From the Department of Radiation Oncology and the
Laboratory of Signal Transduction, Memorial Sloan-Kettering
Cancer Center, New York, New York 10021 and the ¶ Queensland
Institute of Medical Research, P. O. Royal Brisbane Hospital, Herston,
Brisbane, Queensland 4029, Australia
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ABSTRACT |
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DNA double-stranded breaks (dsb) activate
surveillance systems that identify DNA damage and either initiate
repair or signal cell death. Failure of cells to undergo appropriate
death in response to DNA damage leads to misrepair, mutations, and
neoplastic transformation. Pathways linking DNA dsb to reproductive or
apoptotic death are virtually unknown. Here we report that metabolic
incorporation of 125I-labeled 5-iodo-2'deoxyuridine,
which produces DNA dsb, signaled de novo ceramide synthesis
by post-translational activation of ceramide synthase (CS) and
apoptosis. CS activation was obligatory, since fumonisin B1, a fungal
pathogen that acts as a specific CS inhibitor, abrogated DNA
damage-induced death. X-irradiation yielded similar results.
Furthermore, inhibition of apoptosis using the peptide caspase
inhibitor benzyloxycarbonyl-Val-Ala-Asp fluoromethylketone did not
affect CS activation, indicating this event is not a consequence of
induction of apoptosis. ATM, the gene mutated in ataxia
telangiectasia, is a member of the phosphatidylinositol 3-kinase family
that constitutes the DNA damage surveillance/repair system.
Epstein-Barr virus-immortalized B cell lines from six ataxia
telangiectasia patients with different mutations exhibited radiation-induced CS activation, ceramide generation, and apoptosis, whereas three lines from normal patients failed to manifest these responses. Stable transfection of wild type ATM cDNA
reversed these events, whereas antisense inactivation of ataxia
telangiectasia-mutated gene product in normal B cells conferred the
ataxia telangiectasia phenotype. We propose that one of the functions
of ataxia telangiectasia-mutated gene product is to constrain
activation of CS, thereby regulating DNA damage-induced apoptosis.
The predominant form of death induced in mammalian cells by
ionizing radiation is reproductive (also known as clonogenic) cell
death. The target for radiation is the DNA, and double-stranded breaks
(dsb)1 are regarded as the
specific lesions that initiate this lethal pathway (1, 2). Although
most radiation-induced DNA dsb are rapidly repaired by constitutively
expressed repair mechanisms, residual unrepaired or misrepaired breaks
lead to genetic instability, increased frequency of mutations, and
chromosomal aberrations (1-3). Lethal mutations or dysfunctional
chromosomal aberrations eventually lead to progeny cell death (4, 5),
usually after several mitotic cycles (3, 6).
Although this mechanism has been extensively investigated, the
signaling pathways involved are only partially known. Once generated,
DNA damage activates a coordinate network of signal transduction
pathways that detect DNA breaks, arrest the cells temporarily at
G1, S, and G2 checkpoints, and activate DNA
repair (7-15). This signaling network is regulated by a family of
phosphatidylinositol 3-kinases (PI3-K) which in the human includes the
ataxia telangiectasia-mutated gene product (ATM), the catalytic unit of
DNA-dependent protein kinase (DNA-PK), the AT- and
RAD3-related kinase (Atr), and Frap (FKBP12 and rapamycin-binding
protein kinase) (8, 10, 11, 13-16). Repair of DNA damage is believed
to be carried out, at least in part, by non-homologous recombination
and ligase IV-mediated end joining (17).
Radiation-induced DNA dsb can also signal apoptosis, albeit
significantly less frequently than reproductive cell death (18). Definitive evidence for the involvement of radiation-induced DNA dsb in
apoptosis was provided by experiments utilizing metabolic incorporation
of 125I-labeled 5-iodo-2'-deoxyuridine
([125I]dURd). IdURd replaces thymidine in DNA without
affecting cell viability, although it sensitizes mammalian cells to the
lethal effects of radiation (4, 19). 125I decays by
electron capture, emitting cascades of low energy Auger electrons that
deposit radiation energy within less than 40 Å from the decay site
(20, 21). When targeted to DNA via [125I]dURd, the
radioactive iodine produces dsb within a range of 6 base pairs from the
site of 125I incorporation (22, 23).
[125I]dURd incorporation into the DNA was reported to
induce apoptotic responses in murine lymphoid and myeloid cell lines
(5, 24), whereas targeting of 125I to mitochondria (25),
lysosomes (26), or the cytosol (27) was found to be non-toxic. The
apoptotic response in cells treated with [125I]dURd was
shown to correlate with the number of 125I-induced DNA dsb
(5, 24).
Of the PI-3 kinases associated with DNA damage surveillance, only
ATM has been found to be involved in the apoptotic response to radiation. Hypersensitivity to radiation-induced apoptosis was
reported recently in ATM mutant fibroblast lines (28),
ATM knock-out mouse intestinal and dermal epithelial cells
(29), and in M059J human glioma
cells.2 ATM
mutations produce the genetic disorder ataxia telangiectasia (AT), a
multi-system disease manifested by abnormalities of the immune,
nervous, cutaneous, and endocrine systems (16, 30). At the cellular
level the AT phenotype is characterized by chromosomal instability
(31), increased frequency of malignant transformation (16), defective
G1, S, and G2/M cell cycle checkpoints (32, 33), and hypersensitivity to ionizing radiation (34, 35). The
hypersensitivity of some AT cells to radiation-induced apoptosis suggests that concomitant with its role in the regulation of DNA repair
and recovery of the cell from potentially lethal DNA damage, ATM may function as inhibitor of an apoptotic pathway
activated by DNA damage.
