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The Journal of Immunology, 1998, 160: 5605-5615.
Copyright © 1998 by The American Association of Immunologists

Potassium Leakage During the Apoptotic Degradation Phase1

Bruno Dallaporta, Tamara Hirsch, Santos A. Susin, Naoufal Zamzami, Nathanael Larochette, Catherine Brenner, Isabel Marzo and Guido Kroemer2

Centre National de Recherche Scientifique, Villejuif, France


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results and Discussion
 References
 
The subcellular compartmentalization of ions is perturbed during the process of apoptosis. In this work, we investigated the impact of K+ on the apoptotic process in thymocytes and T cell hybridoma cells. Irrespective of the death-inducing stimulus (glucocorticoids, topoisomerase inhibition, or Fas-crosslinking), a significant K+ outflow was observed during apoptosis, as determined on the single-cell level by means of the K+-sensitive fluorochrome, benzofuran isophtalate. This loss of cytosolic K+ only occurs in cells that have completely disrupted their inner mitochondrial transmembrane potential. Inhibition of this mitochondrial transmembrane potential loss by Bcl-2 or by specific inhibitors acting on the mitochondrial permeability transition pore (bongkrekic acid, cyclosporin A) prevents K+ leakage. K+ drops at the same stage at which cells expose phosphatidylserine residues on the outer leaflet of the membrane and reduce the levels of nonoxidized glutathione, but before they hyperproduce reactive oxygen species, undergo massive Ca2+ influx, shrink, and lyse. In a cell-free system of apoptosis, isolated nuclei exposed to the supernatant of mitochondria that have undergone permeability transition only manifest chromatinolysis when the K+ concentration is lowered from physiologic to apoptotic levels. Accordingly, massive DNA fragmentation causing subdiploidy is confined to cells that have undergone K+ leakage. Together, these data point to the step-wise acquisition of membrane dysfunction in apoptosis and indicate an important role for the disruption of normal K+ homeostasis in apoptotic degradation. Derepression of endonucleases due to low K+ concentrations may be a decisive prerequisite for end-stage DNA fragmentation.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results and Discussion
 References
 
The apoptotic process can be subdivided into at least three different phases: the initiation phase, which is "private" in the sense that it depends upon the apoptosis-inducing stimulus; the common effector stage, during which the "decision to die" is made; and the degradation phase, during which the cell proceeds beyond the point-of-no-return and acquires the morphologic and biochemical hallmarks of apoptosis (1, 2). Among these phases, the effector stage has received much attention because it is subject to multiple regulatory effects, including disease-relevant molecules of the Bcl-2 family (3, 4). Although some investigators tend to believe that all key players at the effector stage are proteins that interact among each other in a multiprotein complex or "apoptosome" (caspases, CED-4, cytochrome c, and members of the Bcl-2 family) (5, 6, 7, 8), other researchers include nonprotein structures among the regulators of apoptosis (3, 4). Recent studies indicate that members of the Bcl-2 family that are incorporated into artificial membranes may have channel-like activities (9, 10, 11). Moreover, the current consensus has established that the function of the inner and/or outer mitochondrial membrane is perturbed early during the apoptotic process, leading to the disruption of the submitochondrial compartmentalization of cytochrome c, apoptosis-inducing factor (AIF)3, and protons (3, 4, 5, 12, 13, 14). This mitochondrial alteration is due at least in part to the opening of the permeability transition (PT) pore, which is formed at the contact site between the inner and the outer mitochondrial membranes (3, 12, 15, 16). Other nonprotein-molecules that are hypothetically involved in the apoptotic process are reactive oxygen species (ROS), endogenous antioxidants such as glutathione (GSH), and Ca2+ (1, 2, 3, 4, 17, 18, 19). However, the universal implication of these agents in cell death has not been confirmed, and it appears that they act primarily at the level of degradation, which is the point-of-no-return of the death process, rather than at the effector stage.

Attempts to order the sequence of apoptosis-associated events are mainly based on multiparameter cytofluorometric analyses, the only methodology that allows for the determination of several parameters on a per-cell basis. Using this approach on thymocytes, primary lymphocytes, or lymphoid cell lines (CEM-C7, 2B4.11, etc.), as well as cancer cell lines, a common, death stimulus-independent sequence of apoptosis-associated events has been established (20, 21, 22, 23, 24, 25, 26, 27). After initial mitochondrial transmembrane potential ({Delta}{Psi}m) disruption (stage 1), cells reduce their content in nonoxidized GSH and aberrantly expose phosphatidylserine (PS) residues on the plasma membrane (stage 2). Subsequently, cells hyperproduce ROS, which leads to cardiolipin oxidation, and undergo massive Ca2+ influx (stage 3) shortly before the plasma membrane permeability is lost. Commitment to cell death has already occurred at stage 1. DNA fragmentation is initiated during stage 2 (20, 21, 22, 23, 24, 25, 26).

We and others have investigated the subcellular distribution of Ca2+ and protons during the apoptotic process (15, 20, 21, 22, 23, 24, 25, 26, 28, 29, 30, 31). Moreover, it has recently been shown that K+ extrudes from the cell during the apoptotic process (32, 33). Whereas Ca2+ is much more abundant in extracellular fluid than in the cytosol, K+ concentrates in the cytosol (physiologic concentration in mammalian cells is 100–140 mM) as compared with the extracellular medium (4–5 mM). The asymmetric distribution of K+ is vital for the maintenance of plasma membrane integrity and function. Manipulations of cytosolic K+ concentrations causing a decrease in K+, such as extracellular K+ depletion or the addition of K+-specific ionophores, can cause apoptosis (34, 35). In contrast, an extra supply of external K+ can prevent apoptosis, at least in neuronal cell lines (36, 37, 38).

Based on these premises, we decided to determine the role of K+ in several models of apoptosis. As shown in this work, a loss of cytosolic K+ occurs during programmed cell death and is located downstream of mitochondrial PT and caspase activation. Although this K+ loss becomes detectable relatively late during apoptosis, it appears crucial for the activation of endonucleases as suggested by data obtained in intact cells and cell-free systems. Collectively, our data establish that the disruption of K+ homeostasis constitutes an important step in the apoptotic degradation phase after commitment to death has occurred.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results and Discussion
 References
 
Cells and culture conditions

Thymocytes from female 4- to 8-wk-old BALB/c mice were cultured at 37°C with 5% CO2 in RPMI 1640 supplemented with 10% FCS, L-glutamine, antibiotics, and ß-mercaptoethanol (50 µM, Sigma, St. Louis, MO). Murine 2B4.11 T cell hybridoma cell lines that had been stably transfected with the SFFV.neo vector containing the human bcl-2 gene or the neomycin resistance gene only were a gift from J. Ashwell (National Institutes of Health, Bethesda, MD).

Induction and inhibition of apoptosis

The following cell death inducers were employed: the glucocorticoid receptor agonist dexamethasone (DEX) (at a final concentration 1 µM; Sigma), etoposide (10 µM), or an Ab specific for CD95/Fas (clone 154000D, 500 ng/ml; PharMingen, San Diego, CA). The following apoptosis inhibitors were tested: the antioxidant N-acetylcysteine (50 mM), the protein synthesis inhibitor cycloheximide (35 µM; Sigma), the IL-1ß-converting enzyme-like caspase inhibitor acetyl-Tyr-Val-Ala-Asp-chloromethylketone (100 µM), the broad spectrum caspase inhibitor N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VAD.fmk) (at a final concentration 100 µM; stocks in 10 mM DMSO; Bachem, Basel, Switzerland), the ligand of the adenine nucleotide translocator bongkrekic acid (BA) (at a final concentration of 50 µM; kindly provided by Dr. Duine, Delft University, Delft, The Netherlands), or cyclosporin A (CsA) (1 µM; Novartis, Basel, Switzerland).

