From the Instituto de Biología y
Genética Molecular, CSIC-Universidad de Valladolid, Facultad de
Medicina, Ramón y Cajal 7, 47005 Valladolid, Spain and the
¶ Department of Immunology and Oncology, Centro Nacional de
Biotecnología (CNB-CSIC), Campus de Cantoblanco,
28049 Madrid, Spain
Received for publication, November 7, 2002, and in revised form, January 30, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We address the specific role of cytoplasmic
Ca2+ overload as a cell death trigger by expressing a
receptor-operated specific Ca2+ channel, vanilloid receptor
subtype 1 (VR1), in Jurkat cells. Ca2+ uptake through the
VR1 channel, but not capacitative Ca2+ influx stimulated by
the muscarinic type 1 receptor, induced sustained intracellular
[Ca2+] rises, exposure of phosphatidylserine, and cell
death. Ca2+ influx was necessary and sufficient to induce
mitochondrial damage, as assessed by opening of the permeability
transition pore and collapse of the mitochondrial membrane
potential. Ca2+-induced cell death was inhibited by
ruthenium red, protonophore carbonyl cyanide
m-chlorophenylhydrazone, or cyclosporin A treatment, as
well as by Bcl-2 expression, indicating that this process
requires mitochondrial calcium uptake and permeability transition pore opening. Cell death occurred without caspase activation,
oligonucleosomal/50-kilobase pair DNA cleavage, or release of
cytochrome c or apoptosis inducer factor from mitochondria,
but it required oxidative/nitrative stress. Thus, Ca2+
influx triggers a distinct program of mitochondrial dysfunction leading
to paraptotic cell death, which does not fulfill the criteria for
either apoptosis or necrosis.
Release of mitochondrial intermembrane proteins to the cytosol is
a fundamental step in the cell death machinery during apoptosis and
necrosis (1). Among other triggers, massive calcium uptake into
isolated mitochondria induces the collapse of the mitochondrial membrane potential
( In the ischemic/excitotoxic stress model, the precise role of
Ca2+ overload in neuronal cell death is too complex to
trace (5). Stress stimuli often produce, in addition to
Ca2+ entry through the plasma membrane, other effects,
including (i) changes in membrane potential that can activate a variety
of ion channels, (ii) metabolic changes that may lead to free radical production, (iii) activation of kinases, and (iv) release of
Ca2+ from the intracellular Ca2+ stores.
Release of Ca2+ from the stores may be particularly
relevant, as Ca2+ depletion of the endoplasmic reticulum
(ER) blocks protein synthesis and is proapoptotic per se
(8, 9).
Apoptosis is induced through Ca2+-dependent
pathways in several cell types. This raises the question of whether
Ca2+ influx is sufficient for cell death. An apoptotic
pathway involving up-regulation of the FasL gene has been extensively
studied in T cells (10-12). This activation-induced cell death (AICD)
pathway can be triggered by receptors that stimulate phosphoinositide turnover, such as the T cell receptor or the human muscarinic type 1 receptor (HM1R), which produce an increase in
[Ca2+]c by inositol 1,4,5-trisphosphate-mediated
Ca2+ release from ER and capacitative Ca2+
entry. This is followed (>6 h) by calcium-dependent
up-regulation of FasL mRNA expression (12, 13), FasL binding to
Fas, and activation of caspases (14). Treatment with the
sarcoendoplasmic reticulum calcium ATPase blocker thapsigargin or with
Ca2+ ionophores induces cell death in T cells (8); however,
both treatments produce emptying of the ER (15, 16), which by itself can trigger apoptosis (9).
We address here the specific role of Ca2+ overload as a
cell death trigger by transiently expressing vanilloid receptor type 1 (VR1) (17) in Jurkat J-HM1-2.2 cells. VR1 is a receptor-operated Ca2+ channel, which is naturally expressed in some neurons
(17), peripheral blood T lymphocytes, and Jurkat cells (18, 19). Selective activation of VR1 with capsaicin allows for the control of
Ca2+ influx through the plasma membrane and the study of
specific effects of [Ca2+]c rises in our model.
Since the T cell line used in this study also expresses HM1R, the
effects of Ca2+ influx (through VR1) and Ca2+
release from the ER (through HM1R) can be directly compared. We find
that sustained Ca2+ entry though VR1 but not
Ca2+ release from ER induced by HM1R causes fast
mitochondrial damage with loss of Cell Cultures--
J-HM1-2.2 cells expressing the human HM1R
have been previously described (20). Cells were maintained in
Dulbecco's modified Eagle's medium (Bio-Whittaker Europe)
supplemented with 5% heat-inactivated fetal calf serum, 1 mM L-glutamine, 10 mM HEPES,
penicillin-streptomycin (100 units/ml and 100 µg/ml; Bio-Whittaker
Europe), and 0.5 mg/ml G418 as a selection medium for expression of the
HM1R.
Antibodies and Reagents--
The anti-Fas (clone CH-11)
monoclonal antibody was obtained from Medical and Biological
Laboratories Co., Ltd. Rabbit polyclonal Anti-AIF was a kind
gift from Dr. Susin (Institut Pasteur, Paris, France). Mouse monoclonal
anti-cytochrome C was from Pharmingen (clone 6H2.B4). Human polyclonal
anti-mitochondria Ab was from Dr. A. Serrano. Reagents used
include 8-methyl-N-vanillyl-6-nonenamida (capsaicin; Sigma),
carbonyl cyanide m-chlorophenylhydrazone (CCCP; Sigma), and
Z-Val-Ala-DL-Asp-fluoromethylketone (Z-VAD; Bachem AG).
Monoclonal antibody 12C4 against cytochrome oxidase subunit II was from
Molecular Probes (Eugene, OR), and AC15 monoclonal antibody
anti- Expression Vectors--
The pEF-GFP-VR1 plasmid was generated by
subcloning the VR1 cDNA into the plasmid pEF-GFPC1 (21) using the
SalI and KpnI restriction sites. For that
purpose, the VR1 cDNA was subcloned into pCDNA3 (17) and was
amplified by PCR with ECOTAQ Plus polymerase (Ecogen, Ltd.),
using the oligonucleotides VR1.1
(GGAATTCCGTCGACATGGAACAACGGGCTAGC) and VR1.2
(GCTCTAGAGGTACCGCGGCCGCTTATTTCTCC). The construction of
expression plasmids pEF-p35 and pEF human Bcl-2 (pEFhbcl-2) was
previously described (22). For some experiments involving measurement
of 2',7'-dichlorodihydrofluorescein (DCFH2) oxidation by
flow cytometry in the FL1 channel, VR1 cDNA was subcloned into the
pEF4/Myc-His B expression plasmid (pEF-VR1) (Invitrogen).
Transfection Assays--
For transfection, cells in logarithmic
growth were transfected with 20-30 µg of the pEF-GFP-VR1 plasmid by
electroporation at 270 V/975 microfarads using the Gene Pulser II
(Bio-Rad). Cells were analyzed 48 h after transfection. J-HM1-2.2
cells stably expressing baculoviral caspase inhibitor p35 were obtained
by transfecting J-HM1-2.2 cells with pEF-p35 plasmid and five rounds of selection with anti-Fas. These cells expressed similar levels of Fas
on the cell surface as compared with parental J-HM1-2.2 cells.
GFP-VR1 Expression Analysis--
The expression of the GFP-VR1
protein was analyzed by Western blot, flow cytometry, and fluorescence
microscopy. For the Western blot analysis, transfected cells were lysed
for 10 min in ice-cold radioimmune precipitation lysis buffer
supplemented with 0.1 mM phenylmethylsulfonyl fluoride. The
cell lysate was centrifuged at 15,500 × g for 5 min,
and the supernatant protein was electrophoresed (25 µg/lane) on a 5%
SDS-polyacrylamide gel and analyzed by Western blot (WB). For the flow
cytometry expression assays, cells were resuspended in PBS-propidium
iodide (PI; 200 ng/ml) and then analyzed (Coulter Epics XL-MCL). Data
analysis was performed with System II software (Coulter) by measuring
the green (505-545 mm; FL1) fluorescence in the cells excluding PI
(605-635 mm; FL3).