The best known apoptotic response to radiation-induced DNA damage is
p53-mediated (32, 36, 37). In murine thymic cells, p53-mediated apoptosis was not affected by ATM (29, 38), although in
spermatogonia undergoing meiosis ATM suppressed
p53-dependent apoptosis (39). Furthermore, the
ATM/DNA-PK doubly mutated M059J glioma cell line was shown
to respond to radiation with apoptosis, whereas its sister line M059K,
isolated from a different region of the same tumor, was shown to have
intact ATM/DNA-PK and to be
apoptosis-resistant.2 Both cell lines are p53-mutated (40).
These studies indicate that in some cells p53 and ATM may function
coordinately, whereas in others they may regulate apoptosis independently.
Similar to the lack of effect on thymic apoptosis, ATM does not appear
to regulate sphingomyelinase (SMase)-mediated apoptosis, known to be
activated at the plasma membrane rather than the nucleus. Haimovitz-Friedman et al. (41) reported that radiation
induced in nuclei-free membrane preparations of endothelial cells a
rapid hydrolysis of the phospholipid sphingomyelin to generate ceramide via activation of SMase. The same response was observed in intact cells, and the ceramide generated served as a second messenger in
initiating apoptotic signaling. Definitive evidence for the role of
SMase in initiating radiation-induced apoptosis was provided by the use
of genetic models. ASMase knock-out mice failed to generate ceramide
and to develop typical apoptotic lesions in the pulmonary endothelium
after whole body irradiation, although the apoptotic response in the
thymus was preserved (42). The converse pattern was observed in whole
body irradiated p53 knock-out mice, which failed to exhibit thymic
apoptosis but showed a normal apoptotic response in the lung. These
data indicated that ASMase-mediated apoptosis, expressed in pulmonary
endothelial cells, is distinct from the p53-dependent
pathway that operates in thymocytes. Barlow et al. (38)
reported that both thymic and pulmonary endothelial apoptosis remained
unaltered in whole body irradiated ATM knock-out mice,
indicating that ATM may regulate yet another, currently undefined,
apoptotic pathway.
Whereas ASMase activation represents a general mechanism for
pro-apoptotic generation of ceramide, there is an alternative mechanism
that involves de novo synthesis of ceramide and apoptosis via activation of the enzyme ceramide synthase (CS). De novo
synthesis of ceramide occurs in the endoplasmic reticulum and
mitochondria (43, 44) via CS-mediated condensation of the sphingoid
base sphinganine and fatty acyl-CoA to form dihydroceramide, which is
rapidly oxidized to ceramide (45). Bose et al. (46) have recently reported that daunorubicin activated CS and induced ceramide generation and apoptosis in P388 and U937 cells. Furthermore, the
fungal toxin fumonisin B1 (FB1), a natural specific inhibitor of CS
(47), blocked daunorubicin-induced ceramide generation and apoptosis in
both cell types. FB1 was also reported to abrogate daunorubicin-induced
apoptosis in hen granulosa cells (48) and CPT-11-induced apoptosis in
L929 cells (49). Whether CS activation induced by
daunorubicin and CPT-11 is associated with DNA damage remains unknown.
Since FB1 did not block tumor necrosis factor-induced ceramide
generation nor tumor necrosis factor-induced apoptosis, both of which
involve SMase activation in U937 cells (46), it appears that
CS-mediated apoptosis is distinct and independent of the SMase-mediated mechanism.
In the present study, we investigated whether DNA damage and
ATM regulate the pro-apoptotic function of CS. The data
indicate that radiation-induced DNA damage signals post-translational
activation of CS, ceramide generation, and apoptosis,
whereas ATM down-modulates this response. CS activation
appeared obligatory for DNA damage-induced death since FB1 blocked
[125I]dURd-induced ceramide elevation and apoptosis.
Irradiated AT cells exhibited increased CS activation and apoptosis,
reversed by transfection with wild type ATM, whereas
antisense inactivation of ATM in normal cells recapitulated the AT
phenotype. Thus, signals from DNA damage and ATM appear to reciprocally
regulate the activity of CS and consequently apoptosis after radiation exposure.
Cell Cultures, Mice, and Irradiation--
Cloned populations of
BAEC were established from the intima of bovine aorta and grown as
described previously in DMEM supplemented with 10% calf serum,
penicillin (50 units/ml), and streptomycin (50 µg/ml) at 37 °C in
a 10% CO2 atmosphere (50). During the phase of exponential
growth, purified human recombinant basic fibroblast growth factor (1 ng/ml) was added every other day. After 5-7 days in culture, the cells
reached confluence and exhibited features of contact-inhibited
monolayers. The culture medium was then changed to fresh DMEM
supplemented with 5% heat-inactivated calf serum, penicillin (50 units/ml), and streptomycin (50 µg/ml). These plateau phase cells
were maintained in culture medium for another 3-4 days before experiments.
