Plasma Membrane Potential in Thymocyte
Apoptosis
This information is current as
of June 16, 2017.
Bruno Dallaporta, Philippe Marchetti, Manuel A. de Pablo,
Carine Maisse, Huynh-Thien Duc, Didier Métivier, Naoufal
Zamzami, Maurice Geuskens and Guido Kroemer
J Immunol 1999; 162:6534-6542; ;
http://www.jimmunol.org/content/162/11/6534
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References
Plasma Membrane Potential in Thymocyte Apoptosis1
Bruno Dallaporta,* Philippe Marchetti,*† Manuel A. de Pablo,* Carine Maisse,*
Huynh-Thien Duc,‡ Didier Métivier,* Naoufal Zamzami,* Maurice Geuskens,§
and Guido Kroemer2*
A
poptosis may be defined as a cell death process in which
the activation of catabolic processes and enzymes occurs
before cytolysis, thereby facilitating the recognition, uptake, and digestion of the apoptotic cell by neighboring cells. For
a long time, it has been assumed that the activation of endonucleases and specific proteases (caspases) would not only constitute a
distinctive feature of apoptosis but also determine the decisive step
(the “decision to die” or “commitment point”) that distinguishes
living from dying cells. It is now clear that, in most models of
apoptosis, inhibition of nucleases and caspases does not prevent
cytolysis (1–5), indicating that the decision to die is made before
catabolic enzymes are activated and that the activation of such
enzymes can be a byproduct of the cell death process rather than
a regulatory event (reviewed in Refs. 6 and 7). Accordingly, current attempts to understand the decisive step of the apoptotic process are focusing on the mechanisms leading to caspase and endonuclease activation rather than on the action of these hydrolases.
We and others have shown that defective ion compartmentalization and membrane potentials may have a major impact on
apoptotic regulation (7–10). Lipophilic cations accumulate in the
*Centre National de Recherche Scientifique, Unité Propre de Recherche 420, Villejuif, France; †Institut National de la Santé et de la Recherche Médicale, Unit 459,
Lille, France; ‡Centre Hépatobiliaire de l’Hôpital Paul Brousse, Villejuif, France; and
§
Department of Molecular Biology, Université Libre de Bruxelles, Rhode-Saint-Genèse, Belgium
Received for publication December 21, 1998. Accepted for publication March 15,
1999.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This work was supported by Agence Nationale pour la Recherche sur le SIDA,
Association pour la Recherche contre le Cancer, Centre National de la Recherche
Scientifique, Fondation pour la Recherche Médicale, Ligue Française contre le Cancer, Institut National de la Santé et de la Recherche Médicale, and Programme de
Recherche Fondamentale en Microbiologie et Maladies Infectieuses et Parasitaires du
Ministère de la Recherche, Sidaction (to G.K.). M.G. is a senior research associate of
the Belgian National Fund for Scientific Research.
2
Address correspondence and reprint requests to Dr. Guido Kroemer, 19 rue Guy
Môquet, B.P. 8, F-94801 Villejuif, France. E-mail address: [email protected]
Copyright © 1999 by The American Association of Immunologists
mitochondria matrix, driven by the electrochemical gradient following the Nernst equation, according to which every 61.5-mV
increase in membrane potential (usually 120 –170 mV) corresponds to a 10-fold increase in cation concentration in mitochondria. Therefore, the concentration of such cations is 2–3 logs
higher in the mitochondrial matrix than in the cytosol (CS) (11).
During the process of apoptosis, cells reduce the mitochondrial
retention of such lipophilic cations, including the fluorescent dye
3,39-dihexyloxacarbocyanine iodide (DiOC6(3))3 (7, 8). Thus, the
inner mitochondrial transmembrane potential (DCm), which is
mainly a proton gradient, dissipates relatively early during apoptosis (12, 13) via a process that involves the Bcl-2-inhibited and
Bax-facilitated opening of the permeability transition (PT) pore
(14, 15). Opening of the PT pore (which allows for the free diffusion of solutes of #1500 Da on the inner mitochondrial membrane) causes an increase in mitochondrial matrix volume (16).
Because the surface of the outer mitochondrial membrane is
smaller than that of the inner membrane, the PT pore opening may
cause local mechanic disruption of the outer mitochondrial membrane (17, 18), with consequent release of soluble intermembrane
proteins (19, 20). Several of the mitochondrial intermembrane proteins released upon PT pore opening (e.g., cytochrome c, apoptosis-inducing factor, and procaspases-2, -3, and -9) are endowed
with the capacity to activate caspases or nucleases (21–25). As a
result, mitochondrial membrane permeabilization has been viewed
as a phenomenon that marks the “point-of-no-return” of the apoptotic process (6, 7, 12, 26, 27).