Intracellular Calcium Measurements--
Single cell measurements
of [Ca2+]c were performed by time-resolved
ratiometric digital imaging fluorescence microscopy in cells loaded
with the low affinity Ca2+ dye fura-4F. The dye was loaded
into the cells by incubation with 4 µM fura-4F/AM
(Molecular Probes, Inc., Eugene, OR) for 60 min at room temperature, in
standard solution containing 145 mM NaCl, 5 mM
KCl, 1 mM CaCl2, 1 mM
MgCl2, 10 mM glucose, and 10 mM
HEPES-sodium, pH 7.4. Cells were then washed with fresh medium,
resuspended at 10 × 106 cells/ml, and used for measurements.
Coverslips (12-mm diameter) were coated with fibronectin by incubation
(2 h, 37 °C) in PBS containing 20 µg/ml fibronectin (from human
plasma; Sigma), followed by incubation in PBS containing 1% bovine
serum albumin (1 h, 37 °C). Coverslips were mounted under the
microscope (Diaphot; Nikon) in a 37 °C chamber, and 105
cells loaded with fura-4F were attached to the coverslip during 10 min.
Test solutions were applied by continuous perfusion at 2-3 ml/min.
This allowed >95% exchange of the medium bathing the cells within
5-10 s. For fluorescence measurements, cells were alternately
epi-illuminated at 340 and 380 nm. Light emitted above 520 nm was
recorded by an extended ISIS-M camera (Photonic Science, Robertbridge,
East Sussex, UK) and analyzed using an Applied Imaging Magical image
processor (Sunderland, Tyne and Wear, UK) with 32-megabyte video RAM.
Sixteen video frames of each wavelength were averaged by hardware with
an overall time resolution of about 5 s for each pair of images at
alternate wavelengths. Consecutive frames obtained at 340- and 380-nm
excitation were rationed pixel by pixel, and the
[Ca2+]c was estimated using the following formula
(23): [Ca2+]c = Kd· Assessment of Apoptotic Cell Death--
Detection and
quantification of apoptosis at the single cell level was determined
using the In Situ Cell Death Detection Kit, TMR red (Roche Molecular
Biochemicals), which labels DNA strand brakes (TUNEL assay), following
the manufacturer's instructions, followed by flow cytometry analysis.
DNA degradation was determined by measuring the orange-red (555-600
mm; FL2) fluorescence in both GFP-VR1+ and
GFP-VR1
DNA degradation was determined by staining cellular DNA with PI,
followed by flow cytometry analysis. Briefly, transfected cells
(0.5-1 × 106) were fixed and permeabilized in 70%
ethanol (
For the determination of DNA fragmentation in oligonucleosomal
fragments (DNA laddering), total cellular DNA was extracted from
106 cells using the Easy-DNA kit (Invitrogen) following the
instructions provided by the manufacturer. Half of the DNA obtained was
loaded on 1.5% agarose gel and electrophoresed.
PS exposure was evaluated by staining the cells with annexin V-PE (BD
Pharmingen) and 7-aminoactinomycin D (7-AAD; Sigma) following the
instructions given by the manufacturer. Calcium-free Dulbecco's
modified Eagle's medium supplemented with 0.2 mM EGTA to
chelate calcium from fetal calf serum and the same medium plus 2 mM CaCl2 were used in experiments that required
the absence of extracellular calcium. Briefly, cells treated in
complete Dulbecco's modified Eagle's medium (0.1 µM
capsaicin unless otherwise indicated) were collected by centrifugation,
washed once in PBS, and incubated for 5 min with annexin V-PE in the
appropriate binding buffer. Cells were then incubated for 5 min with
7-AAD (1 µg/ml) and analyzed by flow cytometry. GFP-VR1+
cells and GFP-VR1 Active Caspase-3--
Active caspase-3 in cells transfected with
the pEF-GFP-VR1 plasmid was detected using the PE-conjugated polyclonal
active caspase-3 antibody apoptosis kit 1 (BD Pharmingen), followed by flow cytometry analysis of the orange-red (555-600 mm; FL2)
fluorescence in the GFP-VR1+ and GFP-VR1 Western Blot Analysis--
Total cellular protein was separated
by SDS-PAGE under reducing conditions and transferred to nitrocellulose
membranes (HybondTM ECLTM; Amersham
Biosciences). Membranes were blocked overnight with 5% nonfat dry milk
in PBS buffer. Subsequent antibody incubations and membrane washes were
performed in 0.5% nonfat dry milk in PBS, 0.2% Tween 20 buffer. The
blot was developed with peroxidase-conjugated anti-rabbit or anti-mouse
antibodies using ECL reagents.
Analysis of Changes in Cytochrome c and AIF Release--
Release of cytochrome
c and AIF from the mitochondria to the cytosol in sorted
GFP-VR1+ and GFP VR1 Immunofluorescence--
Transfected cells (106) were
washed in PBS, fixed in 4% paraformaldehyde for 15 min at room
temperature, preincubated in 5% bovine serum albumin, and incubated
for 1 h with primary antibodies (anti-mitochondria,
anti-cytochrome c, anti-AIF) in PBS containing 0.5% bovine
serum albumin and 0.1% Triton X-100. Cells were washed three times
with the same buffer and incubated for 1 h with goat Fab2
anti-human Cy3, goat Fab2 anti-mouse Cy5, or goat Fab2 anti-rabbit Cy5
(Jackson Immunoresearch Inc.), respectively. After antibody staining,
cells were labeled with 4',6-diamidino-2-phenylindole following
standard protocols. Confocal images were captured with Bio-Rad
microscope model Radiance 2000 MP with four lasers, argon (488 nm), helium-neon (543 nm), red laser diode (633 nm), and infrared
multiphoton (Mira 690-1000 nm), mounted on an Olympus IX70 microscope
and using a ×60 Plan Apo NA, 1.4 objective. For visualization of
4',6-diamidino-2-phenylindole we used two-photon excitation at 760 nm
and emission filter HQ390/70.
Measurement of Reactive Oxygen Species (ROS)/Reactive Nitrogen
Species (RNI)--
Oxidative/nitrative stress was measured by
analyzing DCFH2 oxidation to DCF. To this end, cells
transfected with pEF-VR1 plasmid were labeled with 10 µM
cell-permeable DCFH-DA (2',7'-dichlorodihydrofluorescein diacetate) for
30 min, washed, and challenged for the indicated periods of time with
different stimuli. The fluorescence corresponding to the oxidized probe
was followed by measuring the green (505-545 mm; FL1) fluorescence in
the annexin V-PE+ population (555-600 mm; FL2).
Expression of GFP-VR1 Fusion Protein--
We constructed an
N-terminal GFP fusion protein (Fig.
1A) that enabled us to
simultaneously follow the expression of VR1 (by GFP fluorescence) and
the functional consequences of VR1 stimulation with capsaicin in
transient expression experiments with J-HM1-2.2 cells. The
GFP-VR1 Functional Properties of the VR1 Channel in Jurkat Cells--
To
characterize calcium transport through the VR1 channel, we followed the
Ca2+ influx induced by capsaicin by single cell ratiometric
calcium imaging in cells loaded with a fluorescent Ca2+
probe (fura 4F, Kd for Ca2+ = 0.77 µM), which allows for an accurate estimation of
[Ca2+]c in the micromolar range. Cells bound to
coverslips were superfused with the appropriate solutions, and
fluorescence emission was imaged for [Ca2+]c
quantitation (see "Experimental Procedures"). Individual cells
expressing GFP-VR1 were identified by their fluorescence at the end of
each experiment, and Ca2+ influx in GFP-VR1+
and GFP-VR1
We compared Ca2+ influx through VR1 with
receptor-controlled, capacitative calcium influx. To this end, we took
advantage of the fact that J-HM1-2.2 cells stably express HM1R, and
HM1R stimulation by carbachol induces inositol
1,4,5-trisphosphate-mediated Ca2+ release from the ER and
subsequent capacitative calcium influx (20). More than 95% of the
cells responded to carbachol with a rise in
[Ca2+]c of up to 3-4 µM (Fig.
