 ± 176 to 155 ± 94 nA. NPPB was equally effective in blocking the inward current (-178 ± 61 to -110 ± 60 nA; mean of 9 cells, P < 0.05, Fig. 4A). The DMS-induced current was irreversibly suppressed by the trivalent cation lanthanum; 1 mM lanthanum decreased the peak outward current from 357 ± 120 to 146 ± 86 nA (mean of 8 cells, P < 0.05) without any effect on inward current (Fig. 4B). It is important to note that neither NPPB nor lanthanum affected the basal current observed in DMSO-injected oocytes.


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Fig. 4.   Sensitivity of the current to chloride channels inhibitors. A: average steady-state current-voltage relationship of DMS-injected oocytes (mean value on 9 cells) before (solid circles) and after exposure (open circles) with 5-nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB, 0.1 mM) in ND solution. B: average steady-state current-voltage relationship of DMS-injected oocytes (mean value on 8 cells) before (solid circles) and after exposure (open circles) with lanthanum (1 mM) in ND solution.

We investigated whether the current could be a calcium-activated chloride current (ICl.Ca), despite a net difference regarding time dependence and shape. The irreversible block by 1 mM lanthanum clearly distinguish ICl-swell from ICl.Ca (1). The current displayed no requirement for extracellular calcium; oocytes bathed in a solution containing 0.2 mM EGTA plus 1 mM BaCl2 responded identically to oocytes bathed in a solution containing 1.8 mM CaCl2 (Fig. 5A). Niflumic acid (50 µM), a potent blocker of the ICl.Ca, did not significantly affect the DMS-induced current (422 ± 212 to 406 ± 212 nA on peak outward current; mean of 5 cells; Fig. 5B).


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Fig. 5.   DMS-induced current did not appear to be a calcium-sensitive chloride channel. A: the current displayed no requirement for extracellular calcium: oocytes bathed in a solution containing 0.2 mM EGTA plus 1 mM BaCl2 exhibited similar responses to oocytes bathed in a solution containing 1.8 mM CaCl2 (mean value on 6 cells). B: average steady-state current-voltage relationship of DMS-injected oocytes (mean value on 5 cells) before (solid circles) and after exposure (open circles) with niflumic acid (50 µM), an inhibitor of calcium-dependent chloride currents, in ND solution.

We tested another sphingolipid derivative, DS, which is also known to be a potent inductor of apoptosis. DS-injected oocytes exhibited a similar fast-activating, slowly inactivating, slightly rectifying current (Fig. 6A). The peak outward current (+60 mV) was 40 ± 16 nA in DMSO-injected and 194 ± 52 nA for DS-injected cells in KD solution. The peak inward current (-100 mV) was -10 ± 4 nA in DMSO-injected and -87 ± 36 nA in DS-injected cells (mean of 6 cells, P < 0.05).


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Fig. 6.   D-Erythrosphingosine (DS) and the protein kinase C (PKC) inhibitor PKC-(19-31) activated a similar current. A: average steady-state current-voltage relationship of DMSO and DS-injected oocytes (mean value ± SD on 6 cells). DS-injected oocytes exhibited a current similar to the DMS-injected oocytes. B: average steady-state current-voltage relationship of DMSO-, PKC-(19-31)-, and scrambled PKC-(19-31)-injected oocytes (mean value on 6 cells). PKC-(19-31) but not scrambled PKC-(19-31)-injected oocytes exhibited a current similar to the DMS-injected oocytes. C: average steady-state current-voltage relationship of DMS-injected oocytes with (PMA + DMS) and without (DMS) preexposure to 1 µM PMA during 15 min (mean value on 6 cells). Preexposure with PMA abolished DMS-activated current. PMA, phorbol myristate acetate.