A further apoptosis-relevant alteration in ion compartmentalization affects plasma membrane K1 homeostasis. Under normal circumstances, the K1 concentration is much higher in the CS (100 –
3
Abbreviations used in this paper: DiOC6(3), 3,39-dihexyloxacarbocyanine iodide;
DEX, dexamethasone; TEA, tetraethylammonium; TPA, tetrapentylammonium;
CMXRos, chloromethyl-X-rosamine; DCm, mitochondrial transmembrane potential;
DCp, EC, extracellular; CS, cytosol; plasma membrane potential; PI, propidium iodide; Eth, ethidium; HE, hydroethidine; PBFI, benzofuran isophtalate; DEVD, AspGlu-Val-Asp; PT, permeability transition; ROS, reactive oxygen species; Z-VAD.fmk, N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone.
0022-1767/99/$02.00
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Apoptosis is accompanied by major changes in ion compartmentalization and transmembrane potentials. Thymocyte apoptosis is
characterized by an early dissipation of the mitochondrial transmembrane potential, with transient mitochondrial swelling and a
subsequent loss of plasma membrane potential (DCp) related to the loss of cytosolic K1, cellular shrinkage, and DNA fragmentation. Thus, a gross perturbation of DCp occurs at the postmitochondrial stage of apoptosis. Unexpectedly, we found that
blockade of plasma membrane K1 channels by tetrapentylammonium (TPA), which leads to a DCp collapse, can prevent the
thymocyte apoptosis induced by exposure to the glucocorticoid receptor agonist dexamethasone, the topoisomerase inhibitor
etoposide, g-irradiation, or ceramide. The TPA-mediated protective effect extends to all features of apoptosis, including dissipation
of the mitochondrial transmembrane potential, loss of cytosolic K1, phosphatidylserine exposure on the cell surface, chromatin
condensation, as well as caspase and endonuclease activation. In strict contrast, TPA is an ineffective inhibitor when cell death is
induced by the potassium ionophore valinomycin, the specific mitochondrial benzodiazepine ligand PK11195, or by primary
caspase activation by Fas/CD95 cross-linking. These results underline the importance of K1 channels for the regulation of some
but not all pathways leading to thymocyte apoptosis. The Journal of Immunology, 1999, 162: 6534 – 6542.
The Journal of Immunology
6535
140 mM) than in the extracellular (EC) fluid (#5 mM). A
continuous, low K1 efflux, via so-called K1 leakage channels,
accounts for the maintenance of the charge differences on the
plasma membrane, the plasma membrane potential (DCp), and
thus is vital for ion and volume homeostasis. A major decrease of
the intracellular K1 levels has been observed in several cell types
(S49, Jurkat, HL60, and thymocytes) undergoing apoptosis in response to a variety of different inducers (glucocorticoids, A23187,
anisomycin, thapsigargin, staurosporine, anti-Fas/CD95, UV irradiation, and etoposide) (10, 28, 29). Complete loss of the K1 gradient may be expected to have severe metabolic consequences and
to be accompanied by a loss of DCp, because K1 efflux (which
normally follows the concentration gradient) would be interrupted.
Moreover, a loss of intracellular K1 may be linked to cellular
shrinkage (10, 28), one of the hallmarks of apoptosis. Importantly
enough, K1 efflux is mandatory for endonuclease activation, given
that physiological intracellular K1 concentrations inhibit nuclear
endonucleases in both cells and cell-free systems of apoptosis (10,
29, 30). This has lead to the proposal that the derepression of
endonucleases due to low intracellular K1 concentrations may be
a decisive prerequisite for end-stage DNA fragmentation (29).
The mechanisms accounting for K1 efflux during the apoptotic
process have been elusive thus far. On theoretical grounds, this K1
efflux could be due to the activation of K1 channels and/or due to
the inhibition of active K1 transport systems such as K1/Na1
ATPase. Inhibition of K1/Na1 ATPase may be expected to result
in an increase in cellular volume, which is not found in apoptosis
and rather is a feature of necrosis (7). Therefore, we reasoned that
an increased activity of K1 channels rather than an inhibition of
active K1 transport might account for the loss of cellular K1, a
hypothesis that we have tested in this work using K1 channel
inhibitors. Because K1 efflux causes a local charge asymmetry on
the plasma membrane and is the major determining factor for DCp
(60 –70 mV in most cell types, negative inside), we wondered
whether the apoptosis-associated K1 loss would culminate in a
loss of DCp. Therefore, we determined the retention of DiOC6(3)
in the CS of cells undergoing apoptosis (which, based on the
Nernst equation, should reach a cytosolic concentration ;1 log
higher than that of the EC medium at a DCp of 60 mV) and comparatively monitored the DCp and the DCm in cells undergoing
apoptosis.