Cytofluorometric determination of mitochondrial parameters, ROS, GSH levels, and cytosolic Ca2+ or K+

For the determination of the {Delta}{Psi}m, 3,3'-dihexyloxacarbocyanine iodide (DiOC6(3)) (at a final concentration of 40 nM; stock 40 µM in ethanol; excitation wave length of 488 nm, emission 529 nm; Molecular Probes, Eugene, OR) was used (39). The generation of ROS was monitored with hydroethidine (HE) (at a final concentration of 5 µM; stock 10 mM in DMSO; excitation wave length of 488 nM, emission 620 nM; Molecular Probes); the content in nonoxidized GSH was determined using monochlorobimane (at a final concentration of 50 µM; stock 100 mM in ethanol; excitation wave length of 488 nM; emission 620 nM). Ca2+ levels were measured using Fluo-3 acetoxymethyl ester (AM) (at a final concentration of 250 nM; stock 1 mM in DMSO; excitation wave length of 488 nM, emission 525 nM; Sigma) in calcium-free HBSS (Eurobio, Paris, France). After incubation in the presence of the indicated fluorochrome (15–30 min, 37°C), cells were kept on ice (60-min maximum) until analysis. For the determination of intracellular K+ levels, cells were loaded during the 15 to 30 min with cell-permeant potassium-binding benzofuran isophthalate (PBFI)-AM (at a final concentration of 2.5 µM; stock 500 µM in dimethylformamide) (40, 41, 42). The resulting PBFI fluorescence was elicited at 360 nM and measured at 485 ± 20 nm.

Cytofluorometric analyses

The frequency of hypoploid cells was determined by ethanol fixation followed by staining with propidium iodine (PI) as previously described (43) using an EPICS Profile II Analyzer (Coulter, Hialeah, FL). All other stainings were analyzed using a FacsVantage cytofluorometer (Becton Dickinson). Data were analyzed in duplicate, and results were recorded for 10,000 cells while gating was performed either on cells exhibiting normal forward scatter (FSC) and side scatter (SSC) characteristics or on the whole cell population (normal cells plus shrunken cells) as indicated. In control experiments, cells were labeled in the presence of the protonophore carbonyl cyanide m-chlorophenylhydrazone (CCCP) (at a final concentration of 100 µM; intermediate stock 10 mM in ethanol), the prooxidant ter-butylhydroperoxide (200 µM), the redox-cycling agent menadione (which causes the generation of superoxide anions; 1 mM; stock 1 M in water), the calcium ionophore A23187 (which allows for the influx of extracellular Ca2+; 500 nM; stock 500 µM in DMSO), or a combination of valinomycin (0.5 µM)/nigericin (2.5 µM) (which allows for the free distribution of K+) plus different concentrations of KCl/NaCl (with a final osmolarity of 300 mM) and 5 mM HEPES buffer (pH 7.4). Cell sorting was performed using an Elite II cytofluorometer (Coulter, Miami, FL).

Cytofluorometric separation of cells and determination of cellular K+ levels

Thymocytes stimulated for 4 h with DEX (1 µM) were simultaneously stained with DiOC6(3) and PBFI as described above and separated in a FACS (Coulter Elite II or Becton Dickinson FACS Vantage). Cells were sorted into complete culture medium that was kept on ice and analyzed within a maximum of 30 min (for potassium quantitation) or 120 min after separation (for reculture and reanalysis). After cytofluorometric separation, cells (~106/tube) were washed three times (600 x g, 4°C, 10 min) in 140 mM NaCl plus 10 mM HEPES (pH = 7.4) and stored as a pellet at -20°C. Cells were resuspended in 100 µl of H2O, subjected to three freeze-thaw cycles, centrifuged (5000 x g, 4°C, 15 min) and injected into an IL-243 flame spectrophotometer (Laboratory Instrumentation, Lexington, MA) according to standard procedures.

Cell-free system of nuclear apoptosis

Nuclei from HeLa cells were purified on a sucrose gradient (44) and conserved at -20°C in HNB freezing buffer (80 mM KCl, 20 mM NaCl, 250 mM sucrose, 5 mM EGTA, 1 mM DTT, 0.1 mM PMSF, and 10 mM PIPES, pH 7.4) plus 50% glycerol for up to 15 days. Mouse liver mitochondria (10 mg/ml protein) were incubated with atractyloside (5 mM) for 10 min at room temperature to induce PT and the release of AIF and cytochrome c and then ultracentrifuged (1.5 x 105 g, 1 h, 4°C) as previously described (12, 45). Nuclei (103 nuclei/µl) were cultured in the presence of mitochondrial supernatants (200 ng protein/µl) for 90 min at 37°C in cell-free system buffer (220 mM mannitol, 68 mM sucrose, 2 mM NaCl, 2.5 mM PO4H2K, 0.5 mM EGTA, 2 mM MgCl2, 5 mM pyruvate, 0.1 mM PMSF, 1 mM dithiothreitol, and 10 mM HEPES-NaOH; pH 7.2) supplemented with variable amounts of KCl. Nuclei were stained with PI (10 µg/ml; Sigma) for >=5 min at room temperature), and cytofluorometric analysis was subsequently performed in an EPICS Profile II Analyzer (Coulter) while gating the FSCs and SSCs on single-nucleus events (46). Alternatively, nuclei were stained with 4',6-diamidino-2-phenylindone or subjected to agarose gel electrophoresis as previously described (12). In control experiments, the N-benzyloxycarbonyl-Val-Ala-Asp-7-amino-4-methyltrifluoromethylcoumarin (Z-VAD.afc) (30 µM in CFS buffer at 37°C, 30 min; Enzyme Systems, Livermore, CA) cleaving activity of mitochondrial supernatants was assessed. Z-VAD.fmk, an inhibitor of AIF (45, 46), was used for control purposes at a final concentration of 100 µM.


   Results and Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussion
References
 
Cells undergoing apoptosis exhibit a reduction in cytosolic potassium levels

Upon loading with the K+-sensitive dye PBFI (40, 41, 42), thymocytes that undergo apoptosis in response to DEX exhibit a lower fluorescence as compared with untreated cells (Fig. 1Go). Control experiments revealed that the difference in PBFI-dependent fluorescence between viable (PI-excluding) DEX-treated cells and viable control thymocytes disappeared in the presence of a combination of the potassium ionophore valinomycin and the K+/H+ exchanger nigericin (Fig. 1GoA), which together allow for the free distribution of K+ between the cell and the extracellular environment (41). Thus, the difference in PBFI fluorescence is due to a difference in K+ concentration rather than in dye uptake. Calibration of K+-permeant cells with variable doses of extracellular K+ (Fig. 1Go, A and B) revealed that viable, PI-excluding cells, which retain PBFI, exhibit a low (0.6–0.8 logs) although consistent (n = 5) K+-dependent variation of fluorescence. Although this relatively low signal/background ratio would render this dye inappropriate for the quantitation of apoptosis in single parameter studies, it is possible to assess the loss of PBFI fluorescence in combination with additional fluorochromes (see below, Figs. 2Go-4). In the presence of K+ ionophores and external K+, all cells, including the fraction of DEX-treated thymocytes with an originally low PBFI-dependent fluorescence (PBFIlow cells), acquire a PBFIhigh phenotype. The reduction of K+-dependent PBFI fluorescence is detectable in a fraction of normal-sized cells before major volume loss occurs, at least according to the criterium of the FSC (Fig. 1GoC), and is also detectable before cells become permeant to PI (Fig. 1GoA). A gas spectroscopic method for K+ determination confirmed that K+ loss occurred during apoptosis (see below, Fig. 5Go). In conclusion, DEX-induced thymocyte apoptosis is accompanied by both a loss of intracellular K+ that occurs before cell shrinkage and a complete loss of plasma membrane barrier function.