PKC modulation of chloride current. It has been previously reported that DMS was cytotoxic for a variety of cell types acting as an inhibitor of PKC (16, 20). Thus we examined the role of PKC in the modulation of the DMS-induced chloride current. Oocytes injected with the pseudosubstrate inhibitor of PKC, PKC-(19-31) (15), exhibited the same fast-activating, slowly inactivating, slightly rectifying current as described after DMS injection (Fig. 6B). The peak outward current (+60 mV) was 31 ± 12 nA in DMSO-injected cells and 263 ± 89 nA in PKC-(19-31)-injected cells (mean of 6 cells, P < 0.05). The peak inward current (-100 mV) was -14 ± 6 nA and -59 ± 15 nA, respectively (P < 0.05). Oocytes injected with the scrambled peptide PKC-(19-31) did not exhibit any significant current compared with DMSO-injected oocytes (Fig. 6B). Before intracellular injection with DMS, oocytes were incubated during 15 min with PMA (1 µM), a powerful nonphysiological activator of PKC. Preexposure with PMA abolished DMS-activated current and did not induce any significant current compared with DMSO-injected oocytes (Fig. 6C). When the DMS-induced current was expressed in oocytes, adding PMA to the bath did not induce any significant changes in I-V curve (data not shown).

DNA fragmentation and death of oocytes. DNA agarose gel electrophoresis revealed that injection with DMS and PKC-(19-31) induced DNA fragmentation. DNA from noninjected cells, or injected with DMSO, remained unfragmented (Fig. 7).


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Fig. 7.   Injection of DMS and PKC-(19-31) in X. laevis oocytes induced apoptosis. Agarose gel electrophoresis of the DNA of noninjected cells (control), injected with DMSO, DMS, and PKC-(19-31). DNA extraction 24 h after injection. Lane 1: Hae III digest of phi X174 DNA (size markers of 1,353, 1,078, 872, 603, 310, 281, 271, 234, 194, and 118 base pairs). DNA agarose gel electrophoresis revealed that injection with DMS caused DNA fragmentation as well as PKC-(19-31). DNA from noninjected cells or cells injected with DMSO remained unfragmented.

Cell shrinkage. Twenty-four hours after injection, the oocytes injected with DMS and PKC-(19-31) clearly exhibited a loss in cell volume and folds in their membrane. The level of the column containing oocytes injected with DMS and PKC-(19-31) was significantly lower than those injected with DMSO compared with noninjected oocytes (-7.7 ± 1.4%, -7.0 ± 1.2%, and +0.3 ± 0.6%, respectively, compared with noninjected oocytes). In hypotonic medium (105 mosM) the level of the column was increased by +7.1% compared with control oocytes and was -6.1% in hypertonic medium (315 mosM). When the volume of oocytes was calculated as described in MATERIALS AND METHODS, we found a mean volume of 180 ± 19 µm3 in DMSO-injected oocytes and 150 ± 22 µm3 in DMS-injected oocytes, corresponding to a decrease in mean cell volume of -17% (30 measurements in each group). In hypotonic medium the change in cell volume was +24 ± 1.9% compared with control oocytes and was -18 ± 3.8% in hypertonic medium. Taking into account that the volume of oocytes is rather big and that folds can lead to an underestimation of the intracellular volume, the decrease in cell volume observed could correspond to a notable loss of water. DMS-injected oocytes did not exhibit any significant volume change in the presence of 1 mM lanthanum or 0.1 mM NPPB: -1.8 ± 2.2 and -2.4 ± 1.9% decrease in the level of the column; -3.6 ± 3.5 and -4.2 ± 2.7% for volume measurement, respectively.


    DISCUSSION
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The present study demonstrates that, in X. laevis eggs, injection of DMS and DS activates a chloride current and induces apoptosis probably via the inhibition of PKC activity.

