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
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ABSTRACT |
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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
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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- (TNF-
) 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.
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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 M. 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 = 4/3 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 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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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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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.
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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.
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-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).
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.
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ACKNOWLEDGEMENTS |
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We thank Dr. Colin Nichols and Dr. François Vallette for critical reading of the manuscript and Y. Lecarpentier for help.
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FOOTNOTES |
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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.
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