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  		FIGURE 1. Reduction of cytosolic K+ levels in apoptosis. Thymocytes that had been cultured for 6 h in the presence of DEX (1 µM) or freshly prepared control cells were stained with PI and simultaneously loaded with the membrane permeant agent PBFI-AM for 30 min in the presence or absence of valinomycin/nigericin and/or extracellular K+. Both cells with normal SSC and FSC characteristics as well as cells bearing an apoptotic phenotype were gated. A, Double staining profiles. Numbers denote arbitrary units of mean fluorescence intensity (log scale) that were found in the adjacent gate. B, Histograms of K+-dependent PBFI fluorescence among viable (PI-) cells that were gated as in A. Only viable cells with normal SSC and FSC characteristics were analyzed. C, PBFI fluorescence as a function of cell size. PBFI fluorescence was plotted against the FSC. The results are representative of five independent experiments.

 


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  		FIGURE 2. Triple staining of cells for the simultaneous determination of {Delta}{Psi}m (fluorochrome: DiOC6(3)), ROS generation (HE->Eth conversion), and K+ (fluorochrome: PBFI). Thymocytes were cultured for 4 h in the presence or absence of DEX, stained with the three indicated fluorochromes as detailed in Materials and Methods, and subjected to cytofluorometric analyses. The PBFI-staining profiles of normal-sized cells with a DiOC6(3)high HE->Ethlow population (fraction I), DiOC6(3)intermediate HE->Ethlow population (fraction II), DiOC6(3)lowHE->Ethlow population (fraction III), or DiOC6(3)lowHE->Ethhigh population (fraction IV) are shown. Numbers refer to the percentage of cells found in each gate.

 


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  		FIGURE 5. Relationship between {Delta}{Psi}m disruption, K+ loss, and nuclear apoptosis. Thymocytes that had been treated with DEX (1 µM, 4 h) were stained with PBFI plus DiOC6(3) and separated cytofluorometrically into three populations: fraction I, DiOC6(3)highPBFIhigh cells; fraction II, DiOC6(3)intermediatePBFIhigh cells and DiOC6(3)lowPBFIlow cells (A). Only cells having a normal size were gated. Thereafter, sorted cells were subjected to the determination of total H2O-soluble K+ content using a gas spectroscopic method. Values are given as arithmetic means ± SEM (n = 10) (B). Alternatively, cells were cultured for 1 h at 4°C or 37°C in the presence or absence of 1 µM CsA; they were then relabeled with PBFI/DiOC6(3) and reanalyzed (C). The results are representative of five (B) and two independent determinations (C).

 
Kinetics of K+ changes and temporal relationship with other features of the apoptotic process

In most cell types, including thymocytes, the first cytofluorometrically detectable sign of apoptosis is a loss of {Delta}{Psi}m and the subsequent hyperproduction of ROS on the uncoupled respiratory chain (21, 22, 26, 28, 47). To determine the temporal relationship between these signs of apoptosis and K+ loss, we performed a multiparameter analysis taking advantage of the {Delta}{Psi}m-sensitive dye DiOC6(3) (green fluorescence), the ROS-dependent conversion of hydroethidine into ethidium (Eth) (red fluorescence), and PBFI as a K+-sensitive marker (blue fluorescence). As shown in Figure 2Go, phenotypically normal (DiOC6(3)high HE->Ethlow) cells are homogeneously PBFIhigh, whereas DiOC6(3)low HE->Ethhigh cells are PBFIlow. In contrast, cells of the intermediate DiOC6(3)low HE->Ethlow stage split into PBFIhigh and mostly PBFIlow cells, indicating that K+ loss occurs after {Delta}{Psi}m dissipation but before the acquisition of the HE->Ethhigh phenotype. Kinetic multiparameter analyses confirmed this interpretation. Cells undergoing apoptosis manifest the {Delta}{Psi}m disruption before K+ leakage. Of note, DiOC6(3)low but not DiOC6(3)intermediate cells emit low K+-dependent fluorescence (Fig. 3GoA). K+ loss exhibits a strong correlation with PS exposure, as determined with an FITC-annexin V conjugate. All PBFIhigh cells are FITC-annexin Vlow (Fig. 3GoB), suggesting that K+ loss occurs together with or shortly after PS exposure. In contrast, the PBFI fluorescence correlates to a lower degree with ROS generation and Ca2+ influx. PBFIlow cells are distributed between the HE->Ethhigh/Fluo-3high populations (among which all cells are PBFIlow) and a fraction of cells that still have a normal HE->Ethlow phenotype (Fig. 3GoC) and have not yet undergone the Fluo-3-detectable terminal Ca2+ influx (Fig. 3GoD). In conclusion, K+ loss occurs after the initial dissipation of the {Delta}{Psi}m and after PS exposure, but before cells hyperproduce ROS and accumulate Ca2+.



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  		FIGURE 3. Kinetic analysis of DEX-induced changes in apoptosis-associated changes. Cells were cultured during the indicated period in the presence of DEX or cultured for 4 h in its absence; cytofluorometric determinations of cytosolic K+ concentrations (A–D), {Delta}{Psi}m (A), PS exposure (B), ROS generation (C), and cytosolic Ca2+ (D) were subsequently performed. As internal controls, cells were treated for 30 min with the protonophore CCCP (100 µM) (A), the detergent digitonin (0.05%) (B), the ROS-generating agent menadione (1 mM) (C), or the Ca2+ ionophore A12387 (500 nM) (D) before staining. Numbers indicate the percentage of cells encountered in the adjacent gate. Data are shown for all cells (normal-sized plus shrunken cells) while excluding debris on the FSC and SSC. Double-negative cells in A, C, and D have a shrunken phenotype (data not shown). Numbers in dark circles refer to the percentage of hypoploid cells, as determined by PI staining of ethanol-permeabilized cells. Note that both menadione and the protonophore CCCP, but not the Ca2+ ionophore A12387, caused a rapid loss in PBFI fluorescence.