2A and Supplemental Material). We also confirmed the source
of the calcium responsible for these changes by stimulating the cells
with the agonists in Ca2+-free medium. Under these
conditions, the ability of capsaicin to induce the
[Ca2+]c rise was completely blocked, but
carbachol still induced a considerable [Ca2+]c
increase (Fig. 2A and Supplemental Material). These results
indicate unambiguously that the mechanisms for the
[Ca2+]c increases induced by both agonists are
completely different. Capsaicin exclusively induced ligand-gated
Ca2+ influx, whereas carbachol induced Ca2+
release from the ER and subsequent capacitative Ca2+
influx. When cells were stimulated for a sustained period of time (5-6
min), [Ca2+]c remained increased during the full
stimulation period with capsaicin but decreased rapidly from 3-4 to
0.7-0.9 µM within 2 min during stimulation with
carbachol (Fig. 2, B and C). This behavior is
consistent with different mechanisms of action, since release of
Ca2+ from the ER by carbachol ceases once the calcium store
is emptied, and then the [Ca2+]c increase is
sustained only by the stimulated capacitative Ca2+ influx,
which is much smaller than Ca2+ influx through VR1. The
shapes of the frequency distributions of the maximal
[Ca2+]c peaks induced by capsaicin and carbachol
were completely different (Fig. 2C). The VR1 agonist
increased [Ca2+]c in only 20-40% of the cells
(the ones expressing the vanilloid receptor), and 88% of the peak
values were within the 5-7 µM range (and 100% over 3 µM). On the contrary, the HM1R agonist increased
[Ca2+]c in 95% of the cells, and the frequency
distribution was quite uniform in the 1-7 µM range. The
fraction of the total cell population that responded with the largest
Ca2+ peaks (6-7 µM) was quite similar for
both agonists (19% for capsaicin and 17% for carbachol; Fig.
2C, right), but the responses to carbachol were
transient, whereas the responses to capsaicin were sustained over time
(Fig. 2C, left).
Capsaicin Treatment Induces Paraptosis in GFP-VR1+
Cells--
Once the functional expression of the VR1 was observed, we
analyzed whether capsaicin-induced opening of the calcium channel could
induce the exposure of phosphatidylserine (PS) from the inner to the
outer leaflet of the plasma membrane, one feature of cell death. To
this end, we electronically gated on GFP-VR1+ cells and
analyzed annexin V binding by two-color flow cytometry. As shown in
Fig. 3A, treatment with
capsaicin induced early (15-min) translocation of PS in
GFP-VR1+ cells. PS exposure increased in a time- and
dose-dependent manner (Fig. 3B). After 1 h
of treatment with capsaicin, ~70% of the GFP-VR1+ cells
had PS on the outer leaflet of the plasma membrane (Fig. 3A). As an internal control, we examined the ability of
capsaicin to induce PS exposure in the GFP-VR1
PS exposure not only occurs in apoptosis but also in necrosis. Necrotic
cell death is characterized by simultaneous annexin V staining and
increased plasma membrane permeability to small solutes such as 7-AAD.
To distinguish between apoptotic and necrotic cell death, simultaneous
tricolor flow cytometry analysis was performed. As seen in Fig.
3C, GFP-VR1+ annexin V+ cells
excluded 7-AAD during the first 1 h of treatment with capsaicin. This supports the hypothesis that VR1+ cells underwent
apoptotic rather than necrotic cell death upon capsaicin treatment.
GFP-VR1+ annexin V+ cells became permeable to
7-AAD after 2 h of culture with capsaicin (Fig. 3B,
right panel), which demonstrates that these cells
ultimately underwent necrotic cell death.
To gain insight into the pathways that connect calcium influx with cell
death, we determined the different requirements for triggering PS
exposure. First, we tested whether the presence of extracellular
calcium was required for the process. In the absence of extracellular
calcium (nominally calcium-free culture medium), capsaicin-induced PS
exposure was inhibited (Fig.
4A). Therefore, calcium influx
through the VR1 channel was required to induce cell death. Second, we
investigated the contribution of mitochondria to PS exposure.
Cyclosporin A (CSA) has been shown to inhibit apoptotic and necrotic
cell death induced by mitochondrial damage by preventing the opening of
the PTP (1). We therefore analyzed the effect of CSA on
capsaicin-induced PS exposure. As shown in Fig. 4A, CSA
prevented PS exposure on the outer leaflet of the cell membrane,
suggesting that mitochondrial PTP opening is involved in cell death
induced by capsaicin. Overexpression of the antiapoptotic protein
Bcl-2, which protects mitochondria from diverse insults (1),
also reduced capsaicin-induced PS exposure, which supports the
contribution of mitochondria to the observed cell death (Fig.
4B).
We also analyzed whether caspase activation was involved in this
process. Although the general caspase inhibitor Z-VAD prevented anti-Fas-induced apoptosis (measured as annexin-V binding), Z-VAD treatment of GFP-VR1+ cells did not interfere with
capsaicin-induced cell death (Fig. 5A). Furthermore,
GFP-VR1+ cells stably expressing the general caspase
inhibitor p35 from baculovirus exhibited reduced PS exposure in
response to
Recent results suggest that the distinction between apoptosis and
necrosis is less clear than initially thought, and caspase-independent programmed cell death and "paraptotic" cell death have been
described, which display mixed features of apoptosis and necrosis (26, 27). To distinguish among these possibilities, we performed other
classical protocols for the analysis of apoptosis such as caspase-3
activation assays, DNA fragmentation, and TUNEL. These assays were
carried out by two-color flow cytometry analysis on electronically
gated GFP-VR1+ cells or by sorting of GFP-VR1+
cells. The flow cytometry experiments quantified (i) active caspase-3 (Fig. 6A), (ii) cells in the
sub-G0/G1 peak of the cell cycle (Fig.
6B), and (iii) TUNEL+ cells.
GFP-VR1+, capsaicin-treated cells did not express active
caspase-3 (Fig. 6A), did not undergo DNA degradation into
small fragments such as those observed when Capsaicin Treatment Induces Caspase-independent
Pretreatment of the VR1-expressing cells with the general caspase
inhibitor Z-VAD did not prevent the Calcium Influx into Mitochondria Is Necessary for Capsaicin-induced
Cell Death--
PTP opening can be triggered by the increase of
[Ca2+] inside the mitochondrial matrix (1). The results
described so far suggest that the increase of
[Ca2+]c promoted by VR1 activation stimulates
mitochondrial Ca2+ uptake through the MCU, leading to an
increase in intramitochondrial [Ca2+] and subsequent PTP
opening. If this hypothesis were correct, depolarization of
mitochondria before activation of VR1 should inhibit capsaicin-induced
cell death, since Calcium Overload Does Not Induce Cytochrome c or AIF Release from
Mitochondria--
Proapoptotic factors such as cytochrome c
and AIF are released from mitochondria during apoptotic processes.
Since we have demonstrated that mitochondrial damage is essential for
Ca2+-induced cell death, we examined whether cytochrome
c and AIF were released from mitochondria in the cell death
model studied here, which showed a paraptotic phenotype. Cytochrome
c release was assessed by Western blot analysis (Fig.
9A) of soluble and particulate
fractions from GFP-VR1+ and GFP-VR1
We also analyzed whether AIF release from mitochondria was associated
with calcium-induced cell death. We could not detect AIF in either the
soluble fractions (Western blot; Fig. 9C) or cell nuclei
(Fig. 9D) from GFP-VR1+ cells treated with
capsaicin. In contrast, staurosporin treatment induced AIF release into
the cytosol (Fig. 9C), subsequent translocation to the
nucleus, and chromatin condensation (Fig. 9D).
Calcium Overload Induces Oxidative/Nitrative
Stress-dependent Cell Death--
Oxidative/nitrative
stress plays a pivotal role in regulating programmed cell death. ROS as
well as RNI can be generated after mitochondrial damage and may
subsequently mediate apoptosis or necrosis (1). In order to assess free
radical production by calcium overload, we analyzed DCFH2
oxidation in cells expressing the VR1 channel. Capsaicin induced early
(5 min) accumulation of DCF as measured by an increase in fluorescence
(Fig. 10A) in the annexin
V+ population; after 30 min, the increase in DCF
fluorescence was comparable with that of cells treated with the
uncoupler CCCP. Antioxidants such as N-acetyl-1-cysteine
(NAC) are useful tools in determining the involvement of ROS/RNI in
programmed cell death (34). To this end, we measured real time PS
exposure in response to capsaicin, in the presence or absence of NAC.