Like other cells, thymocytes possess voltage-dependent K1
channels that participate in sustaining the resting DCp, in regulating cell volume, and in enabling cellular activation processes (31).
Here, we addressed the functional impact of K1 channels, K1
depletion, and DCp breakdown on thymocyte apoptosis. Our results suggest that gross perturbations of K1 currents leading to
DCp collapse are only observed at a relatively late, postmitochondrial stage of apoptosis. However, inhibition of K1 channels may
prevent the apoptosis induced by some but not all inducers at a
premitochondrial stage. These data underscore the physiological
importance of plasma membrane ion gradients in the control of
apoptosis.
lowing apoptosis inhibitors were tested: the K1 channel blockers tetraethylammonium (TEA) and tetrapentylammonium (TPA) (200-mM standard
dose; Sigma), the glucocorticoid receptor antagonist RU38486 (10 mM;
kindly provided by Roussel-Hoechst-Marion), the broad-spectrum caspase
inhibitor N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VAD.fmk) (100-mM final concentration; stock 10 mM in ethanol; Bachem,
Basel, Switzerland), or the ligand of the adenine nucleotide translocator
bongkrekic acid (50-mM final concentration; kindly provided by Dr. Duine,
Delft University, Delft, The Netherlands).
Materials and Methods
Determination of Asp-Glu-Val-Aspase (DEVDase) activity of CS
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 2-ME (50 mM; Sigma, St. Louis, MO). The following cell death inducers were employed: the glucocorticoid receptor agonist dexamethasone (DEX) (final concentration 1 mM; Sigma), etoposide
(10 mM; Sigma), irradiation (10 Gy; RX30/55 irradiator, Gravaton Industries, Gosport, U.K.), C2 ceramide (50 mM; Sigma), valinomycin (100 mM;
Sigma), PK11195 (200 mM; Sigma), or an Ab specific for CD95/Apo-1/
Fas (clone 154000D, 500 ng/ml; PharMingen, San Diego, CA). The fol-
For the determination of the DCp, DiOC6(3) (at a final concentration of 40
nM, stock 40 mM in ethanol, and an excitation wavelength of 488 nm
(emission 529 nm); Molecular Probes, Eugene, OR) was employed (32).
Alternatively, we used 20 nM of chloromethyl-X-rosamine (CMXRos)
(fluorescence at 600 nm), as described previously (33). The fluorescence
intensity or emission spectrum of these dyes is not influenced by variations
in the pH (data not shown), within a pH range relevant for apoptosis regulation (pH 6.0 – 8.2) (9). To determine the contribution of the DCp, DiOC6
staining was performed in either complete medium, 140 mM NaCl supplemented with 2 mM glucose and 10 mM HEPES (pH 7.4), or 140 mM
KCl plus 2 mM glucose plus 10 mM HEPES (pH 7.4), as indicated. This
latter procedure leads to the collapse of the DCp (34). Nonviable cells were
excluded by simultaneous staining with propidium iodine (PI) (final concentration of 5 mM; stock 10 mM in DMSO; excitation at 488 nM, emission 620 nM; Molecular Probes). Loss of membrane integrity was determined by means of the vital dye ethidium (Eth) bromide (200 ng/ml; 5 min
at room temperature, excitation at 480 nm, emission 600 nm). An FITCannexin V conjugate (1/400 dilution; 1 mg/ml, 15 min at 4°C; Brand Applications, Maastricht, The Netherlands; 525 nm) with a high affinity for PS
(35, 36) was used for the assessment of aberrant PS exposure. Labeling
with FITC-annexin V was performed after the removal of FCS by washing
cells twice in HEPES buffer (10 mM HEPES-NaOH (pH 7.4), 150 mM
NaCl, 5 mM KCl, 1 mM MgCl2, and 1.8 mM CaCl2). The generation of
ROS was monitored with hydroethidine (HE) (final concentration of 2 mM;
stock 10 mM in DMSO, excitation at 488 nM, emission 620 nM; Molecular
Probes). For the determination of intracellular K1 levels, cells were loaded
for 15–30 min with cell-permeant benzofuran isophtalate (PBFI)-acetoxymethyl ester (final concentration of 2.5 mM; stock 500 mM in DMSO).
The resulting PBFI fluorescence was elicited at 360 nM and measured at
485 6 20 nm as described previously (29, 37). The frequency of hypoploid
cells was determined by ethanol fixation followed by staining with PI as
described previously (38) using an Epics Profile II Analyzer (Coulter, Hialeah, FL). All other stainings were analyzed using a FACSvantage cytofluorometer (Becton Dickinson, Mountain View, CA), which was also
used for cell sorting.