 
PBFI-detectable K+ loss is observed in response to various apoptosis inducers, namely DEX ( Figs. 1–4GoGoGoGo) and the topoisomerase type II inhibitor etoposide, and is also seen as a result of crosslinking of the Fas/Apo-1/CD95 Ag (Fig. 4Go). Whenever an inhibitor interrupts the proapoptotic signaling cascade, K+ loss is prevented. Thus, inhibiting protein synthesis with cycloheximide or with the addition of the thiol antioxidant N-acetylcysteine prevents DEX- and etoposide-induced {Delta}{Psi}m disruption, nuclear apoptosis, and K+ outflow (Fig. 4Go). However these agents have no effect on Fas-induced apoptosis in any of its manifestations at the levels of mitochondria, nuclei, and K+. In contrast, acetyl-Tyr-Val-Ala-Asp-chloromethylketone (an inhibitor of caspase-1, formerly called IL-1ß-converting enzyme) prevents caspase-1-dependent Fas-induced apoptosis and K+ loss but has no effect on DEX- and etoposide-induced apoptosis (Fig. 4Go). These data support the conclusion that apoptotic {Delta}{Psi}m disruption, nuclear apoptosis, and K+ outflow are functionally associated with each other.



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  		FIGURE 4. Functional association between apoptotic {Delta}{Psi}m disruption, K+ loss, and nuclear apoptosis. Thymocytes were incubated in the presence of the indicated apoptosis inducers and inhibitors during a 4-h culture period as described in Materials and Methods and stained with PBFI plus DiOC6(3). The percentage of cells in each gate is shown. Numbers in dark circles indicate the percentage of subdiploid cells. The results are representative of two to three different determinations.

 
Functional relationship between K+ outflow and mitochondrial features of apoptosis

As shown in Figures 2 through 5GoGoGoGo, K+ loss is always secondary to {Delta}{Psi}m disruption. No K+ loss is observed in cells whose {Delta}{Psi}m remains stable. Cytofluorometric separation of DiOC6(3)/PBFI-stained cells and flame spectroscopic determination of K+ revealed that DEX-treated DiOC6(3)highPBFIhigh cells contained 10.9 ± 3.6 nmol K+/106 cells (meon ± SEM, n = 10). This value is comparable with that of nontreated control cells. In contrast, DiOC6(3)lowPBFIlow cells contain only 3.1 ± 1.1 nmol K+/106 cells (n = 10; p < 0.001, paired Student’s t test) (Fig. 5Go, A and B). Since this determination has been performed on cells with a normal volume (i.e., cells with normal FSC and SSC parameters), it appears that the K+ concentration drops to less than one-third of the control value.

To further investigate the relationship between mitochondrial PT and K+ loss, cells were treated with two inhibitors of the mitochondrial megachannel, BA and CsA. BA is a ligand of the adenine nucleotide translocator that prevents {Delta}{Psi}m disruption in most models of apoptosis (12, 24, 27, 48) but that fails to protect mitochondria against Fas-elicited caspase-1 (46). This agent stabilizes the {Delta}{Psi}m of DEX- or etoposide-treated cells and simultaneously prevents the loss of K+-dependent PBFI fluorescence (Fig. 4Go, lower panels), suggesting that K+ loss is secondary to mitochondrial PT.

To confirm this hypothesis, cells were stained with DiOC6(3) and PBFI and then cytofluorometrically separated into DiOC6(3)highPBFIhigh, DiOC6(3)intermediatePBFIhigh, and DiOC6(3)lowPBFIlow cells (Fig. 5GoA). Upon culture at 37°C, a fraction of DiOC6(3)high cells became DiOC6(3)intermediate and a significant percentage of DiOC6(3)intermediate cells became DiOC6(3)low and concomitantly reduced PBFI fluorescence (Fig. 5GoC), thus corroborating the precursor product relationship (DiOC6(3)highPBFIhigh -> DiOC6(3)intermediatePBFIhigh -> DiOC6(3)lowPBFIlow) that was suggested by the kinetic analyses (Fig. 3Go). CsA, a transient inhibitor of mitochondrial PT which acts on matrix cyclophilin D (16, 49), was employed as a probe to assess the role of mitochondrial dysfunction in this system. CsA (1–10 µM, Fig. 5Go and data not shown) prevents the initial transition (DiOC6(3)highPBFIhigh -> DiOC6(3)intermediatePBFIhigh) but not the second one (DiOC6(3)intermediatePBFIhigh -> DiOC6(3)lowPBFIlow), suggesting that the CsA-suppressible PT is only responsible for the first phase of {Delta}{Psi}m disruption (Fig. 5GoC). The transfection-enforced overexpression of Bcl-2, an endogenous inhibitor of PT (12, 45, 50), also prevents both {Delta}{Psi}m dissipation and the extrusion of K+ ions (Fig. 6Go).



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  		FIGURE 6. Effect of Bcl-2 on K+ loss. 2B4.11 T cell hybridoma cell lines stably transfected with a SFFV.neo vector containing the human bcl-2 gene or the neomycin resistance gene only were treated for 12 h with 1 µM DEX or 10 µM etoposide and then stained with PBFI and DiOC6(3).

 
Taken together, these data indicate that mitochondrial PT is a prerequisite of K+ loss. However, PT itself is not the mechanism that accounts for K+ release.

Functional relationship between K+ loss and caspase activation

Since caspases play a major role in apoptosis, we evaluated the effect of caspase inhibition on cytofluorometrically detectable apoptotic changes. Z-VAD.fmk, a broad spectrum inhibitor of caspases and other proteases including AIF (45, 46), completely prevents Fas-induced apoptosis at the DiOC6(3)high stage, thus confirming the involvement of caspase cascades early during Fas-induced apoptosis (46, 51, 52). In contrast, Z-VAD.fmk arrests DEX- and etoposide-treated cells at a later stage of the process, namely the DiOC6(3)intermediate stage (Fig. 4Go). Thus, in support of previous results (30, 48, 53), caspase inhibitors fail to prevent mitochondrial PT. Instead, they arrest the apoptotic process in a postmitochondrial phase of the apoptotic process. Cells that have been exposed to DEX plus Z-VAD.fmk (or etoposide plus Z-VAD.fmk) tend to accumulate at the DiOC6(3)intermediate stage (Figs. 4Go and 7Go). At this Z-VAD.fmk-arrested stage, cells still maintain high concentrations of nonoxidized intracellular GSH (as detected by monochlorobimane), lack PS exposure, and remain PBFIhigh (Fig. 7Go).



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  		FIGURE 7. Effect of Z-VAD.fmk on DEX-induced {Delta}{Psi}m disruption, K+ loss, GSH depletion, and PS exposure. Thymocytes were cultured for 4 h in the presence or absence of DEX (1 µM) and/or Z-VAD.fmk (100 µM) and then stained with the indicated fluorescent dyes. The typical data of three independent experiments are shown.

 
This observation indicates that the quasisimultaneous acquisition of these features of apoptosis (complete {Delta}{Psi}m disruption, GSH depletion/oxidation, PS exposure, and K+ loss) requires the action of Z-VAD.fmk-inhibitable proteases. Moreover, these data imply that caspase activation occurs before the K+ loss. Accordingly, physiologic concentrations of K+ (100–140 mM) fail to inhibit the acetyl-Asp-Glu-Val-Asp-7-amino-4-methyl coumarin-cleaving protease activity that accumulates in the cytosol of DEX-treated thymocytes. Similarly, in the presence of 140 mM K+, recombinant human caspases-3, -6, and -7 retain 60 to 80% of their enzymatic activity (100% value at 140 mM NaCl) (data not shown) (32). Thus, caspases can act upstream of K+ loss in cells with a normal K+.