As shown in Fig. 10B, NAC inhibited capsaicin-induced PS
exposure, which demonstrates the contribution of oxidative and/or
nitrative stress to Ca2+ influx-induced cell
death.
We describe a unique model of rapid cell death, triggered by
Ca2+ influx through a receptor-operated channel and
involving mitochondrial damage. This cell death fulfills several
criteria of apoptosis, such as collapse of Cell death associated with excitotoxic or ischemic neural lesions and
ischemia/reperfusion damage in several tissues shows a paraptotic
phenotype (26, 35, 36). It has been proposed that mitochondria play a
prominent role in excitotoxic cell death (4, 37), but this model is too
complex to determine the specific role of Ca2+ overload in
cell death (5) (see Introduction). In our model, cell death is
triggered exclusively by Ca2+ influx through the plasma
membrane by the direct and specific activation of a receptor-operated
plasma membrane channel, VR1. The current study provides evidence that
mitochondrial Ca2+ overload plays a decisive role in this
paraptotic cell death, as shown by the Stimulation of T cells expressing HM1R with carbachol triggers AICD
(12), but the cell death induced by carbachol is much slower than the
one induced by VR1 stimulation (Fig. 3A; compare with Ref.
12). Direct comparison of the effects of capsaicin and carbachol on
[Ca2+]c showed important differences (Fig. 2).
The effect of the VR1 agonist was much more sustained over time
compared with carbachol, as expected from the different mechanisms of
action. Whereas capsaicin treatment sustains Ca2+ influx
during the entire stimulation period, the effect of carbachol is
transient, consistent with cessation of Ca2+ release once
the ER empties. Subsequently, the plasma membrane Ca2+
ATPase pumps out the Ca2+ load, and
[Ca2+]c quickly returns toward lower levels. At
this stage, [Ca2+]c is still maintained
moderately high (<1 µM) (Fig. 2B) by
Ca2+ influx through capacitative mechanisms (calcium
release-activated channels), which are activated by the emptying of the
ER (15, 16, 38). In response to these, [Ca2+]c
calcium release-activated channels are partially inactivated, since
[Ca2+]c is kept above resting levels (39).
However, regarding mitochondrial Ca2+ accumulation,
[Ca2+]c below 1 µM, the
level maintained by carbachol, produces slow mitochondrial
Ca2+ uptake (7), whereas a [Ca2+]c of
7 µM, which is sustained with capsaicin treatment, produces a fast mitochondrial uptake. The increase in
[Ca2+]c from 1 to 7 µM accelerated
mitochondrial uptake more than 30 times in chromaffin cells (7). Our
results show that the mitochondrial calcium overload resulting from
capsaicin treatment induced PS externalization within 15 min
(Fig. 3A), whereas 1-h treatment with carbachol
had no significant effect on this parameter (Fig. 3A). Such
an early commitment to death suggests that an irreversible event, most
likely PTP opening, has occurred by this time. This hypothesis is
supported by the observation that the maximal decrease in
Release of proapoptotic factors cytochrome c and AIF from
mitochondria occurs during apoptotis. Cytochrome c release
may eventually lead to the assembly of the cytochrome c,
ATP, Apaf-1, caspase-9, and caspase-3 complex (apoptosome), which
drives activation of the caspase cascade (40) and oligonucleosomal DNA
fragmentation (41). In contrast, released AIF translocates to the
nucleus and produces 50-kbp DNA fragmentation (1). Under conditions of
severe ATP depletion and/or inefficient cytochrome c
release, it is thought that caspase activation is not achieved, and
cells fail to manifest some of the caspase-dependent
features of apoptosis, this favoring the adoption of a more
"necrotic" phenotype (1, 42). In our model, capsaicin-induced cell
death occurred without release of AIF or cytochrome c from
mitochondria (Fig. 9). Also calcium-induced mitochondrial damage and
cell death occurred without concomitant caspase-3 activation and were
not inhibited by general caspase inhibitors. In addition, neither 50 kbp nor oligonucleosomal DNA degradation were found. The lack of AIF
release correlated with an absence of 50-kbp DNA fragmentation, which
is one hallmark of AIF-induced programmed cell death (42).
Capsaicin-induced cell death was instead associated with cleavage of
the DNA into ~1000-kbp fragments (Fig. 6C). Interestingly,
such a DNA degradation pattern has been reported during cell death
induced by glutamate in cerebellar granule cells (43). This pattern is
different from the features of AICD and resembles characteristics
reported for caspase-independent cell death (3, 26) and the so-called paraptotic programmed cell death (26, 27).
We observed rapid DCFH2 oxidation upon VR1 stimulation, and
cell death was inhibited by the antioxidant NAC, which supports the
role of oxidative/nitrative stress in calcium overload-induced paraptosis. Future experiments using specific NO probes and scavengers will help to identify the nature of the molecules involved in the
oxidation of DCFH2 (ROS and/or NOI) and their relative
contribution to paraptosis. Our preliminary
data2 demonstrate that
Finally, our results indicate that T cells possess all of the
mechanisms necessary for coupling mitochondrial Ca2+
overload to cell death. The paraptotic cell death resulting from activation of the transfected VR1 receptors resembles
ischemia/reperfusion or excitotoxic damage found in excitable tissues,
which possess mechanisms for massive Ca2+ entry. The
emerging inference is that the type of stress a cell undergoes
determines the cell death response (apoptosis, paraptosis, or
necrosis) by evoking the corresponding death mechanisms, which may be universal for all cell types.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
m)1 (1),
the opening of the mitochondrial permeability transition pore or
megachannel (PTP) and the release of proapoptotic factors such as
cytochrome c (2) and/or apoptosis inducer factor (AIF) (3).
It has been shown that high [Ca2+] (100-500
µM) is necessary for PTP opening in isolated mitochondria (1), but this [Ca2+] is not attained in the cytosol of
intact cells. It has been proposed that mitochondrial Ca2+
overload in intact cells can trigger the opening of PTP and release of
proapoptotic factors during neuronal ischemic and/or excitotoxic cell
death (4, 5), but the downstream events that couple the rise in calcium
to cell death are unknown. Mitochondrial Ca2+ overload
would be a side result of the rise in the cytosolic Ca2+
concentration ([Ca2+]c) promoted by
Ca2+ entry through plasma membrane receptor-operated and
voltage-dependent Ca2+ channels. When the
[Ca2+]c increases, Ca2+ is taken up
into the mitochondria via the mitochondrial Ca2+ uniporter
(MCU), a low affinity, high capacity Ca2+ transport system
driven by the
M. Since uptake through MCU is a
function of ([Ca2+]c) (6), its extent critically
depends on both the magnitude and duration of the
[Ca2+]c rise (7).
M and
caspase-independent, paraptotic cell death. This cell death pattern is
completely different from AICD, thus revealing that different
programmed cell death pathways can coexist in the same cell and be
selectively induced by diverse stress stimuli.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-actin was from Sigma.
·{(R
Rmin)/(Rmax
R)}, where R is the ratio between the
fluorescence emissions measured at 340- and 380-nm excitation,
Rmax and Rmin are the
ratios obtained at saturation of the dye with Ca2+ and in
the absence of Ca2+, respectively, Kd is
the dissociation constant for the dye (0.77 µM), and
is the ratio of the maximal (in the absence of Ca2+) and
the minimum (at saturation with Ca2+) fluorescence
emissions measured at 380-nm excitation. The values of
Rmax and Rmin and
were determined in cells permeabilized to Ca2+ with
ionomycin and perfused with media containing either no Ca2+
(5 mM EGTA) or 10 mM Ca2+. These
values were similar to the ones obtained with fura-2. These procedures
have been previously described (24). The standard perfusion solution
was as described above for fura-4F loading except that bovine serum
albumin (1 mg/ml) and EGTA (50 µM) were added to prevent
spontaneous activation of cells during measurements. For the
Ca2+-free solution, CaCl2 was omitted from the
standard perfusion solution, and 100 µM EGTA was added.
cells.