Confocal microscopy
Cells were stained with DiOC6(3) as described above and were examined
with a Leica TCS NT confocal microscope (Leica Microsystemes, Rueil
Malmaison, France) equipped with a 15-mW argon-krypton laser configured with an inverted Leica DM IRBE. The 488-nm line was used to excite
DiOC6(3). Each image consisted of the projection of 50 optical sections
performed at intervals of 200 nm in the z-axis. xy fields of 1024 3 1024
pixels were scanned using an oil Pl Apo 1003 (not applicable 5 1.4)
objective. Chats (0.2 3 0.2-mm size) were quantified, and images were
processed using the Leica TCS NT software (PowerScan module) and
printed on a Sony HD-8800 color printer.
A total of 100 ml of CS (1 3 107 cells/100 ml in CFS 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 DTT, and 10 mM
HEPES-NaOH, pH 7.4) supplemented with additional protease inhibitors:
1 mg/ml leupeptin, 1 mg/ml pepstatin A, 50 mg/ml antipain, and 10 mg/ml
chymopapain) were prepared by five freeze/thaw cycles in liquid nitrogen,
followed by centrifugation (1.5 3 105 g, 4°C, 1 h) as described previously
(39). The capacity of CS or purified caspase-3 to cleave the caspase-3
recognition site DEVD was determined using Ac-DEVD-amino-4-methylcoumarin (Bachem) as fluorogenic substrate (40).
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Induction and inhibition of apoptosis
Cytofluorometric determination of DCp, viability,
phosphatidylserine exposure, reactive oxygen species (ROS),
cytosolic K1, and DNA loss
6536
APOPTOSIS AND PLASMA MEMBRANE POTENTIAL
DNA fragmentation analysis and electron microscopy
DNA fragmentation (5 3 105 cells/lane) was determined by agarose gel
electrophoresis (32). Electron microscopy was performed on ultrathin sections of glutaraldehyde/osmiumtetroxide-fixed, Epon-embedded cells after
uranyl acetate and lead citrate staining as described previously (41).
Results and Discussion
Sequential loss of DCm and DCp in thymocyte apoptosis
FIGURE 1. Alterations of DCm and DCp induced by DEX or TPA. A,
Determination of DiOC6(3) incorporation into untreated thymocytes (control) or cells cultured in the presence of DEX (1 mM, 6 h) or TPA (200 mM,
2 h). Cells were stained with DiOC6(3) in the presence of a buffer containing either 140 mM NaCl or 140 mM KCl. Results are representative of
five independent experiments. B, Confocal image of DiOC6(3)-stained control thymocytes, DEX-treated cells (DiOC6(3)low population), or TPAtreated cells. Black, background intensity; red, 20 –100 arbitrary units of
fluorescence intensity; green, 100 –200 arbitrary units; and blue, .200 arbitrary units. C, Quantitation of signals measured by confocal microscopy.
The fluorescence intensity of selected areas of cells (mitochondrion-rich
cytoplasm, CS) treated as described in A as well as DiOC6(3) background
fluorescence (in the EC area) were measured by confocal microscopy as
described in Materials and Methods. Each graph contains curves for three
different representative cells obtained by moving the cursor measuring the
fluorescence intensity on different subcellular regions. Data are shown on
a linear scale of arbitrary units of fluorescence intensity for the EC medium, the CS, and mitochondrion-rich areas (Mito). Each point represents
a 0.2 3 0.2-mm area within the cell. This experiment has been repeated
once, yielding similar data.
apoptosis, have undergone shrinkage both of the cytoplasm and of
the mitochondria (Refs. 13, 33, and 44 and data not shown). In
conclusion, thymocytes undergoing apoptosis dissipate their DCm
and manifest a transient mitochondrial swelling before they lose DCp.
K1 channel blockade inhibits thymocyte apoptosis induced by
DEX and etoposide
The specific K1 channel blocker TPA causes a dose-dependent
loss of DCp without affecting DCm, which can be measured independently from DCp by assessing the ratio between cytosolic
and mitochondrial DiOC6(3) retention (Figs. 1 and 3A). This has
been shown by comparative DiOC6(3) staining in the presence or
absence of 140 mM NaCl or KCl (Figs. 1A and 3A) or by confocal
quantification of the DiOC6(3) concentration gradients on the
plasma and mitochondrial membranes (Fig. 1, B and C and Fig.
3C). The DCp decrease induced by TPA appears to be almost total,
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Lipophilic cations such as DiOC6(3) accumulate in the mitochondrial matrix, driven by the electrochemical gradient following the
Nernst equation. Therefore, the concentration of such cations is
2–3 logs higher in the mitochondrial matrix than in the CS (11).