K+ loss is a prerequisite for endonuclease activation

In several models of apoptosis induction and inhibition, the K+ loss exhibits a strong correlation with nuclear DNA loss (Fig. 4Go). This correlation has been confirmed by cytofluorometric purification of DiOC6(3)highPBFIhigh, DiOC6(3)intermediatePBFIhigh, and DiOC6(3)lowPBFIlow cells (as Fig. 5GoA) and by the subsequent determination of the frequency of hypoploid cells. Only DiOC6(3)lowPBFIlow cells exhibit advanced DNA fragmentation (Fig. 8Go), suggesting the possibility that K+ may act as an endogenous regulator of apoptotic degradation. Since manipulations of potassium have pleiotropic and toxic effects on intact cells (Fig. 1Go) (54), we tested the effect of variable K+ concentrations on a cell-free system of apoptosis in which isolated HeLa nuclei are exposed to supernatants from mitochondria that have been treated with atractyloside, an inducer of PT leading to the release of AIF (12) and cytochrome c (55). Physiologic concentrations of KCl corresponding to the normal cytosolic K+ concentration (100–140 mM) inhibit chromatin condensation and DNA fragmentation (as measured on gels or as nuclear DNA loss) to background levels (Fig. 9Go, A and B). In contrast, K+ has a far less spectacular inhibitory effect on the Z-VAD.afc cleavage mediated by the mitochondrial supernatant (Fig. 9GoC). Thus, K+ inhibits endonuclease activation much more efficiently than proteolytic activity, causing Z-VAD.afc cleavage (Fig. 9Go, B and C). Lowering of the KCl concentration itself is not sufficient to activate endonucleases (Fig. 10GoB). However, it does facilitate their activation by mitochondrial supernatants. The apoptosis-inhibitory effect of KCl is dose dependent, with an approximate ED50 of 80 mM (Fig. 9GoB). As a consequence, K+ loses most of its suppressive effect on endonuclease activation at the concentration that is found in the viable PBFIlow population, about one-third of the physiologic concentration. K+ outflow is a prerequisite for apoptotic endonuclease activation.



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  		FIGURE 8. Relationship between K+ loss and nuclear DNA loss. After 4 h of culture in the presence of DEX, thymocytes were labeled with PBFI and DiOC6(3), separated on a FACS (as shown in Fig. 5GoA) (A), permeabilized with ethanol, and stained with PI to determine the DNA content (B). Note that only DiOC6(3)lowPBFIlow cells become hypoploid.

 


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  		FIGURE 9. Effect of K+ on a cell-free system of nuclear apoptosis. A, Isolated nuclei from HeLa cells were cultured with AIF in the presence or absence of 120 mM KCl as described in Material and Methods. Nuclei were stained with PI, and the frequency of hypoploid nuclei was determined by cytofluorometry. Alternatively, nuclei were stained with 4',6-diamidino-2-phenylindole or subjected to agarose gel electrophoresis. Fluorescence micrographs of representative nuclei are shown. Note the peripheral chromatin condensation and the translucid aspect of AIF-treated nuclei. B, Dose response effect of increasing K+ concentrations on the AIF effect on isolated nuclei. C, Effect of K+ on AIF-mediated cleavage of the fluorescent substrate Z-VAD.afc. For control purposes, AIF-containing mitochondrial supernatants were treated with Z-VAD.fmk (100 µM, 15 min) before they were added to nuclei or Z-VAD.afc.

 


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  		FIGURE 10. Stages of the common phase of thymocyte apoptosis. The diagram covers the data contained in this paper as well as information from References 20, 21, 23, 25, and 48.

 
Concluding remarks

The data reported in this work are compatible with a step-wise sequence of the common phase of apoptosis. The cytofluorometric manifestations of this sequence are summarized in Figure 10Go. It may be assumed that various proapoptotic signal transduction or damage pathways converge at the level of the mitochondrion, the organelle which determines the commencement of the common pathway of apoptosis (3, 4, 5, 12, 13, 14). One of the first events of the common phase of apoptosis is the disruption of mitochondrial membrane integrity, with the dissipation of the {Delta}{Psi}m and/or the release of proteins that are normally sequestered in the intermembrane space (cytochrome c, AIF) in a sequence that remains to be determined and that may be cell type- and/or inducer-specific (13, 14, 15, 20, 21, 22, 23, 24, 25, 26, 29, 30, 31, 46). Thus, the sequence of molecular and cellular events leading to apoptosis, as described here, applies to thymocytes and lymphoid cells but may be different in other cell types.

The dissipation of the inner membrane proton gradient (stage 1) is mediated by PT, a process that is inhibited by CsA, BA, and Bcl-2 ( Figs. 4–6GoGoGo) (3, 15, 16, 21, 24, 45, 49). After initial {Delta}{Psi}m disruption, cells are irreversibly committed to death and consequently have entered the effector phase of apoptosis (20, 30, 53). However, early after {Delta}{Psi}m disruption, cells still maintain a normal morphology (20) and lack major changes in K+/Ca2+ homeostasis ( Figs. 1–3GoGoGo), redox potentials (Figs. 2Go, 3Go, and 7Go), or plasma membrane structure (Figs. 1Go, 3Go, and 7Go) (stage 1, Fig. 10Go).

The activation of Z-VAD.fmk-inhibitable proteases, which are triggered as a consequence of mitochondrial changes (5, 12, 13, 14, 24, 26, 31, 45, 46), is rate-limiting for the manifestation of a subsequent block of alterations (stage 2, Fig. 10Go), namely PS exposure on the cell surface, loss of nonoxidized GSH, and K+ leakage (Figs. 4Go and 7Go). The functional relationship among these events is unknown. As a possibility, the rearrangement of those plasma membrane lipids that are linked to PS exposure might favor the extrusion of GSH and K+ (56). Alternatively, the inhibition of Na/K adenosine triphosphatase and/or the opening of K+ channels may be connected to redox regulation and/or plasma membrane function (37, 57). Thus, perturbation of redox metabolism by protonophores or menadione favors rapid K+ efflux (Fig. 3Go). However, the exact molecular mechanism of K+ loss that occurs during apoptosis remains elusive. The cytosolic accumulation of those apoptogenic activities (AIF, cytochrome, etc.) that are normally confined to mitochondria (12, 45, 46, 58) stimulates nuclear endonucleases that can only be activated in the context of a low K+ concentration (Fig. 9Go).

The progressive exhaustion of antioxidant defense subsequently causes the manifest (HE-detectable) hyperproduction of ROS, which entails massive Ca2+ influx into the cytosol (stage 3, Fig. 10Go) shortly before cells shrink, form apoptotic bodies, and lyse ( Figs. 1–3GoGoGo) (25).

As a caveat, it would be erroneous to assume that the pleiotropic molecules that are involved in the common pathway of apoptosis such as K+, Ca2+, ROS, or caspases only participate in the degradation phase of apoptosis. Indeed, it has been established that elevations in cytosolic Ca2+ and ROS can serve as facultative transmitters in particular (private) proapoptotic signal transduction pathways (reviewed in 3 . Similarly, caspases can be activated at the premitochondrial stage of apoptosis, for instance after Fas crosslinking (46, 51, 52), although this is not a general feature of apoptosis. By analogy, it is possible that the K+ efflux can participate in special cases of apoptosis initiation. Indeed, a reversible K+ loss has been observed early after IL-3 withdrawal in a murine hematopoietic cell line (59) and an outward K+ current may be decisive for neuronal apoptosis that is induced by staurosporine or serum deprivation (38). However, at least in the models that we have studied in this work, K+ release occurs at a relatively late stage of apoptosis, after commitment to cell death.