20 °C) for 5 min in order to preserve GFP fluorescence.
Cytosolic DNA fragments were then extracted by incubation with DNA
extraction buffer (0.2 M Na2HPO4, 1 mM citric acid, 10 min). Finally, cells were resuspended in
DNA staining solution (PBS; 100 µg/ml RNase A, 20 µg/ml PI) before
analysis by flow cytometry. DNA degradation was determined as the
percentage of DNA located in the sub-G0/G1 peak
of the cell cycle in both GFP-VR1+ and
GFP-VR1
cells.
cells were electronically gated, and PS
(phosphatidylserine) exposure and 7-AAD permeability were analyzed in
both populations. For continuous recording of annexin-V binding, cells
were incubated with Hanks' balanced salt solution supplemented with up
to 2.5 mM CaCl2, and annexin-V PE binding was
measured in gated cell populations by the increase in FL2 fluorescence.
cells.
m--
Changes in
m were determined by staining the cells after
transfection with 100 nM MitoTracker Red (CMX-Ros;
Molecular Probes) for 15 min at 37 °C in a humidified atmosphere
containing 5% CO2. Cells were then washed in Hanks'
balanced salt solution, supplemented or not with calcium (2 mM) when required, and analyzed by flow cytometry by
measuring the red (605-635 mm; FL3) fluorescence in both
GFP-VR1+ and GFP-VR1
cells. The decrease in
m is seen as a decrease in CMX-Ros fluorescence. Control experiments were performed using CCCP (10 µM;
Sigma). For the continuous recording of measurements determining
m changes, cell suspensions were kept at 37 °C
during data acquisition (20 min).
cells was analyzed by
Western blot analysis as previously described (25). Supernatants
(cytosolic extracts free of mitochondria) and pellets (particulate
fraction that contains mitochondria) from digitonin-permeabilized cells
were electrophoresed on a 15% SDS-polyacrylamide gel and then
analyzed by WB using anti-cytochrome c antibody (7H8.2C12;
BD Pharmingen) or polyclonal anti-AIF antibody as described above. To
verify proper separation of the mitochondria, membranes were reprobed
with anti-cytochrome oxidase antibody. Protein loading was normalized
by using an anti-
-actin monoclonal antibody.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
cells in this heterogeneous cell population
constituted an excellent internal control for the functional
experiments. In routine experiments, GFP-VR1 expression was achieved in
20-40% of Jurkat cells, as evidenced by flow cytometry (Fig.
1B). Expression of the construct was also confirmed by
Western blot analysis (Fig. 1C), and its localization to the
plasma membrane was assessed by confocal microscopy (Fig.
9B).
View larger version (38K):
[in a new window]
Fig. 1.
Transient expression of GFP-VR1.
A, GFP-VR1 construct. GFP was fused to N-terminal rat VR1,
and the resulting open reading frame was under the control of the
pEF-Bos promoter. B, flow cytometry analysis of J-HM1-2.2
Jurkat cells after transfection with the pEFGFP-VR1 construct or an
empty vector (pEF4). C, kinetics of GFP-VR1 expression. WB
analysis of GFP-tagged proteins was performed in cell lysates at
different time points after transfection. As a negative control,
J-HM1-2.2 cells were transfected with an empty vector (pEF4). An
80-kDa GFP-tagged molecule (GFP-DGK ) (21) was used as a positive
control.
cells present in the same field was analyzed
separately. Sequential stimulation of cells with capsaicin and
carbachol induced changes in [Ca2+]c; the time
course of this response is shown in Fig. 2A. The traces have been
averaged for all of the cells in each group, identified by GFP
fluorescence. Stimulation with capsaicin (0.1 µM) in
Ca2+-containing medium induced a fast increase of
[Ca2+]c from 0.1 to 6 µM in a
subpopulation of the cells, which quickly returned to base line after
the removal of capsaicin. The capsaicin-sensitive cells corresponded to
cells expressing GFP-VR1, as assessed by superimposition of
fluorescence images excited at 490 nm (to visualize GFP) and the
ratiometric Ca2+ images (fura-4F, excited at 340 and 380 nm; see upper panels in Fig. 2A).
View larger version (30K):
[in a new window]
Fig. 2.
Functional expression of GFP-VR1
Ca2+ channel. A, single-cell
[Ca2+]c image kinetic analysis of short
pulse-treated cells expressing GFP-VR1. Transiently transfected cells
were loaded with fura 4F and subjected to digital imaging
fluorescence microscopy, and [Ca2+]c images were
obtained as indicated under "Experimental Procedures." The
first color frame (GFP) at
the top corresponds to the green fluorescence image, and
subsequent ratiometric images for [Ca2+]c were
recorded every 5 s. Representative frames corresponding to the
different [Ca2+]c peaks were taken at the
indicated time points and are shown at the top.
[Ca2+]c profiles were averaged for the two cell
populations (GFP-VR1+, n = 45;
GFP-VR1 , n = 62) and represented at the
bottom. Cells were superfused during the indicated periods
with the different solutions containing the VR1 agonist capsaicin (0.1 µM) and the HM1R agonist carbachol (50 µM),
in the absence (empty bar) or the presence
(gray bar) of calcium in the external medium. See
also the video file provided in Supplemental Material. B,
single-cell [Ca2+]c measurements corresponding to
long pulse-treated cells. Average [Ca2+]c profile
analysis was performed as indicated in A, and treatment with
the different agonists was maintained for the time indicated. The
solid line corresponds to average
[Ca2+]c of GFP-VR1+ cells
(n = 30), whereas the dashed line
shows the average response of GFP-VR1
cells
(n = 69). The S.D. in all cases was below 10% of the
mean. C, same as in B, but representing
individual cell [Ca2+]c traces (left
side). On the right, the frequency distribution
of calcium responses is shown (n = 335).
cells.
Capsaicin did not induce PS exposure in this cell group. Although
carbachol induced a [Ca2+]c increase peaking at
6-7 µM in 17% of the cells (about the same percentage
as capsaicin; see above), PS exposure did not increase above background
in carbachol-stimulated cells (Fig. 3, A and left
panel in B). This observation indicates that a
sustained [Ca2+]c increase, such as the one
produced by capsaicin, rather than a high transient
[Ca2+]c, is required to trigger PS exposure.
View larger version (37K):
[in a new window]
Fig. 3.
VR1 triggering induces PS exposure in the
absence of permeability to 7-AAD. J-HM1-2.2 cells expressing
GFP-VR1 were treated with capsaicin (0.1 and 0.04 µM) or
carbachol (50 µM) for the indicated time points.
A, gated GFP-VR1+ and GFP-VR1
cells were analyzed for PS exposure (annexin V binding). B,
kinetics of PS exposure (left) and permeability to 7-AAD
(right) in gated populations. C, dot plot
analysis of permeability to 7-AAD in GFP-VR1+, annexin
V+ cells. Where appropriate, the percentage of cells
in each subpopulation is indicated.
View larger version (40K):
[in a new window]
Fig. 4.
Calcium influx-induced PS exposure requires
extracellular calcium and involves mitochondrial PTP opening.
A, J-HM1-2.2 cells expressing GFP-VR1 were treated for
1 h with capsaicin (0.1 µM) in the absence or
presence of calcium (2 mM) in the culture medium and CSA
(10 µM). B, cells were cotransfected with 10 µg of pEFGFP-VR1 construct and either 30 µg of pEF empty vector
(control) or an equivalent amount of the pEFhbcl-2 expression vector
(hbcl-2). hbcl-2 expression was analyzed by WB analysis in the upper
panel. Transfected cells were stimulated with 0.1 µM
capsaicin or 2 µM staurosporin for the indicated periods.
The numbers represent the percentage of annexin
V+ cells in the GFP-VR1+ population.
-Fas but not in response to capsaicin (Fig.