AT lines are EBV-transformed lymphoblastoid cells derived from the
peripheral blood of adult AT patients and have been previously characterized, whereas control lymphoblastoid lines are derived from
normal donors (35). Cells were grown in suspension culture in RPMI 1640 medium supplemented with 10% fetal calf serum (Life Technologies,
Inc.) at 37 °C in a 5% CO2 atmosphere. Stably
transfected cell lines containing ATM or antisense
ATM cDNA expression vectors were generated by Lipofectin
(Life Technologies, Inc.) transfection of pMAT1 and pMAT2 vectors,
respectively (51, 52), and grown in medium containing 0.2 mg/ml
hygromycin (Roche Molecular Biochemicals). For experiments, expression
of ATM or antisense ATM cDNA constructs in
C3ABR and AT1ABR cells was achieved by treating cells with 3-5
µM CdCl2 (Sigma) for 8-16 h and was
evaluated by Western blotting using ATM-4BA antibody as described (51).
Cell numbers were determined using a Coulter counter, model ZM (Coulter
Electronics), and cell viability was assessed by trypan blue exclusion.
LNCaP, MS1418, GM0988, and A431 cells were grown as described
previously (42, 53).
Irradiation of cultured cells was carried out at 25 °C in a
Gamma-cell 40 chamber containing two sources of 137Cs
(Atomic Energy of Canada) at a dose rate of 100 cGy/min. One hour prior
to irradiation, the culture medium was changed to RPMI 1640 medium
containing 0.1% human albumin (Alpine Biologics). For irradiation of
mice, 4-6-week-old male C3H/HeJ or homozygous acid sphingomyelinase
knock-out mice (54) received whole body irradiation, delivered using a
Cs-137 Irradiator (Shepherd Mark-I, model 68, SN643) at a dose rate of
270 cGy/min, as previously reported (42). Our animal housing facility
is approved by the American Association for Accreditation of Laboratory
Animal Care and is maintained in accordance with the regulations and
standards of the United States Department of Agriculture and the
Department of Health and Human Services, National Institutes of Health.
Treatment of BAEC with
[125I]dURd--
Incorporation of
[125I]dURd was carried out using pre-confluent BAEC,
pulsed with [125I]dURd (0.25 nM IdURd; 0.5 µCi 125I/ml) for 16 h at 37 °C. The cells were
then washed twice with Hanks' buffered saline solution and incubated
in fresh DMEM supplemented with 10% calf serum. After 3 h, cells
were detached from the monolayer by 2 min incubation with 0.05%
trypsin, 0.02% EDTA at 25 °C, pelleted, and resuspended in the
freezing medium (25% calf serum, 10% Me2SO, 65% DMEM).
The cell pellet was frozen at Quantification of Apoptosis--
Morphological changes in the
nuclear chromatin of cells undergoing apoptosis were detected by
staining with the DNA-binding fluorochrome bisbenzimide
trihydrochloride (Hoechst 33258; Sigma) as described previously (46). A
minimum of 500 cells was scored for the incidence of apoptosis for each
data point.
Caspase activity was measured using the fluorogenic caspase substrate
Z-DEVD-AFC (55). For these studies, cells were lysed in RIPA buffer (50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 0.5% sodium deoxycholate) and nuclei and cell debris
pelleted by centrifugation at 11,000 × g. Reactions
were performed using 20 µg of cell extract in 0.2 ml of reaction
buffer containing 8 µM Z-DEVD-AFC according to the
manufacturer's instructions (Kamiya Biomedical Company, Seattle, WA).
Lipid Studies--
At the indicated times after irradiation,
monolayers of BAEC or A431 cells were washed once with cold
phosphate-buffered saline (PBS), and lipids were extracted with two
incubations of methanol for 10 min at 4 °C, followed by an equal
volume of chloroform and 0.6 volume of buffered saline solution/EDTA
(135 mM NaCl, 4.5 mM KCl, 1.5 mM
CaCl2, 0.5 mM MgCl2, 5.6 mM glucose, 10 mM HEPES, pH 7.2, 10 mM EDTA). For cells in suspension culture, cells were
pelleted at 800 × g for 5 min, washed twice with cold
PBS, resuspended in 0.6 ml of cold PBS, and extracted with 1 ml of chloroform/methanol, 1 N HCl (100:100:1, v/v/v), as
described (46).
Lipids in the organic (lower) phase were dried under N2 and
subjected to mild alkaline hydrolysis (1 N methanolic KOH
for 1 h at 37 °C) to remove glycerophospholipids. Ceramide was
quantified by the diacylglycerol kinase (DAG) assay as described
previously (46). Sphingomyelin content was measured using cells labeled to isotopic equilibrium with [3H]choline and verified by
phospholipid phosphorus as described (42).
Evaluation of de Novo Synthesis of Ceramide in Intact
Cells--
[125I]dURd-treated BAEC were thawed, plated
in culture medium, and incubated at 37 °C for 12 h as described
above. Cells were then placed on ice followed by the addition of
[9,10-3H]palmitic acid (60 Ci/mmol; American Radiolabeled
Chemicals Inc.) and incubation at 37 °C to start incorporation. At
the indicated times, lipids were extracted and subjected to mild
alkaline hydrolysis as described above. 3H-Labeled ceramide
was resolved by thin layer chromatography on silica gel 60 plates using
a solvent system of chloroform/methanol/acetic acid (94:2:4), detected
by comigration with ceramide standards, and quantified by liquid
scintillation counting (46). Nonspecific incorporation, which
represents less than 10% of the total incorporation, was determined at
4 °C and subtracted from each data point.