After submission of this work, several studies reporting the efflux of K+ from cells undergoing apoptosis have been published (32, 33, 60). These studies clearly establish the role of the K+ (and Na+) efflux as a mechanism leading to cytosolic H2O loss and consequent volume loss (32, 33, 60). Thus, as shown here (Fig. 1Go), all cells with a shrunken phenotype exhibit low K+-dependent PBFI fluorescence, and only a minor fraction of cells that still have a normal FSC demonstrate a PBFIlow phenotype (33). Moreover, in agreement with our data, a rise in the concentration of extracellular K+ preventing the drop in cytosolic K+ can prevent nuclear DNA fragmentation (32, 33). However, in contrast to our observations, Cidlowski and coworkers (32, 33) suggest that K+ is an endogenous inhibitor of caspase activation and has a primary role in the regulation of cell death. This interpretation is based on the observations that 1) the cytochrome c/deoxy ATP-driven activation of caspase-3 is inhibited by K+ (32), and that 2) exogenous K+ can prevent endonuclease activation both in cells and in cell-free systems of apoptosis. As shown here, the action of a Z-VAD.afc-specific enzyme that is thought to be responsible for caspase-3 activation (46) is not inhibited by K+ (Fig. 9Go). This may imply that during early apoptosis, when K+ is still high, caspase activation is triggered by factors other than cytochrome c that are present in mitochondrial supernatants (46). Clearly, in our cell-free system of nuclear apoptosis, in which isolated nuclei are mixed with mitochondrial supernatant in the absence of cytosol (46, 61), cytochrome c and deoxy ATP are inactive, because they require the addition of cytosolic proteins for caspase-3 and endonuclease activation (62). In intact cells, the inhibition of caspase activation by Z-VAD.fmk prevents the K+ efflux, suggesting that caspase activation occurs upstream of this efflux (Fig. 7Go). Since the inhibition of caspases by Z-VAD.fmk fails to prevent the initial {Delta}{Psi}m dissipation (Fig. 7Go) and the subsequent cytolysis (53), it appears that the commitment point which seals the fate of the cell is located upstream of caspase activation and consequently upstream of K+ efflux. Therefore, although K+ outflow is critical for the acquisition of several hallmarks of apoptotic morphology (cell shrinkage and DNA fragmentation), it is probably not important for the life/death/ decision, which is made before K+ drops and before caspases and endonucleases are activated (53). This observation applies to the models of thymocyte apoptosis studied in this work. At present we cannot exclude the possibility that K+ levels may play a role during the initiation or effector stages of apoptosis in other cell types or in response to other apoptosis inducers.

In conclusion, we show in this paper that a loss of cytosolic K+ occurs as a common event in the apoptotic degradation phase of thymocytes, after mitochondrial PT and caspase activation but before cell shrinkage and before the plasma membrane becomes permeable. K+ leakage is closely associated with the loss of plasma membrane asymmetry and facilitates endonuclease-mediated chromatin degradation. These findings emphasize the functional impact of ion gradient dissipation on the apoptotic process.


   Acknowledgments
 
We thank Dr. P.X. Petit (Centre National de Recherche Scientifique, Villejuif) for technical advice, Dr. F Leviel (Institut National de la Santé et de la Recherche Médicale Unit 356, Paris, France) for help in K+ determinations, and D. Métivier (Centre National de Recherche Scientifique, Villejuif) for cell sorting.


   Footnotes
 
1 This work was supported by Agence Nationale pour la Recherche sur le SIDA, Association pour la Recherche contre le Cancer, Centre Nationale de la Recherche Scientifique, Fondation de France, Fondation pour la Recherche Médicale, Ligue Française contre le Cancer, Institut National de la Santé et de la Recherche Médicale, North Atlantic Treaty Organization, Roussel Uclaf-Hoechst, Sidaction, and the French Ministry of Science (to G.K.). S.A.S. was the recipient of a fellowship from the European Union. I.M. was the recipient of a fellowship from the Spanish Ministry of Science. Back

2 Address correspondence and reprint requests to Dr. Guido Kroemer, CNRS-UPR420, 19 rue Guy Môquet, BP8, F-94801 Villejuif, France. E-mail address: Back

3 Abbreviations used in this paper: AIF, apoptosis-inducing factor; FSC, forward scatter; SSC, side scatter; BA, bongkrekic acid; DEX, dexamethasone; PS, phosphatidylserine; DiOC6(3), 3,3'-)dihexyloxacarbocyanine iodide; {Delta}{Psi}m, AM, acetoxymethyl ester; mitochondrial transmembrane potential; CsA, cyclosporin A; Eth, ethidium; GSH, glutathione; HE, hydroethidine; PI, propidium iodine; CCCP, carbonyl cyanide m-chlorophenylhydrazone; PBFI, potassium-binding; benzofuran isophthalate; PT, permeability transition; ROS, reactive oxygen species; Z-VAD.afc, N-benzyloxycarbonyl-Val-Ala-Asp-7-amino-4-methyltrifluoromethylcoumarin; Z-VAD.fmk, N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone. Back

Received for publication November 24, 1997. Accepted for publication February 3, 1998.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussion
 References
 