Given that the resting DCp of lymphocytes is usually 60 –70 mV,
the cytosolic concentration is ;10-fold higher than in the incubation medium. This implies that two potentials (DCm plus DCp)
influence the cellular uptake of DiOC6 (3). To distinguish the relative contribution of DCmand DCp to DiOC6(3) uptake, thymocytes were stained with DiOC6(3) in the presence of either 140
mM NaCl (which is compatible with the maintenance of DCp) or
140 mM KCl (which abolishes DCp). As expected, staining thymocytes in the presence of high EC K1, which causes a DCp depolarization, induces a reduction in DiOC6(3) fluorescence (Fig. 1A). This
K1-dependent reduction is not seen when cells are treated with the
K1 channel blocker TPA, indicating that TPA-inhibitible, voltagegated K1 channels maintain the DCp in this cell type, as reported
previously (42). Moreover, no K1-mediated reduction is found for
DiOC6(3)low thymocytes stimulated with DEX, indicating that
such cells do not have DCp or DCm (Fig. 1A), a finding which is
in line with our previous observation that such cells have undergone massive K1 efflux (29) and thus must be unable to generate
DCp. As a result, it appears that both TPA and DEX cause a DCp
collapse, although both reagents affect DCm in a differential fashion.
This interpretation was confirmed using confocal analysis of
DiOC6(3)-stained control, DEX-, or TPA-treated cells. TPA selectively affects the cytosolic (but not the mitochondrial) accumulation of DiOC6(3). The gradient between the EC and the cellular
DiOC6(3)-dependent fluorescence disappears after preincubation
of cells with TPA (Fig. 1C). In contrast, DEX reduces both the
plasma membrane and the mitochondrial DiOC6(3) concentration
gradients (Fig. 1, B and C).
Previously, we have reported the existence of a minor (,5%)
DiOC6(3)intermediate population among preapoptotic thymocytes (4,
43). These cells represent an intermediate stage of the apoptotic
process because, once purified, they quickly become DiOC6(3)low
(44). To clarify the DCm/DCp status of these cells, DEX-treated
thymocytes were stained with DiOC6(3), followed by cytofluorometric separation of cells into DiOC6(3)high (I), DiOC6(3)intermediate
(II), and DiOC6(3)low (III) cells (Fig. 2B). These cells were then
restained for 15 min in NaCl (which would maintain the DCp) or
KCl (which would depolarize the DCp) with either DiOC6(3) (Fig.
2C) or CMXRos (Fig. 2D), a DC-sensitive dye emitting a wavelength other than DiOC6(3) (33). As expected, DiOC6(3)high cells
possess DCp, whereas DiOC6(3)low cells have a greatly reduced, if
any, DCp. DiOC6(3)intermediate cells treated with 140 mM KCl
lower their fluorescence upon restaining with DiOC6(3) or de novo
staining with CMXRos (Fig. 2, C and D), indicating that the cells
have undergone an at least partial DCm collapse; however, these
cells retain their DCp. Electron microscopy revealed that this
DiOC6(3)intermediate population lacks advanced chromatin condensation and cell shrinkage (Fig. 2F), yet manifests mitochondrial
matrix swelling compared with DiOC6(3)high cells, which have a
normal phenotype (Fig. 2E). The mitochondrial swelling of
DiOC6(3)intermediate cells must be transient, because DiOC6(3)low
cells, which represent a later stage of the process with full nuclear
The Journal of Immunology
6537
because the DiOC6 (3) retention of TPA-treated cells is not reduced by treatment with 2 mM ouabain (data not shown), an inhibitor of the Na/K ATPase that is specifically involved in the
maintenance of DCp yet dispensable for that of the inner mito-
chondrial membrane (45). Even upon prolonged incubation (4 h in
Fig. 3A), TPA does not cause thymocyte apoptosis, as assessed by
measuring nuclear hypoploidy (numbers in cycles in Fig. 3A). In
contrast, incubating cells with DEX during the same period readily
FIGURE 3. Effect of TPA on DiOC6(3) uptake. Thymocytes were cultured in the absence (A) or presence (B) of DEX. In addition, the indicated amount
of TPA or TEA was added to the culture medium. After 4 h of incubation, cells were stained with DiOC6(3) in complete culture medium (A and B). The
DiOC6(3) incorporation profiles obtained in the absence of TPA are shown as an internal control in each individual graph. Alternatively, the concentration
of DiOC6(3) within the mitochondrion-rich and mitochondria-free cytoplasm was measured by confocal microscopy as described in the legend to Fig. 1C.
The fluorescence intensity was measured for each subcellular region of 10 different cells. Data are expressed as mean values 6 SEM (n 5 10).
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FIGURE 2. Temporal sequence of DCm
and DCp collapse induced by DEX. Untreated cells (A) or DEX-treated (1 mM, 4 h)
thymocytes (B) were stained with DiOC6(3),
followed by cell sorting into DiOC6(3)high (I,
41% of total), DiOC6(3)intermediate (II, 3% of
total), or DiOC6(3)low (III, 23% of total) populations, as indicated by the windows in B.