Ceramide Synthase Assay--
Microsomal membranes were prepared
as described previously (46). Briefly, cells were washed with cold PBS,
scraped off plates, pelleted, and resuspended in 300 µl of
homogenization buffer (25 mM HEPES, pH 7.4, 5 mM EGTA, 50 mM NaF, and 10 µg/ml each of
leupeptin and soybean trypsin inhibitor). Cells were disrupted using a
motor-driven 7-ml Tenbroeck tissue homogenizer (Bellco Glass), and
lysates were pelleted at 800 × g for 5 min. The
postnuclear supernatant was centrifuged at 250,000 × g
for 35 min. The microsomal membrane pellet was resuspended in 1.0 ml of
homogenization buffer. Membranes were prepared fresh daily.
Assays of ceramide synthase activity were performed as described
previously (46). Briefly, microsomal membrane protein (75 µg) was
incubated with 2 mM MgCl2, 20 mM
HEPES, pH 7.4, 20 µM fatty acid-free bovine serum albumin
(Sigma), escalating concentrations (0-20 µM) of
sphinganine (Biomol), 70 µM unlabeled palmitoyl-coenzyme A (Amersham Pharmacia Biotech), and 1.8 µM (0.2 µCi)
[1-14C]palmitoyl-coenzyme A (55 mCi/mmol) (Amersham
Pharmacia Biotech). Sphinganine was dried under N2 from a
stock solution in 100% methanol and dissolved by sonication in the
reaction mixture prior to addition of microsomal membranes. The
reaction mixture (1 ml) was incubated at 37 °C for 1 h and then
stopped by extraction of lipids using 2 ml of chloroform/methanol
(1:2). The lower phase (500 µl) was removed, dried under
N2, and applied to a silica gel 60 thin layer chromatography plate. Dihydroceramide was resolved using a solvent system of chloroform/methanol, 3.5 N ammonium hydroxide
(85:15:1), detected by comigration with ceramide standards, and
quantified by liquid scintillation counting. Under the conditions used,
the substrate was not rate-limiting, and the reaction was linear for enzyme concentration and time.
Assays for Neutral and Acidic Sphingomyelinase--
BAEC were
irradiated on ice and subsequently incubated at 37 °C for various
periods. The cells were then harvested by scraping, washed twice with
PBS at 4 °C, and pelleted. Neutral and acidic sphingomyelinase
activities were measured in total cell lysates as described (42).
Statistical Analysis--
Statistical analysis was performed by
Student's t test and t test for correlation
coefficient. Linear regression analysis was by the method of least squares.
DNA Damage Signals Ceramide Elevation and Apoptosis in Endothelial
Cells--
Direct DNA damage was produced in bovine aortic endothelial
cell (BAEC) by metabolic incorporation of [125I]dURd.
Since the half-life time for decay of 125I is 60.14 days,
the accumulation of 125I decays and DNA dsb is slow. In
contrast, the repair of DNA dsb and transit through the cell cycle is
rapid, attenuating the effectiveness of [125I]dURd
treatment. It is, therefore, necessary to slow down cellular metabolism
for extended periods to permit sufficient generation of DNA dsb to
affect survival (23, 56). Prior studies showed that freezing over
liquid nitrogen optimized the formation of biologically relevant DNA
dsb in [125I]dURd-treated cells. In mouse L cells and
Chinese hamster V79 cells, Radford and Hodgson (23) demonstrated a
linear relationship between 125I decays per cell, DNA dsb,
and the lethal lesions per cell using this procedure. Story et
al. (24) showed that DNA dsb, but not single-stranded DNA breaks,
signaled apoptosis in [125I]dURd-treated LY-TH mouse
lymphoma cells. Freezing and thawing per se did not affect
the level of DNA dsb and, if anything, attenuated the lethal effects of
radiation, perhaps by reducing the indirect effect of radiation via
free radicals, such as reactive oxygen species (23).
For the present studies, pre-confluent BAEC were pulsed for 16 h
(approximately 1.5 doubling times) with [125I]dURd (0.25 nM IdURd; 0.5 µCi of 125I/ml). Excess
[125I]dURd not incorporated into the DNA was chased by
multiple washings with 10% calf serum in DMEM and a 3-h incubation in
the same medium. The cells were resuspended in 10% calf serum in DMEM
and stored at
Fig. 1A shows that treatment
of BAEC with [125I]dURd for various periods resulted in
an apoptotic response 24 h after thawing, whereas apoptosis was
not detected in control IdURd-untreated cells, or in cells treated with
non-radioactive IdURd (data not shown). In
[125I]dURd-treated cells, apoptosis became apparent at
approximately 12 h after thawing and reached a maximal level after
18-24 h (data not shown). The extent of apoptosis was proportional to
the duration of [125I]dURd treatment. Low levels of
apoptosis were detected as early as after 5 days of
[125I]dURd exposure, peaking at approximately 30% after
12 days.
Fig. 1B shows that [125I]dURd-treated cells
also exhibited an increase in cellular ceramide. The freeze-thaw
procedures used in these experiments did not affect the base-line
levels of ceramide. In control and [125I]dURd-treated
cells incubated at DNA Damage Induces Apoptosis via Activation of Ceramide
Synthase--
In previous studies in BAEC, we demonstrated that
radiation induced ceramide generation via activation of plasma membrane SMase (41). To test whether SMase was also activated in response to DNA
damage, BAEC were treated with [125I]dURd for 17 days and
tested for neutral sphingomyelinase and ASMase activity at time points
ranging from 10 min to 24 h after thawing. Control cells displayed
detectable neutral sphingomyelinase and ASMase activities with maximal
velocities of 302 ± 63 and 4051 ± 631 pmol/min/mg protein,
respectively. There was no increase in either SMase activity in
untreated or irradiated BAEC within the first 24 h after thawing
(n = 3).