5B). Taken together, these results indicate that caspases
are not involved in capsaicin-induced cell death. Additional
experiments were performed to test whether capsaicin treatment could
also induce other apoptotic events such as cell shrinkage and increase
in cell complexity. Treatment with capsaicin induced cell shrinkage
(measured by a decrease in forward scatter) and increased cell
complexity (increase in side scatter) as assessed by flow cytometry in
GFP-VR1+ cells (not shown). Z-VAD-insensitive,
CSA-inhibitable PS exposure, cell shrinkage, and increase in cell
complexity were also induced by capsaicin in three other cell lines
transiently expressing GFP-VR1: NCB20 (neuroblastoma), HEK 293 (human
embryonic kidney cells), and WEHI231 (pre-B cell line), which indicates
that this process is not restricted to Jurkat cells (results not
shown).
View larger version (33K):
[in a new window]
Fig. 5.
Activation of caspases is not involved in
calcium overload-induced PS exposure. A, J-HM1-2.2
cells expressing GFP-VR1 were treated with capsaicin (0.1 µM) for 1 h in the presence or absence of Z-VAD (100 µM) and -Fas (100 ng/ml) as a control. B,
J-HM1-2.2 or J-HM1-2.2 cells stably expressing the p35 caspase
inhibitor were transfected with GFP-VR1 expression vector. PS exposure
was assessed after capsaicin and
-Fas treatment for 1 h. p35
expression was analyzed by WB analysis (upper
panel). The numbers represent the percentage of
annexin V+ cells in the GFP-VR1+
population.
-Fas was used (Fig.
6B), and were TUNEL
(not shown). Consistent
with the results of the cell cycle analysis, GFP-VR1+-sorted cells treated with capsaicin did not
undergo oligonucleosomal DNA laddering as assessed by agarose gel
electrophoresis of genomic DNA (Fig. 6C, left).
Pulse field electrophoresis of DNA from these cells demonstrated the
progressive appearance of 1000-kbp DNA fragments (Fig. 6C,
right) that did not progress to 50 kbp and/or oligonucleosomal fragmentation as occurs in AIF-dependent
apoptosis or in classical apoptosis (26-28). In contrast, treatment
with staurosporin, a well known apoptosis inducer, produced initial fragmentation of the DNA in 1000 kbp and subsequent processing to 50 kbp (Fig. 6C, right). Taken together, the data
show that capsaicin-induced cell death fulfills several criteria for
both apoptosis and necrosis.
View larger version (41K):
[in a new window]
Fig. 6.
Ca2+ influx does not induce
caspase-3 activation or oligonucleosomal/50-kbp DNA fragmentation.
A, two-color flow cytometry analysis of gated
GFP-VR1+ cells was performed for detection of active
caspase-3. B, cell cycle analysis on gated
GFP-VR1+ cells was performed by DNA staining with PI.
C, oligonucleosomal DNA fragmentation (left) and
pulse-field agarose (right) gel electrophoresis of genomic
DNA from sorted GFP-VR1+ cells, treated for the indicated
time points with capsaicin (0.1 µM), staurosporin
(Sts; 2 µM), or -Fas (100 ng/ml) as
positive controls.
m
Dissipation--
Protection by CSA and Bcl-2 suggested the
involvement of mitochondrial PTP opening in capsaicin-induced cell
death (Figs. 4A and 5). To further investigate the role of
mitochondria, we labeled cells with CMX-Ros (29) and measured
m by two-color flow cytometry. Treatment with
capsaicin induced a decrease of CMX-Ros fluorescence (depolarization)
in GFP-VR1+ cells but not in GFP-VR1
cells
(Fig. 7A). The decrease in
CMX-Ros fluorescence induced by capsaicin was comparable with the one
induced by treatment with the protonophore CCCP, a mitochondrial
uncoupler that collapses
m (Fig. 7, B and
C). To circumvent the possibility that CMX-Ros may be
retained into the mitochondria upon depolarization, which could
undervalue the
m loss, we performed some experiments
in which cells were treated with capsaicin before the staining with CMX-Ros. The uptake of CMX-Ros in GFP-VR1+ cells pretreated
with capsaicin was also decreased in comparison with untreated
GFP-VR1+ (Fig 7B, top). In addition,
we performed experiments prelabeling cells with a reversible,
potential-sensitive dye, tetramethyl rhodamine methyl ester (TMRE). We
observed a rapid decrease in the TMRE fluorescence of
GFP-VR1+ cells upon the addition of capsaicin or CCCP,
confirming the data obtained with CMX-Ros (Fig. 7B,
bottom). The capsaicin-induced mitochondrial depolarization
occurred rapidly;
m dissipation was evident after
2-3 min and reached a plateau in 10-15 min (Fig. 7A).
Consistent with the annexin V binding results (see above), removal of
extracellular calcium or the addition of the PTP inhibitor CSA blocked
the ability of capsaicin to dissipate
m (Fig. 7, A and C). These results suggest that dissipation
of
m was not only due to mitochondrial calcium uptake
but most probably was a consequence of PTP opening induced by
mitochondrial calcium overload. The fact that HM1R triggering did not
induce such a
m loss (not shown) provides further
support for this hypothesis.
View larger version (40K):
[in a new window]
Fig. 7.
Ca2+ influx induces
caspase-independent
m dissipation.
A, kinetic analysis of
m changes on gated
GFP-VR1+ and GFP VR1
cells, in the absence or
presence of extracellular calcium. The arrows indicate
capsaicin addition (0.1 µM). B,
top, GFP-VR1+ cells were left untreated
(CONTROL) or pretreated with capsaicin 0.1 µM
(CAPS) or 10 µM CCCP uncoupler for 5 min and
then were labeled for an additional 15-min period with CMX-Ros (100 nM), and CMX-Ros uptake was analyzed by flow cytometry.
Bottom, GFP-VR1+ cells were preloaded for 15 min
with 20 nM of the potential-sensitive dye TMRE.
Subsequently, cells were left untreated (CONTROL) or treated
with capsaicin 0.1 or 10 µM CCCP uncoupler for 5 min, and
m dissipation (TMRE fluorescence decrease) was
measured by flow cytometry. C, the experiment was performed
as in A, in the presence or absence of a 10-min pretreatment
period with CSA (10 µM). CCCP uncoupler (10 µM) was used as a positive control for
m dissipation. D, same as in A,
but pretreatment with Z-VAD (100 µM) was performed 20 min
prior to the capsaicin addition.
m collapse
induced by capsaicin (Fig. 7D), suggesting that caspases do
not contribute to the observed
m dissipation. In
summary, these results indicate that the cell death triggered by
capsaicin is mediated by calcium-induced mitochondrial damage but does
not involve caspase activation.
m is the driving force for
mitochondrial Ca2+ uptake (7, 30, 31). To test this
possibility, we studied the effect of capsaicin when mitochondria had
been depolarized by prior treatment with CCCP. Fig.
8A shows that pretreatment with CCCP at two different doses prevented PS exposure induced by
capsaicin. In another series of experiments, the MCU was blocked by
treatment with ruthenium red during the electroporation of cells (32).
This treatment also prevented capsaicin-induced PS exposure (Fig.
8B). Thus, inhibition of mitochondrial Ca2+
uptake, either by collapsing
m with CCCP or by
inhibiting MCU with ruthenium red, prevented cell death. These results
indicate that accumulation of Ca2+ into mitochondria is
essential for PTP activation and cell death induced by VR1 opening.
View larger version (42K):
[in a new window]
Fig. 8.
Ca2+ influx-dependent
PS exposure requires mitochondrial calcium uptake. A,
kinetics of annexin V binding on GFP-VR1+ cells was
performed by incubating cells with Hanks' balanced salt solution
supplemented with calcium and annexin V-PE, in the absence or the
presence of a 10-min pretreatment with CCCP uncoupler (0.025 and 0.01 µM in left and right
panel, respectively). B, fixed time point
analysis of annexin-V binding to GFP-VR1+ cells incubated
during electroporation with ruthenium red (0.01 and 0.05 µM) and treated with capsaicin (0.1 µM)
36 h later.
sorted
cells and by immunofluorescence analysis (Fig. 9B) using anti-cytochrome c antibody. Actinomycin D was used as a
positive control (33). Sorted GFP-VR1+ cells underwent
massive PS exposure (82% annexin-V+ cells; not shown)
within 45 min after the capsaicin addition. However, this treatment did
not induce detectable cytochrome c release (Fig.