After sorting, cells were washed in 140 mM
NaCl or 140 mM KCl, followed by restaining with DiOC6(3) (C) or de novo staining
with CMXRos (D) and reanalysis. This experiment was repeated three times, yielding
similar results. In addition, DiOC6(3)high (E)
or DiOC6(3)intermediate (F) cells were fixed
and subjected to electron microscopy. Mitochondria-rich areas of representative cells
are shown.
6538
APOPTOSIS AND PLASMA MEMBRANE POTENTIAL
induced a total collapse in DiOC6 (3) retention and nuclear apoptosis in ;40% of the cells. Unexpectedly, when thymocytes were
kept in culture in the continuous presence of both DEX and TPA
(Fig. 3B), we found that TPA treatment prevented the cells from
responding to DEX by advancing to the DiOC6(3)low phenotype
and caused them to lose their DCm (Fig. 3, B and C). Dose-response analyses revealed a strong correlation between the TPAmediated collapse of DCp and the prevention of DEX-induced
DCm dissipation (Fig. 3, A and B). Whereas TPA was an efficient
inhibitor of the DEX-triggered DCm collapse at doses as low as 20
mM, a dose also reported to prevent UV-induced cell shrinkage of
HL60 cells (28), another quaternary ammonium, TEA, which has
a lower affinity for K1 channels than TPA (46), had to be employed at millimolar concentrations to obtain a similar effect (Fig.
3, A and B). Again, this effect correlated with the induction of a
DCp collapse (Fig. 3, A and B).
In a further series of experiments, we addressed the question of
whether TPA would selectively affect some features of DEX-induced apoptosis or rather block the entire apoptotic program. As
shown in Fig. 4, we found that TPA prevented all characteristics of
apoptosis, including the mitochondrial hypergeneration of ROS
(measured by determining the oxidation of HE to Eth, Fig. 4A), the
exposure of phosphatidylserine on the plasma membrane surface
(measured by FITC-annexin V conjugates, Fig. 4B), cytosolic K1
efflux (measured with PBFI, Fig. 4C), and nuclear DNA loss
(quantitated by PI staining of ethanol-fixed cells, Fig. 4D). TPA
also prevents the activation of caspases capable of cleaving the
substrate DEVD (Fig. 5A), as well as oligonucleosomal DNA fragmentation (Fig. 5B). Electron microscopy confirmed that thymocytes treated with both TPA and DEX exhibited a normal morphology (Fig. 6). Similar data were obtained when the
topoisomerase type II inhibitor etoposide was used instead of DEX
(Fig. 4). The dose of TPA necessary for the blockade of nuclear
apoptosis was found to correlate with that causing a DCp collapse
(numbers in circles in Fig. 3, A and B). Thus, TPA can block the
thymocyte apoptosis induced by the glucocorticoid agonist DEX
or by the genotoxic agent etoposide.
FIGURE 5. Effect of TPA on caspase and endonuclease activation. Thymocytes treated as described in the legend to Fig. 4 were subjected to lysis and
determination of DEVD-aminotrifluoromethylcoumarin cleavage (A). Alternatively, DNA was extracted (5 3 105 cells/lane) and subjected to agarose gel
electrophoresis for the detection of oligonucleosomal DNA fragmentation (B).
Cells were cultured for 4 h with the indicated combination of DEX, etoposide,
anti-CD95, and/or TPA, followed by analysis of the indicated parameter.
Shaded boxes indicate addition of the corresponding substance.
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FIGURE 4. Cytofluorometric assessment of various apoptosis-associated parameters and their modulation by TPA. Thymocytes were cultured for 4 h
with DEX, etoposide, or a mAb specific for Fas/CD95 and/or TPA, followed by staining with DiOC6(3) plus HE (A), Eth bromide plus FITC-annexin V
(B), or PBFI (C). Alternatively, cells were permeabilized with ethanol, and the nuclear DNA content was determined with PI (D). Results are representative
of four independent experiments.
The Journal of Immunology
6539
K1 channel blockade blocks a specific rather than general
apoptosis pathway
To determine whether TPA would block DEX-induced apoptosis
at an early or a late stage, we performed time-course inhibition
experiments. RU38486, a glucocorticoid receptor antagonist, can
block DEX-induced thymocyte apoptosis (12, 41) when added
early after DEX (Fig. 7a). In contrast, the caspase inhibitor ZVAD.fmk prevents the advent of nuclear apoptosis at a later stage,
in accordance with the idea that caspase-dependent endonuclease
activation occurs after the commitment to apoptosis triggered via
glucocorticoid receptor occupation (4). Compared with Z-VAD.fmk, TPA has to be added at a relatively early stage to obtain a
suppression of DEX-induced apoptosis (Fig. 7a); this finding is in
line with the fact that TPA prevents all signs of DEX-induced
apoptosis, including the early mitochondrial changes (Figs. 3 and 4),
whereas Z-VAD.fmk blocks the process leading to DNA fragmentation after DCm dissipation and mitochondrial swelling (4, 43).