  1. Thompson, C. B.. 1995. Apoptosis in the pathogenesis and treatment of disease. Science 267:1456.[Abstract/Free Full Text]
  2. Kroemer, G., P. X. Petit, N. Zamzami, J.-L. Vayssière, B. Mignotte. 1995. The biochemistry of apoptosis. FASEB J. 9:1277.[Abstract]
  3. Kroemer, G.. 1997. The proto-oncogene Bcl-2 and its role in regulating apoptosis. Nat Med. 3:614.[Medline]
  4. Reed, J. C.. 1997. Double identity for proteins of the Bcl-2 family. Nature 387:773.[Medline]
  5. Liu, X., C. N. Kim, J. Yang, R. Jemmerson, X. Wang. 1996. Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86:147.[Medline]
  6. Spector, M. S., S. Desnoyers, D. J. Hoeppner, M. O. Hengartner. 1997. Interaction between the C. elegans cell-death regulators CED-9 and CED-4. Nature 385:653.[Medline]
  7. Chinnaiyan, A. M., K. O’Rourke, B. R. Lane, V. M. Dixit. 1997. Interaction of CED-4 with CED-3 and CED-9: a molecular frame for cell death. Science 275:1122.[Abstract/Free Full Text]
  8. Wu, D., H. D. Wallen, G. Nuñez. 1997. Interaction and regulation of subcellular localization of CED-4 by CED-9. Science 275:1126.[Abstract/Free Full Text]
  9. Minn, A. J., P. Vélez, S. L. Schendel, H. Liang, S. W. Muchmore, S. W. Fesik, M. Fill, C. B. Thompson. 1997. Bcl-XL forms an ion channel in synthetic lipid membranes. Nature 385:353.[Medline]
  10. Schendel, S., Z. Xie, M. O. Montal, S. Matsuyama, M. Montal, J. C. Reed. 1997. Channel formation by antiapoptotic protein Bcl-2. Proc. Natl. Acad. Sci. USA 94:5113.[Abstract/Free Full Text]
  11. Antonsson, B., F. Conti, A. Ciavatta, S. Montessuit, S. Lewis, I. Martinou, M. Bernasconi, A. Bernard, J.-J. Mermod, G. Mazzei, K. Maundrell, F. Gambale, R. Sadoui, J.-C. Martinou. 1997. Inhibition of Bax channel-forming activity by Bcl-2. Science 277:370.[Abstract/Free Full Text]
  12. Zamzami, N., S. A. Susin, P. Marchetti, T. Hirsch, I. Gomez-Monterrey, M. Castedo, G. Kroemer. 1996. Mitochondrial control of nuclear apoptosis. J. Exp. Med. 183:1533.[Abstract/Free Full Text]
  13. Yang, J., X. Liu, K. Bhalla, C. N. Kim, A. M. Ibrado, J. Cai, T.-I. Peng, D. P. Jones, X. Wang. 1997. Prevention of apoptosis by Bcl-2: release of cytochrome c from mitochondria blocked. Science 275:1129.[Abstract/Free Full Text]
  14. Kluck, R. M., E. Bossy-Wetzel, D. R. Green, D. D. Newmeyer. 1997. The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science 275:1132.[Abstract/Free Full Text]
  15. Pastorino, J. G., G. Simbula, K. Yamamoto, P. A. J. Glascott, R. J. Rothman, J. L. Farber. 1996. The cytotoxicity of tumor necrosis factor depends on induction of the mitochondrial permeability transition. J. Biol. Chem. 271:29792.[Abstract/Free Full Text]
  16. Bernardi, P.. 1996. The permeability transition pore: control points of a cyclosporin A-sensitive mitochondrial channel involved in cell death. Biochim. Biophys. Acta 1275:5.[Medline]
  17. McConkey, D. J., P. Hartzell, P. Nicotera, S. Orrenius. 1989. Calcium-activated DNA fragmentation kills immature thymocytes. FASEB J. 3:1843.[Abstract]
  18. McConkey, D. J.. 1996. Calcium-dependent, interleukin 1 beta-converting enzyme inhibitor-insensitive degradation of lamin B-1 and DNA fragmentation in isolated thymocyte nuclei. J. Biol. Chem. 271:22398.[Abstract/Free Full Text]
  19. Slater, A. F. G., C. Stefan, I. Nobel, D. J. van den Dobbelsteen, S. Orrenius. 1996. Intracellular redox changes during apoptosis. Cell Death Differ. 3:57.
  20. Zamzami, N., P. Marchetti, M. Castedo, C. Zanin, J.-L. Vayssière, P. X. Petit, G. Kroemer. 1995. Reduction in mitochondrial potential constitutes an early irreversible step of programmed lymphocyte death in vivo. J. Exp. Med. 181:1661.[Abstract/Free Full Text]
  21. Zamzami, N., P. Marchetti, M. Castedo, D. Decaudin, A. Macho, T. Hirsch, S. A. Susin, P. X. Petit, B. Mignotte, G. Kroemer. 1995. Sequential reduction of mitochondrial transmembrane potential and generation of reactive oxygen species in early programmed cell death. J. Exp. Med. 182:367.[Abstract/Free Full Text]
  22. Castedo, M., A. Macho, N. Zamzami, T. Hirsch, P. Marchetti, J. Uriel, G. Kroemer. 1995. Mitochondrial perturbations define lymphocytes undergoing apoptotic depletion in vivo. Eur. J. Immunol. 25:3277.[Medline]
  23. Castedo, M., T. Hirsch, S. A. Susin, N. Zamzami, P. Marchetti, A. Macho, G. Kroemer. 1996. Sequential acquisition of mitochondrial and plasma membrane alterations during early lymphocyte apoptosis. J. Immunol. 157:512.[Abstract]
  24. Marchetti, P., M. Castedo, S. A. Susin, N. Zamzami, T. Hirsch, A. Haeffner, F. Hirsch, M. Geuskens, G. Kroemer. 1996. Mitochondrial permeability transition is a central coordinating event of apoptosis. J. Exp. Med. 184:1155.[Abstract/Free Full Text]
  25. Macho, A., T. Hirsch, I. Marzo, P. Marchetti, B. Dallaporta, S. A. Susin, N. Zamzami, G. Kroemer. 1997. Glutathione depletion is an early and calcium elevation a late event of thymocyte apoptosis. J. Immunol. 158:4612.[Abstract]
  26. Backway, K. L., E. A. McCulloch, S. Chow, D. W. Hedley. 1997. Relationships between the mitochondrial permeability transition and oxidative stress during ara-C toxicity. Cancer Res. 57:2446.[Abstract/Free Full Text]
  27. Polyak, K., Y. Xia, J. L. Zweier, K. W. Kinzler, B. Vogelstein. 1997. A model for p53-induced apoptosis. Nature 389:300.[Medline]
  28. Petit, P. X., H. LeCoeur, E. Zorn, C. Dauguet, B. Mignotte, M. L. Gougeon. 1995. Alterations of mitochondrial structure and function are early events of dexamethasone-induced thymocyte apoptosis. J. Cell. Biol. 130:157.[Abstract/Free Full Text]
  29. Shimizu, S., Y. Eguchi, W. Kamiike, S. Waguri, Y. Uchiyama, H. Matsuda, Y. Tsujimoto. 1996. Bcl-2 blocks loss of mitochondrial membrane potential while ICE inhibitors act at a different step during inhibition of death induced by respiratory chain inhibitors. Oncogene 13:21.[Medline]
  30. Xiang, J., D. T. Chao, S. J. Korsmeyer. 1996. Bax-induced cell death may not require interleukin 1ß-converting enzyme-like proteases. Proc. Natl. Acad. Sci. USA 93:14559.[Abstract/Free Full Text]
  31. Boise, L. H., C. B. Thompson. 1997. Bcl-XL can inhibit apoptosis in cells that have undergone Fas-induced protease activation. Proc. Natl. Acad. Sci. USA 94:3759.[Abstract/Free Full Text]
  32. Hughes, F. M., C. D. Bortner, G. D. Purdy, J. A. Cidlowski. 1997. Intracellular K+ suppresses the activation of apoptosis in lymphocytes. J. Biol. Chem. 272:30567.[Abstract/Free Full Text]
  33. Bortner, C. D., F. M. Hughes, J. A. Cidlowski. 1997. A primary role for K+ and Na+ efflux in the activation of apoptosis. J. Biol. Chem. 272:32436.[Abstract/Free Full Text]