9A). In contrast, treatment with actinomycin D (Act. D) for 4 h induced a lower extent of PS exposure (60%
annexin-V+ cells; not shown), but cytochrome c
was released from mitochondria in both GFP-VR1+ and
GFP-VR1
cells as assessed by Western blot analysis of
cytosolic fractions (Fig. 9A) and confocal
immunofluorescence (Fig. 9B).
View larger version (23K):
[in a new window]
Fig. 9.
Ca2+
influx-dependent paraptosis does not involve cytochrome
c or AIF release. A,
GFP-VR1 and GFP-VR1+ sorted cells were
analyzed for cytochrome c release to cytosol by WB after
treatment with capsaicin (0.1 µM, 45 min) or actinomycin
D (2 µg/ml, 4 h) as a control. B, cytochrome
c (Cyt c) redistribution was analyzed by confocal
immunofluorescence in GFP-VR1+ and GFP-VR1
cells treated as indicated in A. A polyclonal anti-human
mitochondria (MIT) antibody was used for co-localization
(47). C, same as in A, but AIF release to soluble
fractions was assessed by WB by using polyclonal anti-AIF. As a
control, cells were treated with staurosporin (sts; 2 µM) for 1 h. Mitochondria contamination in the
soluble fractions was assessed by using anti-cytochrome oxidase subunit
II (COX). D, AIF redistribution was assessed by
confocal immunofluorescence. DAPI,
4',6-diamidino-2-phenylindole.
View larger version (43K):
[in a new window]
Fig. 10.
Oxidative-nitrative stress is involved in
Ca2+ influx-dependent paraptosis.
A, cells transfected with pEF-VR1 expression vector were
loaded with DCFH-DA probe as indicated under "Experimental
Procedures." They were subsequently challenged with capsaicin (0.1 µM) and CCCP (10 µM) for the indicated
periods, and the fluorescence corresponding to oxidized probe (DCF) was
measured in the gated annexin V+ population. B,
kinetics of annexin-V binding was performed on GFP-VR1+
cells by directly incubating cells in phenol red-free Dulbecco's
modified Eagle's medium and supplemented up to 2.5 mM
calcium and annexin V-PE in the presence or absence of antioxidant NAC
(10 mM). At the indicated times, cells were challenged with
capsaicin (0.1 µM).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
m,
decrease in cell size, increase in cell complexity, exposure of
phosphatidylserine on the outer leaflet of the plasma membrane, and
lack of permeability to small solutes (7-AAD). Nonetheless, it occurred
in the absence of detectable AIF and cytochrome c release,
caspase activation, or 50-kbp/oligonucleosomal DNA degradation. Thus,
the calcium-induced cell death described in this paper resembles the
so-called paraptotic programmed cell death, which displays
mixed features of both apoptotic and necrotic death (26, 27).
M collapse
upon VR1 receptor activation leading to PTP opening and cell death, as
expected from the electrogenic Ca2+ uptake through MCU and
activation of PTP. It is remarkable that death is prevented by blocking
MCU with ruthenium red (Fig. 8B). Consistent with calcium's
role in cell death, removal of extracellular Ca2+ prevented
mitochondrial depolarization (Fig. 7A) and cell death (Fig.
4A). Also, CSA, a PTP blocker (1), prevented both the
m collapse and the PS externalization induced by
capsaicin (Figs. 7B and 4A, respectively). In
addition, depolarization of mitochondria with CCCP, a mitochondrial
Ca2+ uptake inhibitor (7, 31), before VR1 activation
prevents the early events of cell death (Fig. 8A). These
findings demonstrate that sustained [Ca2+]c
levels in the low micromolar range are sufficient to trigger PTP
opening, mitochondrial damage, and cell death.
m has also taken place after 15 min of treatment (Fig. 7A). In contrast, commitment to apoptosis during AICD
only takes place after 3-4 h, when enough FasL protein has been
synthesized (12).
m loss still occurs when ROS accumulation is
inhibited with antioxidant NAC, and these results locate
oxidative/nitrative stress downstream of calcium-induced mitochondrial
damage. Taken together, our data provide the first evidence for a
direct involvement of cellular calcium influx on mitochondrial damage,
subsequent oxidative/nitrative stress, and paraptotic cell death.
Supporting the phatophysiological importance of our model, it is
remarkable that stimulation of the VR1 receptor with anandamide, an
endocannabinoid that accumulates during in vivo induced
excitotoxic necrosis/apoptosis (44) and in
lipopolysaccharide-stimulated lymphocytes (45), induces mitochondrial
damage and subsequent neural and immune cell death (46).
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. J. Alvarez, Dr. J. A. García-Sanz, Dr. M. A. Gijón, Dr. A. Ruiz-Vela, Dr. A. Van Linden, and Dr. C. Villalobos for critical reading of the manuscript and Dr. Susin for the generous gift of the anti-AIF antibody. We thank E. Olea, M. C. Moreno, I. López-Vidriero, and E. Ruifernández for help with flow cytometry.
![]() |
FOOTNOTES |
---|
* This work was supported by grants from Ministerio de Ciencia y Tecnología/European Union and Plan Nacional de Investigación Científica, Desarrollo e Innovación Tecnológica (FEDER, 1FD97-1725-C02-01 and C02-02) and Comisión Interministerial de Ciencia y Tecnología, Spain, Grant SAF2000-0118-C03-02).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The on-line version of this article (available at
http://www.jbc.org) contains a video file.
§ Recipient of a fellowship from the Spanish Ministerio de Ciencia y Tecnología.
To whom correspondence should be addressed. Tel.:
34-983-423000 (ext. 4589); E-mail: mizdo@ibgm.uva.es.
Published, JBC Papers in Press, February 5, 2003, DOI 10.1074/jbc.M211388200
2 E. Jambrina, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
m, mitochondrial membrane potential;
7-AAD, 7-amino-actinomycin D;
Ab, antibody;
AICD, activation-induced cell death;
AIF, apoptosis inducer
factor;
Cap, capsaicin;
CMX-Ros, MitoTracker Red;
CSA, cyclosporin A;
CCCP, m-chlorophenyl hydrazone;
DCF, 2',7'-dichlorofluorescein;
DCFH2, 2',7'-dichlorodihydrofluorescein;
DCFH-DA, 2',7'-dichlorodihydrofluorescein diacetate;
GFP, green fluorescent
protein;
ER, endoplasmic reticulum;
HM1R, human muscarinic receptor
type 1;
kbp, kilobase pairs;
MCU, mitochondrial calcium uniporter;
NAC, N-acetyl-1-cysteine;
PBS, phosphate-buffered
saline;
PI, propidium iodide;
PS, phosphatidylserine;
PTP, permeability
transition pore;
RNI, reactive nitrogen intermediates;
ROS, reactive
oxygen species;
TMRE, tetramethyl rhodamine methyl ester;
TUNEL, terminal deoxynucleotidyltransferase-mediated dUTP nick and labeling;
VR1, vanilloid receptor subtype-1;
WB, Western blot;
Z-VAD, benzyloxycarbonyl-Val-Ala-DL-Asp-fluoromethylketone..
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Kroemer, G., and Reed, J. C. (2000) Nat. Med. 6, 513-519[CrossRef][Medline] [Order article via Infotrieve] |
2. | Adrain, C., and Martin, S. J. (2001) Trends Biochem. Sci. 26, 390-397[CrossRef][Medline] [Order article via Infotrieve] |
3. | Susin, S. A., Lorenzo, H. K., Zamzami, N., Marzo, I., Snow, B. E., Brothers, G. M., Mangion, J., Jacotot, E., Costantini, P., Loeffler, M., Larochette, N., Goodlett, D. R., Aebersold, R., Siderovski, D. P., Penninger, J. M., and Kroemer, G. (1999) Nature 397, 441-446[CrossRef][Medline] [Order article via Infotrieve] |
4. |
Schinder, A. F.,
Olson, E. C.,
Spitzer, N. C.,
and Montal, M.
(1996)
J. Neurosci.
16,
6125-6133 |
5. |
Duchen, M. R.