The above findings indicate that TPA acts at a relatively early
stage of the DEX-induced apoptotic process, suggesting that it
could interfere with the activation of the apoptotic machinery
rather than with the machinery itself. Therefore, we determined
whether TPA would have a vast spectrum of antiapoptotic action
or instead whether it would fail to prevent apoptosis induction by
some apoptosis triggers. We explored the effects of TPA on apoptosis induction by 1) ceramide, a proapoptotic second messenger
elicited by glucocorticoids and genotoxic stress (47, 48), 2) valinomycin, a K1 ionophore that dissipates cellular and mitochondrial ion gradients (49), 3) PK11195, a ligand of the peripheral
benzodiazepine receptor that facilitates the opening of the PT pore
(50), and 4) Fas/CD95 cross-linking, a manipulation leading to
primary caspase activation (51). TPA prevents the apoptosis induced by ceramide but has no inhibitory effect on the apoptosis
caused by valinomycin, PK11195 (Fig. 7b), or anti-Fas (Figs. 4, 5,
and 7b). Thus, TPA can prevent the cell death induced by DEX
and DNA damage after the stage of ceramide generation, yet remains ineffective when apoptosis is enforced by direct K1 depletion, PT pore opening, or caspase activation.
Concluding remarks
The present work demonstrates that, during thymocyte apoptosis,
gross alterations of plasma membrane function and structure occur
after signs of mitochondrial dysfunction have manifested (Fig. 8).
Thus, dissipation of the DCm and transient mitochondrial swelling
are observed before several changes affect the plasma membrane:
loss of the DCp (Figs. 1 and 2), reduction of intracellular K1 (29),
cell shrinkage, and loss of membrane asymmetry with aberrant
exposure of phosphatidylserine residues on the plasma membrane
surface (33). TPA collapses the DCp of thymocytes, indicating that
TPA-inhibitible, voltage-gated K1 channels generate this DCp.
However, TPA has no major toxic effects on thymocytes (Fig. 6),
does not cause a nonspecific permeabilization of membranes with
Ca21 influx (data not shown), and does not cause PS exposure
(Fig. 4B), indicating that DCp collapse by itself does not perturb
the distribution of lipids in the plasma membrane. Unexpectedly,
we found that the prevention of K1 efflux by TPA fully inhibits the
thymocyte apoptosis induced by a variety of stimuli that may be
p53-dependent (etoposide, g-irradiation) or p53-independent
(DEX, ceramide). These pathways have in common that they require de novo mRNA and protein synthesis (52) and that they are
inhibited by PT pore blockade (26, 41, 53) or the knock-out of
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FIGURE 6. Ultrastructure of thymocytes cultured with DEX and/or TPA. Cells treated as described in the legend to Fig. 4 or Fig. 5 were examined by
electron microscopy.
6540
Apaf-1 or caspase-9 (54, 55), two molecules which link mitochondrial damage to downstream events of apoptosis. TPA intercepts
the apoptotic pathway at the early, premitochondrial level, because
1) it prevents the early DCm dissipation and later mitochondrial
ROS generation induced by DEX or etoposide (Figs. 3B and 4A),
2) has no effect when added late (Fig. 7a) or when added to
DiOC6(3)intermediate cells that have already disrupted of their DCm
(data not shown), and 3) has no inhibitory effect when apoptosis is
enforced by PK11105 (Fig. 7b), an agent that acts directly on the
PT pore. Moreover, TPA fails to prevent the apoptosis induced by
valinomycin, a K1-specific ionophore (Fig. 7a). This latter negative result suggests that TPA exerts its antiapoptotic effect by virtue of is modulatory effect on K1 fluxes rather than via a yet-to-be
discovered nonspecific effect. TPA does not suppress the apoptosis
induced by anti-Fas/CD95 (Figs. 4 and 5), a pathway that causes
direct caspase activation without any need of protein synthesis,
Apaf-1, or caspase-9 (51, 54 –56). Hence, TPA acts on a specific
rather than on a general pathway of apoptosis (Fig. 8). Of note,
TPA does not prevent the K1 efflux triggered by anti-Fas/CD95
(Fig. 4C), indicating that other TPA-resistant mechanisms of K1
efflux can intervene in a late stage of apoptosis, at least in the
Fas/CD95 pathway. The exact mechanisms by which TPA prevents the early mitochondrial changes triggered by DEX, DNA
damage, or ceramide remain elusive. Potassium channel blockers
have been shown previously to inhibit lymphocyte activation processes (31, 57), and it may be possible that an analogous effect
underlies the apoptosis-inhibitory effect of TPA.