An alternative pathway for generation of ceramide is by neo-synthesis
via CS activation (46). Neo-synthesis was assessed by the rate of
[9,10-3H]palmitic acid incorporation into ceramide. Fig.
2A shows that BAEC treated
with [125I]dURd for 17 days and then labeled with
[9,10-3H]palmitic acid at 12 h after thawing
manifested enhanced incorporation of radiolabeled palmitic acid into
ceramide over controls (p < 0.05). In contrast,
incorporation of [9,10-3H]palmitic acid into other
cellular lipids was not enhanced (data not shown). To assess the
activity of CS, microsomal membranes were prepared from the
[125I]dURd-treated cells at 12 h after thawing, and
CS activity was assayed under conditions determined as linear for time
and enzyme concentration and where substrate was not rate-limiting
(data not shown). Fig. 2B shows the kinetics of CS activity
using increasing concentrations of sphinganine.
[125I]dURd enhanced the reaction velocity
(p < 0.05 versus control, n = 6). Eadie-Hofstee transformation of the data revealed that CS from
control cells manifested a maximal velocity of 102 pmol/min/mg protein
and a Km of 1.01 µM. In
[125I]dURd-treated cells, an increase of 91% in the
Vmax (p < 0.05) was observed
with no significant change in Km (Fig.
2C). A small increase in CS activity was detectable in
[125I]dURd-treated cells as early as 4 h after
thawing, and a maximal effect was observed at 12 h (data not
shown). To test whether activation of CS by [125I]dURd
was pre- or post-translational, the effect of the protein synthesis
inhibitor cycloheximide (1 µg/ml) was tested. This concentration of
cycloheximide inhibited in BAEC >90% of protein synthesis, evaluated
by [35S]methionine incorporation into total cellular
protein (data not shown). Fig. 3,
A and B, shows that cycloheximide did not
significantly affect the pattern of CS activation by
[125I]dURd. There was a 94% increase in the
Vmax of CS from cycloheximide-untreated cells,
compared with an 89% increase in CS from cells treated with
cycloheximide (n = 3).
To assess further the role of CS in [125I]dURd-induced
apoptosis, experiments were performed using fumonisin B1
(FB1), a natural product of the fungus Fusarium moniliforme
which acts as a specific inhibitor of CS (47). Treatment of BAEC with
1-25 µM FB1 for as long as 24 h did not affect cell
proliferation or viability nor did it affect basal level of cellular
ceramide (data not shown). Concentrations of 50 µM or
higher were cytotoxic to BAEC, causing rapid cell lysis (data not
shown). This toxicity is well defined, occurring in cells with high
basal metabolism of sphingolipids (57, 58). Toxicity results from
accumulation of the sphingoid bases sphingosine and sphinganine, the
precursors for ceramide synthesis, which are cationic sphingolipids
with detergent-like properties. Fig. 3C shows that 10 µM FB1 and 25 µM FB1, a maximally tolerated
dose, significantly inhibited [125I]dURd-induced
elevation of ceramide (p < 0.05 as compared with control) and apoptosis (p < 0.05) (Fig.
3D). These data provide proof-in-principle of a role for CS
in the mechanism of the apoptotic response induced by DNA damage.
X-irradiation also Induces Apoptosis via Activation of Ceramide
Synthase--
Since external beam x-irradiation, like
[125I]dURd, damages DNA and produces DNA dsb (2, 23), we
tested whether external radiation also signals CS activation. Confluent
cultures of BAEC were exposed to increasing doses of external beam
x-irradiation, and 12 h later microsomal membranes were prepared
and assayed for CS activity. Fig.
4A shows that there was a
dose-dependent increase in CS activity, peaking with a dose
of 10 Gy. At this dose, minimal CS activation was observed at 3-4 h,
and a maximal increase from a base-line of 103-158 pmol/min/mg protein
(p < 0.05) occurred at 12 h without a significant
change in the Km of the reaction (Fig. 4,
B and C). There was also a
dose-dependent increase in de novo synthesis of
cellular ceramide, as detected by increased incorporation of
[3H]palmitic acid into ceramide in BAEC exposed to 10 Gy
(data not shown). Radiolabeling of other cellular lipids was, however,
not increased (data not shown). Radiation-induced activation of CS was
a post-transcriptional and post-translational event, since both
actinomycin D (10 ng/ml) and cycloheximide (1 µg/ml) failed to
inhibit the activation of the enzyme after exposure to 10 Gy (data not
shown).
Fig. 5 shows the impact of ceramide
synthase activation on the patterns of cellular ceramide elevation and
apoptosis. In previous studies, we reported radiation exposure of BAEC
resulted, within seconds to minutes, in SMase activation and ceramide
elevation and provided evidence that ceramide generation was required
for apoptosis to ensue (41). In the present studies we recapitulated these observations, demonstrating that within 30 s of exposure to
a dose of 5 Gy there was a 1.42-fold increase in cellular ceramide levels (p < 0.025 versus control)
(inset in Fig. 5A). Ceramide subsequently
decreased, returning to the base-line level at approximately 120 min
after radiation (data not shown). This early wave of ceramide generation, which occurred well before CS activation became detectable (Fig. 4B), was not affected by treatment with 25 µM FB1 (data not shown). Further follow-up of ceramide
levels revealed that at 4 h after irradiation a second wave of
ceramide elevation became apparent, which reached a maximal 4.2-fold of
control at 16 h (Fig. 5A). This late phase of ceramide
generation coincided with the pattern of CS activation shown in Fig.