Chloride channel similar to ICl-swell. Injection of DMS, DS, or PKC-(19-31) activates a fast-activating, slowly inactivating, slightly rectifying conductance that we characterized as a chloride conductance similar to the ICl-swell previously described in oocytes (1). The dependence on extracellular chloride concentration was clearly demonstrated, although the change in reversal potential (15 mV) was less than the 50 mV-shift predicted if the current was entirely chloride sensitive. Ackerman et al. (1) previously found such a discrepancy and proposed that it could be explained if gluconate is somewhat permeable through the channel and if intracellular chloride concentration decreases when a lower external chloride concentration is applied. Then the expected shift in the reversal potential for a chloride-selective conductance might be substantially <50 mV when lower external chloride concentration is applied. The present results are consistent with this hypothesis, since the change in reversal potential is notable when extracellular chloride concentration is decreased from 98 to 52 mM (13 mV) but only minor when extracellular chloride concentration is decreased from 52 to 4 mM (2 mV), i.e., when the extracellular chloride concentration is below the estimated intracellular concentration. Moreover, when a ramp stimulation protocol was repeated on the same cell, the shift of the reversal potential observed was consistent with a loss of intracellular chloride. This channel appears to be distinct from the calcium-activated chloride channel because of its lack of sensitivity to calcium, to niflumic acid, and block by lanthanum. (1). The channel displays an anion conductivity sequence of SCN- > I- > Cl-, and the current is blocked by lanthanum and NPPB. Regarding the I-V relationship upon changing sodium with potassium chloride, we cannot exclude a cationic contribution to the current. However, the presence of a cationic permeability is a common feature of several anion channels (1, 33). These data strongly suggest that this current correspond to a volume-regulated chloride channel similar to ICl-swell. Indeed, ICl-swell is known to be sensitive to volume or osmotic changes, but injection of the same volume of DMSO in each batch did not activate any significant current.

The transport systems most often activated by cell swelling are potassium or anion channels. Cloned chloride channels involved in cell volume regulation include the MDRI gene product, pICln, P-glycoprotein, ClC-2, and ClC-3, (for review, see Refs. 19 and 25). pICln and P-glycoprotein are no longer serious candidates (25). The following pore properties of ClC-2 are quite distinct from those found in our experiments: inward rectification is marked and the anion selectivity sequence is Cl- > I-. The ClC-3 hypothesis is viable since outward rectification and anion selectivity (I- > Cl-) appears to be similar to that found in our experiments. Moreover, Duan et al. (9) found that a serine residue in ClC-3 links phosphorylation-dephosphorylation to chloride regulation by cell volume. Our results are consistent with a regulation of the channel by PKC and therefore make the ClC-3 hypothesis the more relevant.

Sphingolipids as inductor of apoptosis. Ohta et al. (24) have shown that sphingosine and its methylated derivative, DMS, induce apoptosis in HL-60 human promyelocytic leukemia cells as did pharmacological inhibitors of PKC. In myeloid and lymphoid cells, ceramide analogs caused early, potent and specific internucleosomal DNA fragmentation (23). In the present study, 24 h after DMS or PKC-(19-31) injection, DNA exhibited a ladder on agarose gel electrophoresis, indicating internucleosomal DNA fragmentation, which is a characteristic feature of apoptosis (15); however, this fragmentation was not observed in control or DMSO-injected cells.

Sphinganine, sphingosine, and ceramide have been found to inhibit PKC (22). Recently, ceramide has been shown to selectively inhibit PKC-alpha translocation (20), demonstrating the existence of crosstalk between the two important lipid-mediated pathways of signal transduction. The role of PKC in apoptosis remains, however, unclear. In many cell systems, activation of PKC inhibits apoptosis (28), although stimulatory effects have also been reported (for review, see Ref. 6). The apoptotic capacity of PKC inhibitors, including DMS and DS, may involve cell cycle-related factors as downregulation of c-myc gene expression, which plays an important role in the regulation of both cell proliferation and apoptosis (12). Moreover, a number of potential targets for ceramide have been identified, which include a novel membrane-associated protein kinase (7), protein phosphatase (8), and the mitogen-activated protein kinase cascade (23).