(2000)
J. Physiol.
529,
57-68 |
6. | Gunter, T. E., and Pfeiffer, D. R. (1990) Am. J. Physiol. 258, C755-C786[Medline] [Order article via Infotrieve] |
7. | Montero, M., Alonso, M. T., Carnicero, E., Cuchillo-Ibanez, I., Albillos, A., Garcia, A. G., Garcia-Sancho, J., and Alvarez, J. (2000) Nat. Cell Biol. 2, 57-61[CrossRef][Medline] [Order article via Infotrieve] |
8. | McConkey, D. J., Nicotera, P., and Orrenius, S. (1994) Immunol. Rev. 142, 343-363[Medline] [Order article via Infotrieve] |
9. |
Alvarez, J.,
Montero, M.,
and Garcia-Sancho, J.
(1999)
News Physiol. Sci.
14,
161-168 |
10. | Strasser, A. (1995) Nature 373, 385-386[CrossRef][Medline] [Order article via Infotrieve] |
11. | Golstein, P. (1998) Science 281, 1283[CrossRef][Medline] [Order article via Infotrieve] |
12. | Izquierdo, M., Ruiz-Ruiz, M. C., and López-Rivas, A. (1996) J. Immunol. 157, 21-28[Abstract] |
13. | Dhein, J., H., W., Baumler, C., Debatin, K.-M., and Krammer, P. (1995) Nature 373, 438-441[CrossRef][Medline] [Order article via Infotrieve] |
14. | Izquierdo, M., Grandien, A., Criado, M. L., Robles, S., Leonardo, E., Albar, J. P., González de Buitrago, G., and Martínez-A, C. (1999) EMBO J. 18, 101-111 |
15. | Premack, B. A., McDonald, T. V., and Gardner, P. (1994) J. Immunol. 1994, 5226-5240 |
16. | Montero, M., Alvarez, J., and Garcia-Sancho, J. (1992) Biochem. J. 288, 519-525[Medline] [Order article via Infotrieve] |
17. | Caterina, M. J., Schumacher, M. A., Tominaga, M., Rosen, T. A., Lavine, J. D., and Julius, D. (1997) Nature 389, 816-824[CrossRef][Medline] [Order article via Infotrieve] |
18. |
Schumacher, M. A.,
Moff, I.,
Sudanagunta, S. P.,
and Levine, J. D.
(2000)
J. Biol. Chem.
275,
2756-2762 |
19. | Macho, A., Calzado, M. A., Munoz-Blanco, J., Gomez-Diaz, C., Gajate, C., Mollinedo, F., Navas, P., and Munoz, E. (1999) Cell Death Differ. 6, 155-165[CrossRef][Medline] [Order article via Infotrieve] |
20. | Desai, D. M., Newton, M. E., Kadlecek, T., and Weiss, A. (1990) Nature 348, 66-69[CrossRef][Medline] [Order article via Infotrieve] |
21. |
Sanjuan, M. A.,
Jones, D. R.,
Izquierdo, M.,
and Merida, I.
(2001)
J. Cell Biol.
153,
207-220 |
22. |
Claveria, C.,
Albar, J. P.,
Serrano, A.,
Buesa, J. M.,
Barbero, J. L.,
Martinez, A. C.,
and Torres, M.
(1998)
EMBO J.
17,
7199-7208 |
23. | Grynkiewicz, G., Poenie, M., and Tsien, R. Y. (1985) J. Biol. Chem. 260, 3440-3450[Abstract] |
24. |
Villalobos, C.,
Nunez, L.,
and Garcia-Sancho, J.
(1996)
FASEB J.
10,
654-660 |
25. |
Chandra, J.,
Niemer, I.,
Gilbreath, J.,
Kliche, K. O.,
Andreeff, M.,
Freireich, E. J.,
Keating, M.,
and McConkey, D. J.
(1998)
Blood
92,
4220-4229 |
26. | Leist, M., and Jaattela, M. (2001) Nat. Rev. Mol. Cell. Biol. 2, 589-598[CrossRef][Medline] [Order article via Infotrieve] |
27. |
Sperandio, S.,
de Belle, I.,
and Bredesen, D. E.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
14376-14381 |
28. |
Susin, S. A.,
Daugas, E.,
Ravagnan, L.,
Samejima, K.,
Zamzami, N.,
Loeffler, M.,
Costantini, P.,
Ferri, K. F.,
Irinopoulou, T.,
Prevost, M. C.,
Brothers, G.,
Mak, T. W.,
Penninger, J.,
Earnshaw, W. C.,
and Kroemer, G.
(2000)
J. Exp. Med.
192,
571-580 |
29. |
Marzo, I.,
Brenner, C.,
Zamzami, N.,
Jurgensmeier, J. M.,
Susin, S. A.,
Vieira, H. L.,
Prevost, M. C.,
Xie, Z.,
Matsuyama, S.,
Reed, J. C.,
and Kroemer, G.
(1998)
Science
281,
2027-2031 |
30. |
Babcock, D. F.,
Herrington, J.,
Goodwin, P. C.,
Park, Y. B.,
and Hille, B.
(1997)
J. Cell Biol.
136,
833-844 |
31. |
Montero, M.,
Alonso, M. T.,
Albillos, A.,
Garcia-Sancho, J.,
and Alvarez, J.
(2001)
Mol. Biol. Cell
12,
63-71 |
32. |
Duchen, M. R.
(1999)
J. Physiol.
516,
1-17 |
33. | Ruiz-Vela, A., Albar, J. P., and Martinez, C. A. (2001) FEBS Lett. 501, 79-83[CrossRef][Medline] [Order article via Infotrieve] |
34. | Curtin, J. F., Donovan, M., and Cotter, T. G. (2002) J. Immunol. Methods 265, 49-72[CrossRef][Medline] [Order article via Infotrieve] |
35. |
Crompton, M.
(2000)
J. Physiol.
529,
11-21 |
36. |
Gill, C.,
Mestril, R.,
and Samali, A.
(2002)
FASEB J.
16,
135-146 |
37. | Nicholls, D. G., and Budd, S. L. (1998) Biochim. Biophys. Acta 1366, 97-112[Medline] [Order article via Infotrieve] |
38. |
Alvarez, J.,
Montero, M.,
and Garcia-Sancho, J.
(1992)
FASEB J.
6,
786-792 |
39. | Lewis, R. S. (2001) Annu. Rev. Immunol. 19, 497-521[CrossRef][Medline] [Order article via Infotrieve] |
40. |
Green, D. R.,
and Reed, J. C.
(1998)
Science
281,
1309-1312 |
41. | Enari, M., Sakahira, H., Yokoyama, H., Okawa, K., Iwamatsu, A., and Nagata, S. (1998) Nature 391, 43-50[CrossRef][Medline] [Order article via Infotrieve] |
42. |
Daugas, E.,
Susin, S. A.,
Zamzami, N.,
Ferri, K. F.,
Irinopoulou, T.,
Larochette, N.,
Prevost, M. C.,
Leber, B.,
Andrews, D.,
Penninger, J.,
and Kroemer, G.
(2000)
FASEB J.
14,
729-739 |
43. | Slagsvold, H. H., Marvik, O. J., Eidem, G., Kristoffersen, N., and Paulsen, R. E. (2000) Exp. Brain Res. 135, 173-178[CrossRef][Medline] [Order article via Infotrieve] |
44. | Hansen, H. H., Ikonomidou, C., Bittigau, P., Honoré Hansen, S., and Hansen, H. S. (2001) J. Neurochem. 76, 39-46[CrossRef][Medline] [Order article via Infotrieve] |
45. | Maccarrone, M., De Petrocellis, L., Bari, M., Fezza, F., Salvati, S., Di Marzo, V., and Finazzi-Agro, A. (2001) Arch. Biochem. Biophys. 393, 321-328[CrossRef][Medline] [Order article via Infotrieve] |
46. |
Maccarrone, M.,
Lorenzon, T.,
Bari, M.,
Melino, G.,
and Finazzi-Agrò, A.
(2000)
J. Biol. Chem.
275,
31938-31945 |
47. |
Ruiz-Vela, A.,
Gonzalez de Buitrago, G.,
and Martinez, A. C.
(1999)
EMBO J.
18,
4988-4998 |