  34. D’mello, S. R., C. Galli, T. Ciotti, P. Calissano. 1993. Induction of apoptosis in cerebellar granule neurons by low potassium: inhibition of death by insulin-like growth factor I and cAMP. Proc. Natl. Acad. Sci. USA 90:10989.[Abstract/Free Full Text]
  35. Duke, R. C., R. Z. Witter, P. B. Nash, J. D.-E. Young, D. M. Ojcius. 1994. Cytolysis mediated by ionophores and pore-forming agents: role of intracellular calcium in apoptosis. FASEB J. 8:237.[Abstract]
  36. Galli, C., O. Meucci, A. Scorziello, T. M. Werge, P. Calissano, G. Schettini. 1995. Apoptosis in cerebellar granule cells is blocked by high KCl, forskolin, and IGF-1 through distinct mechanisms of action: the involvement of intracellular calcium and RNA synthesis. J. Neurosci. 15:1172.[Abstract]
  37. Mark, R. J., J. N. Keller, I. Kruman, M. P. Mattson. 1997. Basic FGF attenuates amyloid ß-peptide-induced oxidative stress, mitochondrial dysfunction, and impairment of Na+/K+ ATPase activity in hippocampal neurons. Brain Res. 756:205.[Medline]
  38. Yu, S. P., C.-H. Yeh, S. L. Sensi, B. J. Gwag, L. M. T. Canzoniero, Z. S. Farhangrazi, H. S. Ying, M. Tian, L. L. Dugan, D. W. Choi. 1997. Mediation of neuronal apoptosis by enhancement of outward potassium current. Science 278:114.[Abstract/Free Full Text]
  39. Kroemer, G., B. Lisardo, P. Zamzami, S. Hortelano, C.-A. Martinez. 1997. Detection of apoptosis and apoptosis associated alterations. R. Lefkovitz, ed. The Immunology Methods Manual, Ch. 14.2 1111.-1125. Academic Press, San Diego.
  40. Minta, A., R. Y. Tsien. 1989. Fluorescent indicators for cytosolic sodium. J. Biol. Chem. 264:19449.[Abstract/Free Full Text]
  41. Kasner, S. E., M. B. Ganz. 1992. Regulation of intracellular potassium in mesangial cells: a fluorescence analysis using the dye, PBFI. Am. J. Physiol. 262:F462.[Abstract/Free Full Text]
  42. Meuwis, K., N. Boens, F. D. De Schryver, J. Gallay, M. Vincent. 1995. Photophysics of the fluorescent K+ indicator PBFI. Biophys. J. 68:2469.[Abstract/Free Full Text]
  43. Nicoletti, I., G. Migliorati, M. C. Pagliacci, C. Riccardi. 1991. A rapid simple method for measuring thymocyte apoptosis by propidium iodide staining and flow cytometry. J. Immunol. Methods 139:271.[Medline]
  44. Wood, E. R., W. C. Earnshaw. 1990. Mitotic chromatin condensation in vitro using somatic cell extracts and nuclei with variable levels of endogenous topoisomerase II. J. Cell Biol. 111:2839.[Abstract/Free Full Text]
  45. Susin, S. A., N. Zamzami, M. Castedo, T. Hirsch, P. Marchetti, A. Macho, E. Daugas, M. Geuskens, G. Kroemer. 1996. Bcl-2 inhibits the mitochondrial release of an apoptogenic protease. J. Exp. Med. 184:1331.[Abstract/Free Full Text]
  46. Susin, S. A., N. Zamzami, M. Castedo, E. Daugas, H.-G. Wang, S. Geley, F. Fassy, J. Reed, G. Kroemer. 1997. The central executioner of apoptosis: multiple links between protease activation and mitochondria in Fas/Apo-1/CD95- and ceramide-induced apoptosis. J. Exp. Med. 186:25.[Abstract/Free Full Text]
  47. Kroemer, G., N. Zamzami, S. A. Susin. 1997. Mitochondrial control of apoptosis. Immunol. Today 18:44.[Medline]
  48. Marchetti, P., T. Hirsch, N. Zamzami, M. Castedo, D. Decaudin, S. A. Susin, B. Masse, G. Kroemer. 1996. Mitochondrial permeability transition triggers lymphocyte apoptosis. J. Immunol. 157:4830.[Abstract]
  49. Zoratti, M., I. Szabò. 1995. The mitochondrial permeability transition. Biochem. Biophys. Acta 1241:139.[Medline]
  50. Decaudin, D., S. Geley, T. Hirsch, M. Castedo, P. Marchetti, A. Macho, R. Kofler, G. Kroemer. 1997. Bcl-2 and Bcl-XL antagonize the mitochondrial dysfunction preceding nuclear apoptosis induced by chemotherapeutic agents. Cancer Res. 57:62.[Abstract/Free Full Text]
  51. Los, M., M. Van de Craen, L. C. Penning, H. Schenk, M. Westendorp, P. A. Bauerle, W. Dröge, P. H. Krammer, W. Fiers, K. Schulze-Osthoff. 1995. Requirement of an ICE/CED-3 protease for Fas/Apo-1-mediated apoptosis. Nature 375:81.[Medline]
  52. Muzio, M., A. M. Chinnaiyan, F. C. Kischkel, K. O’Rourke, A. Shevchenko, J. Ni, C. Scaffidi, J. D. Bretz, M. Zhang, R. Gentz, M. Mann, P. H. Krammer, M. E. Peter, V. M. Dixit. 1996. FLICE, a novel FADD-homologous ICE/CED-3-like protease, is recruited to the CD95 (Fas/APO-1) death-inducing signaling complex. Cell 85:817.[Medline]
  53. Hirsch, T., P. Marchetti, S. A. Susin, B. Dallaporta, N. Zamzami, I. Marzo, M. Geuskens, G. Kroemer. 1997. The apoptosis-necrosis paradox: apoptogenic proteases activated after mitochondrial permeability transition determine the mode of cell death. Oncogene 15:1573.[Medline]
  54. Sun, D. Y., S. Jiang, L. M. Zheng, D. M. Ojcius, J. D. E. Young. 1994. Separate metabolic pathways leading to DNA fragmentation and apoptotic chromatin condensation. J. Exp. Med. 179:559.[Abstract/Free Full Text]
  55. Ellerby, H. M., S. J. Martin, L. M. Ellerby, S. S. Naiem, S. Rabizadeh, G. S. Salvese, C. A. Casiano, N. R. Cashman, D. R. Green, D. E. Bredesen. 1997. Establishment of a cell-free system of neuronal apoptosis: comparison of premitochondrial, mitochondrial, and postmitochondrial phases. J. Neurosci. 17:6165.[Abstract/Free Full Text]
  56. van den Dobbelsteen, D. J., C. S. I. Nobel, J. Schlegel, I. A. Cotgreave, S. Orrenius, A. F. G. Slater. 1996. Rapid and specific efflux of reduced glutathione during apoptosis induced by anti-Fas/APO-1 antibody. J. Biol. Chem. 271:15420.[Abstract/Free Full Text]
  57. Duprat, F., E. Guillemare, G. Romey, M. Fink, F. Lesage, M. Lazdunski. 1995. Susceptibility of cloned K+ channels to reactive oxygen species. Proc. Natl. Acad. Sci. USA 92:11796.[Abstract/Free Full Text]
  58. Liu, X., H. Zou, C. Slaughter, X. Wang. 1997. DFF, a heterodimeric protein that functions downstream of caspase-3 to trigger DNA fragmentation during apoptosis. Cell 89:175.[Medline]
  59. Laskay, G., T. Varhelyi, R. E. Daler, T. M. Dexter. 1995. Role of interleukin-3 in the regulation of intercellular K+. Biochem. Biophys. Res. Commun. 214:348.[Medline]
  60. McCarthy, J. V., T. G. Cotter. 1997. Cell shrinkage and apoptosis: a role for potassium and sodium ion efflux. Cell Death Differ. 4:756.
  61. Susin, S. A., N. Zamzami, N. Larochette, B. Dallaporta, I. Marzo, C. Brenner, T. Hirsch, P. X. Petit, M. Geuskens, G. Kroemer. 1997. A cytofluorometric assay of nuclear apoptosis induced in a cell-free system: application to ceramide-induced apoptosis. Exp. Cell Res. 236:397.[Medline]
  62. Zhou, H., W. J. Henzel, X. Liu, A. Lutschg, X. Wang. 1997. Apaf-1, a human protein homologous to C. elegans Ced-4, participates in cytochrome c-dependent activation of caspase-3. Cell 90:405.[Medline]



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