4B. Consistent with this notion, treatment with 25 µM FB1 attenuated the late phase of ceramide generation,
shutting it off after 12 h (Fig. 5A).
The elevations of cellular ceramide were followed by apoptotic
responses. Fig. 5B shows the evolution of apoptosis as a
function of time after exposure to a dose of 5 Gy. Apoptosis was
initially detected at 4 h and increased at a constant rate,
reaching a plateau of 40% at about 16 h after irradiation. These
data were corroborated by measuring caspase activity using the
fluorimetric caspase substrate Z-DEVD-AFC. Irradiation with 2.5, 5, and
10 Gy resulted in 5 ± 1-, 20 ± 1-, and 34 ± 1-fold
increases in caspase activity over unirradiated controls at 16 h,
which were maintained for as long as 24 h. This profile did not
enable distinction between the apoptotic fraction effected by SMase
activation versus that mediated via CS. Treatment with 25 µM FB1 did not alter the apoptotic response up to 8 h after irradiation (Fig. 5B). Thereafter, apoptosis began to slow and like ceramide generation was shut off after 12 h (Fig. 5B). These data suggest the existence of a late phase
apoptotic response mediated via CS activation, the beginning of which
coincides with post-radiation activation of CS (Fig.
4B).
Since these studies indicated that radiation may activate two distinct
mechanisms for generation of ceramide, we investigated SMase and CS
responses to ionizing radiation in other cell types, and we compiled
this information with already published data from our laboratory.
Evaluation of SMase activity was carried out at multiple time points
between 30 s and 120 min after irradiation and of CS activity
between 30 min and 24 h, as detailed under "Materials and
Methods." Fig. 5C shows that some cells activate CS
selectively (AT1ABR and TPA-treated LNCaP cells), whereas other cells
activate only SMase (GM0988 human B cell and C3H/HeJ mice lung
endothelium), neither enzyme (C3ABR, 1418 NPD, A431, and LNCaP cells),
or both (BAEC). In instances where radiation stimulated ASMase selectively, such as in the wild type B cell line
GM0988 or the murine lung endothelium (42), our present studies
disclosed no activation of CS within the first 24 h of irradiation
with doses (10-20 Gy) previously shown to produce maximal apoptotic responses in these cells (64). Furthermore, genetic inactivation of
ASMase, such as occurred in 1418 NPD cells or in the ASMase knock-out
mouse, abrogated apoptosis, whereas restoration of ASMase activity by
retroviral transduction restored radiation-induced ceramide generation
and apoptosis (42). Alternatively, in cells where CS activation
predominated (AT1ABR and TPA-treated LNCaP cells), increased SMase
activity was not detected within 120 min of doses up to 20 Gy, whereas
FB1 uniformly inhibited radiation-induced ceramide generation and
apoptosis (Figs. 3D, 5B, and 6F). In
C3ABR, 1418 NPD human B cells and A431 cells there was no evidence of SMase or CS activation, or apoptosis, with any dose up to 20 Gy. In
human prostate LNCaP cells, radiation with doses up to 50 Gy failed to
activate SMase or CS, generate ceramide, or induce apoptosis within
48 h of treatment. However, in cells pretreated with 10 ng/ml TPA,
10 Gy induced a 1.5-fold increase in CS activity from base line of 156 pmol/mg prot/min by 3 h, a 50-100% increase in ceramide levels
from a base line of 650 pmol/106 cells by 6 h, and
induction of apoptosis beginning at 12 h. These studies indicate
that signaling through CS and/or SMase is cell type-specific and
suggest that these mechanisms, when activated, may play a crucial role
in the induction of apoptosis in response to x-irradiation.
ATM Inhibits Radiation-induced Ceramide Synthase Activation and
Apoptosis--
As discussed in the Introduction, ATM is involved in
several responses to radiation-induced DNA damage, one of which appears to be the inhibition of radiation-induced apoptosis. Since the CS-mediated mechanism of radiation-induced apoptosis constitutes a
response to DNA damage, we investigated whether ATM might regulate this
process. EBV-transformed peripheral blood lymphoblastoid lines from a
normal adult donor (C3ABR) and a homozygous AT patient (AT1ABR) (35)
were used for these initial experiments. Exposure of C3ABR cells to
doses of up to 10 Gy failed to induce an apoptotic response as measured
by bisbenzimide staining (Fig.
6A), whereas AT1ABR cells
exhibited both time- and dose-dependent apoptosis (Fig.
6B). When treated with a dose of 10 Gy, apoptosis was
evident by 16 h and peaked at 36 h. Consistent with these
data, irradiation of AT1ABR cells with 5 and 10 Gy induced 6 ± 1- and 8 ± 1-fold increases in caspase activity, respectively, over
unirradiated AT1ABR cells at 24 h. Similar elevations were
detected for as long as 36 h post-irradiation. In contrast, in
wild type C3ABR cells, irradiation had no effect on caspase activity at
any time from 24 to 36 h. Fig. 6C shows that exposure
of C3ABR cells to 10 Gy did not induce CS activation. In contrast to
AT1ABR cells, 10 Gy induced a significant increase of CS activity (Fig.