Several published reports suggest that the regulation of K1
fluxes has a major impact on apoptosis regulation. Valinomycin, a
K1-specific ionophore, is a potent inducer of apoptosis in many
cell types, including neurons (58), hepatoma cells (59), pre-B cells
(49), and thymocytes (Refs. 60 and 61 and Fig. 7b). K1 depletion
suffices to cause apoptosis in neurons (3, 62, 63), and K1 efflux
underlies a murine model of cerebellar cell death in which the
weaver mutation causes the constitutive activation of the G protein-regulated inward rectifier 2 K1 channel (64). In neocortical
cells, attenuating outward K1 current with TEA or elevated EC
K1 rescues the apoptosis induced by staurosporine, serum withdrawal (58), or b-amyloid (65, 66). Plasma membrane channels
mediating a K1 efflux can cause apoptosis in a variety of different
cell types. Thus, ATP-activated P2Z/P2X purinergic receptors in
macrophages simultaneously mediate pore formation, DCp collapse, and apoptosis (67). Similarly, b-amyloid (68) and the HIVencoded Vpr protein (69) can incorporate into plasma membranes,
thereby perturbing ion homeostasis and causing cell death. In strict
contrast to these data, several apoptosis-inducing agents have been
reported to inhibit K1 channels. This applies to ceramide, which
inhibits the voltage-gated potassium channels of T lymphocytes
via tyrosine kinases (70), 4-aminopuridine, which blocks an outward rectifier K1 channel (71), or the apoptogenic peptides Reaper
and Grim (72). It remains to be confirmed that these effects are
truly responsible for apoptosis induction by these agents.
At first glance, the relationship between the mitochondrial PT
pore opening and K1 efflux appears paradoxical. On one hand,
DCm breaks down before a major CS K1 loss and concomitant
DCp loss occur (Figs. 1 and 2 and Ref. 29). On the other hand, a
blockade of K1 channels can prevent the mitochondrial dysfunction required for DEX- or etoposide-mediated apoptosis (Figs. 3
and 4). As a possibility, subtle changes in K1 homeostasis occurring together with the PT pore opening may participate in a positive feed-forward loop, facilitating the apoptosis-related mitochondrial dysfunction. This possibility is suggested by the recent
finding that the PT pore possesses a low conductance mode of
opening that is K1-selective and thus may cause selective alterations of local K1 concentrations (73). Alternatively, K1 fluxes
might play a pleiotropic role at several levels of the apoptotic
cascade, in analogy to cytosolic free Ca21, which can function, in
the low micromolar range, as a facultative premitochondrial second messenger-signaling PT pore opening (74) and increase as an
obligate consequence of the PT pore opening to levels of .200
mM at a late, postmitochondrial stage of the apoptotic process (75,
76). Thus, subtle changes in K1 fluxes could be involved in the
early apoptotic induction phase, whereas a major K1 loss would
only occur during the late degradation phase of apoptosis. Irrespective of these theoretical possibilities, the present data underline the importance of K1 as an endogenous apoptosis modulator.
FIGURE 8. Stages of the common phase of thymocyte apoptosis and
effects of TPA on apoptosis induction. The diagram resumes the data contained in this paper, as well as information from References 29 and 33. The
difference between stages 2 and 3 is the capacity of enhanced HE3 Eth
conversion (low in stage 2, high in stage 3; Ref. 29). Note that TPA has the
capacity to prevent apoptosis induction by some but not all inducers.
Downloaded from http://www.jimmunol.org/ by guest on June 16, 2017
FIGURE 7. Time course and spectrum of apoptosis-inhibitory action of
TPA. A, Time course of apoptosis inhibition mediated by RU38480, TPA,
or Z-VAD.fmk. Thymocytes were cultured for 5 h in the presence of DEX.
The indicated agents were added 30, 60, 90, 120, 180, or 240 min after
initiation of the culture. Hypoploidy was assessed after 5 h, and the percentage of inhibition of this parameter was determined. Control values of
untreated thymocytes were 15 6 2%; values of thymocytes treated with
DEX only were 52 6 3%. B, TPA-mediated inhibition of apoptosis induced by a panel of different inducers. Thymocytes were cultured for 4 h
with the indicated apoptosis inducer in the presence or absence of TPA
(200 mM), and the frequency of subdiploid cells was measured. Results are
mean values of three independent determinations 6 SEM.
APOPTOSIS AND PLASMA MEMBRANE POTENTIAL
The Journal of Immunology
Acknowledgments
We thank Martial Flactif (Institut Federatif de Recherche 22, Lille, France)
for confocal microscopy.
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