Mechanism of cell shrinkage during apoptosis. One of the most ubiquitous and distinctive features of apoptosis is the loss of cell volume, which is common to all examples of apoptosis (for review, see Ref. 5). The few studies that have attempted to investigate the mechanisms behind apoptotic cell shrinkage have nearly all involved ionic transport. Studies examining the buoyant density change, which occurs in thymocytes undergoing apoptosis, found that the observed transient increase in cell density was in full agreement with the theoretical increase expected if the mechanism for cell shrinkage was based on the extrusion of water (2). Recent studies performed in a dexamethasone-induced apoptosis model in CEM-C7A lymphoblastoid cells have shown that cell shrinkage was accompanied by a net loss in intracellular K+, but without change in K+ efflux rate (3). Apoptosis in the L1210 B cell line is associated with a decrease in the Na+-K+-2Cl- cotransport activity (31). Two recent studies have involved the inhibition of Na+/H+ exchange in apoptosis (18, 26). Activation of Na+-K+-2Cl- cotransporter (10, 14) and Na+/H+ exchanger (13) are well-recognized mechanisms of electrolyte accumulation in shrunken cells. Conversely, inhibition of these two systems does not seem to be a crucial mechanism leading to cell shrinkage (for review, see Ref. 10).

Chloride channels have a well-defined role in cell volume regulation. An initial response to swelling is to reduce the intracellular concentration of K+ and Cl- so that water will flow out of the cell, allowing the volume to return to normal. The volume-regulated anion channels provide the trigger linking cell swelling to the loss of osmolytes (11).

A role of chloride currents in the proliferation and activation of lymphocytes has also been proposed (11). Recently, Szabo et al. (30) demonstrated that tyrosine kinase-mediated activation of outwardly rectifying chloride channel may play a role in CD95-induced apoptosis in T lymphocytes. Our data are consistent with this work, since we found that apoptosis induces the activation of a volume-regulated chloride channel via the inhibition of PKC. Moreover, the lack of cell volume decrease when oocytes are incubated with lanthanum or NPPB just after sphingolipids injection supports the hypothesis that the activation of a volume-regulated chloride current could participate to the cell shrinkage observed during apoptosis in X. laevis oocytes. However, our results cannot be generalized, since the volume regulation can be very different between cell types; it has been reported that several lymphoid cells (S49 Neo, CEM-C7, primary thymocytes) undergo apoptosis in response to hypertonic conditions, whereas several other cell types (L cells, COS, HeLa, GH3) do not (4).

Conclusion. The present results suggest that induction of apoptosis in X. laevis oocytes, using sphingolipids derivatives, activates a chloride current. Activation of this chloride current could contribute to the cell shrinkage observed during apoptosis in X. laevis oocytes. Further studies are needed to understand whether activation of this chloride current can be demonstrated in other cell types during apoptosis and whether it plays a crucial role during programmed cell death.


    ACKNOWLEDGEMENTS

We thank Dr. Colin Nichols and Dr. François Vallette for critical reading of the manuscript and Y. Lecarpentier for help.


    FOOTNOTES

This study was supported by grants from Fondation pour la Recherche Médicale and BQR, Faculté de Médecine Paris Sud (1998).

Address for reprint requests and other correspondence: P. Henry, Service de Cardiologie, Hôpital Lariboisière, 2 rue Ambroise Paré, 75475 Paris Cedex 10, France (E-mail: patrick.henry{at}lrb.ap-hop-paris.fr).

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

Received 9 November 1999; accepted in final form 30 December 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Ackerman, MJ, Wickman KD, and Clapham D. Hypotonicity activates a native chloride current in Xenopus oocytes. J Gen Physiol 103: 153-179, 1994[Abstract].

2.   Arends, MJ, and Wyllie AH. Apoptosis: mechanisms and roles in pathology. Int Rev Exp Pathol 32: 223-254, 1991[ISI][Medline].

3.   Benson, RSP, Heer S, Dive C, and Watson AJM Characterization of cell volume loss in CEM-C7A cells during dexamethasone-induced apoptosis. Am J Physiol Cell Physiol 270: C1190-C1203, 1996[Abstract/Free Full Text].

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