6D), from a base line of 86-146 pmol/min/mg protein at
20 h (p < 0.05) without a significant change in
the Km of the reaction. The peptide caspase
inhibitor benzyloxycarbonyl-Val-Ala-Asp fluoromethylketone (40 µM) abolished apoptosis as measured by caspase activation or morphology but was without effect on CS activation, indicating CS
stimulation is not a consequence of induction of apoptosis. Concomitant
with CS activation, there was a time-dependent rise of
cellular ceramide in AT1ABR but not in C3ABR cells (Fig. 6E) and was noted initially at 8 h and peaked at 24 h (data not
shown). To confirm a role for CS activation in ceramide generation and apoptosis in AT1ABR cells, the effects of FB1 were explored. Fig. 6E shows that at 24 h, there was a 6-fold increase of
ceramide that was almost completely blocked by 50 µM FB1.
The apoptotic response was also inhibited by 50 µM FB1,
which was non-toxic in these cells (Fig. 6F). There was no
significant effect of FB1 on cellular ceramide and apoptosis in C3ABR
cells. These data suggest, although do not provide definitive evidence,
that ATM suppresses the CS-mediated mechanism of radiation-induced
apoptosis.
To determine whether CS activation is a general characteristic of the
AT phenotype of B cells, we compared three normal lines to six AT lines
derived from patients with different AT mutations (Table
I). All six AT lines displayed
radiation-induced ceramide synthase activation and apoptosis, whereas
none of the normal lines manifested CS activation or significant
apoptosis. For the most part, the AT lines that displayed brisk CS
responses showed greater apoptosis.
To explore further this hypothesis, we investigated whether
reconstitution of ATM function would revert the CS response to radiation in AT cells. We used AT1ABR cells stably transfected with the
pMAT1 vector, which contains a full-length ATM cDNA
plasmid under the regulation of a metallothionein II promoter (51). Recent studies demonstrated that expression of ATM in these cells improved the survival, reduced chromosomal aberrations, and partially corrected defective cell cycle check points after radiation (51). In
the present studies, pMAT1-transfected AT1ABR cells were irradiated with 4 Gy, and the promoter was activated with 4 µM
CdCl2, and the cells were evaluated for apoptosis at
24 h after irradiation. Expression of ATM, which was confirmed by
Western blot (Fig. 7A, inset),
resulted in a 50% reduction in the apoptotic response as compared with
pMAT1-transfected AT1ABR cells not stimulated with CdCl2 or
AT1ABR cells transfected with an empty vector (data not shown). To
assess the effect of ATM on CS activation, pMAT1-transfected AT1ABR
cells were irradiated with a dose of 10 Gy, and 16 h later microsomal membranes were prepared and subjected to the CS assay using
10 µM sphinganine as substrate. Fig. 7A shows
that in the absence of induction with CdCl2, radiation
induced, as in naive AT1ABR cells, a 1.8-fold increase in the maximal
velocity of CS in cells stably transfected with empty vector or pMAT1
(p < 0.005 each versus control). 5 µM CdCl2 alone did not significantly affect base-line CS activity in AT1ABR cells (data not shown). However, ATM
expression stimulated by CdCl2 prevented radiation-induced CS activation almost entirely (10 Gy + CdCl2 in
ATM-transfected cells p < 0.005 versus 10 Gy + CdCl2 in cells transfected with vector only). Similar
effects of the reconstitution of ATM function on radiation-induced CS
activation and apoptosis were observed in pMAT1-transfected AT3ABR
cells (data not shown).
A converse pattern was observed in normal C3ABR cells that were
converted to an AT-like phenotype by stable transfection with an
antisense construct of ATM cDNA, as recently described
(52). The vector used for transfection (pMAT2) contains an antisense construct of full-length ATM cDNA under the control of a
metallothionein II promoter. Induction of antisense ATM
leads to loss of 70-80% of the endogenous ATM protein by 9 h
(Fig. 7B, inset) and to increased radiation sensitivity
(52). pMAT2-mediated inactivation of ATM with 5 µM
CdCl2 was associated with significant CS activation 16 h after exposure to 10-Gy irradiation (10 Gy + CdCl2 in
anti-ATM-transfected cells, p < 0.005, versus 10 Gy + CdCl2 in cells transfected with vector only; Fig. 7B) and an increase in
apoptosis at 24 h from the 6 ± 2% in irradiated control
cells to 24 ± 1% (p < 0.05 versus
control) (data not shown). In C3ABR cells, CdCl2 alone did
not affect base-line CS activity, caspase activity, or apoptosis which
represented 8 ± 1% of the population as measured by bisbenzimide staining (data not shown). Together these data provide compelling genetic and biochemical evidence supporting the concept that ATM regulates CS activation, ceramide generation, and apoptosis in response
to DNA damage.
The present studies provide evidence that radiation-induced DNA
damage signals apoptosis via post-translational activation of CS. The
dose dependences for CS activation, ceramide generation, and apoptosis
correlated closely. Inhibition of apoptosis using a peptide caspase
inhibitor did not affect CS activation, indicating this event is not a
consequence of induction of apoptosis. An obligatory role for CS
activation in this pathway was defined by the use of FB1 that blocked
[125I]dURd- and x-radiation-induced ceramide elevation
and apoptosis. The pro-apoptotic signals that activate CS are regulated
by ATM. Not only do immortalized B cells from AT patients exhibit
increased radiation-induced CS activation, ceramide generation, and
apoptosis but stable transfection of wild type ATM reversed
these events. Furthermore, antisense inactivation of ATM in normal B
cells recapitulated the AT phenotype. Thus, signals from DNA damage and
ATM appear to reciprocally regulate the activity of CS and apoptosis
after radiation exposure.
The prevalence of the pro-apoptotic responses of CS and SMase to
radiation is unknown. Our data indicate that the CS response is cell
type-specific. Some cells, such as the AT1ABR and TPA-treated LNCaP
cells, activate CS selectively, whereas others, such as BAEC, activate
it coordinately with SMase. The data define a number of issues. We
examined two pairs of EBV-immortalized human B cell lines generated in
different laboratories. The C3ABR line, derived from peripheral blood
of a normal adult (35), activated neither SMase nor CS and was
radioresistant. Genetic inactivation of ATM in the sister
line AT1ABR conferred both CS activation and radiosensitivity but did
not confer SMase activation. In contrast, the wild type B cell line
GM0988, derived from the peripheral blood of an 8-year-old (42),
responded to ionizing radiation with SMase activation and apoptosis but
not CS stimulation. Inactivation of ASMase in its sister line 1418 derived from a child with NPD did not permit radiation-induced CS
activation but rather resulted in radioresistance. These studies
suggest that radiation-induced ASMase activation, which occurs at the
plasma membrane, and CS activation, which results from DNA damage, are
independent stress response systems. Furthermore, these studies suggest
that cell lines developed from the same apparent source may respond
differently depending on developmental contexts or the
microenvironmental milieu. Although it is possible that selection of
specific B cell populations occurred due to the ages of the donors,
more likely the differences reflect known clonal variations associated
with strain-specific differences in EBV gene expression (59-61).
Similarly, we noted differences between the radiation responses of
primary cultures of bovine macrovascular endothelium in
vitro and pulmonary microvascular endothelium in vivo.
Whereas the former appeared to utilize both SMase and CS, the latter
used only SMase. These systems also appear subject to transmodulation.
Whereas LNCaP cells were noted to be radioresistant and to activate
neither SMase nor CS, low doses of the phorbol ester TPA conferred
radiation-induced CS activation, ceramide generation, and apoptosis
onto these cells. Ultimately, it would appear necessary to understand
cell type-specific signaling in tissues in vivo if
pharmacologic intervention is to be successful. Nonetheless, the
description of generic stress response mechanisms involved in the
decision to live or die would appear to be a prerequisite for devising
future pharmacologic strategies.
The biosynthetic pathway of ceramide synthesis involving CS occurs in
cytosolic organelles, as CS was shown to localize in mammalian cells at
the endoplasmic reticulum and the mitochondrial membrane (43, 44).
Consistent with this observation, we found no CS activity in isolated
control or irradiated nuclei from
BAEC.3 Thus, damaged DNA must
transmit signals from the nucleus to cytoplasmic organelles to initiate
the death program, as radiation-induced CS activation appears
independent of transcription and translation. The nature of these
signals or the post-translational modifications to CS remains unknown.
Whether the localization of CS at mitochondrial membranes is relevant
to the evolution of CS-mediated apoptosis is the topic of ongoing
studies. In this regard, ceramide has been shown to induce cytochrome
c release from mitochondria (62), permeability transition
inhibitable by Bcl-2 (63), and generation of reactive oxygen species by
inactivation of complex III (64, 65).
Although it is not presently possible to molecularly order CS
activation within the myriad events that encompass ATM action, the
delineation of it as transcriptionally independent defines a select
group of potential ATM partners, such as those involved in the system
that surveys and repairs DNA damage. One of the targets for ATM is the
nuclear protein p53 (32, 66, 67). Since the interaction between ATM and
p53 is complex and apparently cell type-specific, it is currently not
possible to define a link between ATM, CS, and p53. CS is not likely to
play a role in radiation-induced apoptosis in thymic cells, since ATM
signals p53/p21-mediated G1 arrest in these cells (29, 33,
68, 69) but not apoptosis (38). Alternatively, CS could be involved in
the spontaneous apoptosis that occurs in meiotic spermatogonia lacking
ATM, an event abrogated by loss of p53 (39). p21 might also play a role in CS activation. Leder and co-workers (70) demonstrated that loss of
p53 accelerated tumorigenesis in ATM knock-out mice, whereas loss of p21 led to reduced tumor size. Pathologic examination revealed
that the ATM/p21 double knock-out tumors underwent
substantial spontaneous apoptosis. The tumors were also remarkably
sensitive to radiation-induced apoptosis. These studies suggest that
p21 restrains apoptosis in ATM-deficient mice and may be required for
tumors to grow unabated. Whether the p21 knock-out tumor phenotype predisposes cells to greater increases in CS activity or enhances the
response to equimolar ceramide elevations is at present unclear.
In summary, the present investigations link direct DNA damage to
apoptotic signaling through CS. This coupling requires activation of an
unknown translationally independent signal from the nucleus to the
cytoplasmic compartment, which is suppressed by ATM. Whether failure to
respond to x-irradiation with CS activation and apoptosis may be a
manifestation of strict ATM regulation and whether it represents a
clinically relevant component of radiation resistance are presently
uncertain. The present studies may also provide a target to address the
mechanisms by which other DNA damage/repair proteins such as p53,
DNA-protein kinase, or c-Abl affect the apoptotic response. Integration
of the ATM/CS pathway into the set of events that regulate DNA damage
responses holds the promise of understanding mechanisms involved in the
decision of cells to repair DNA breaks or to die.
INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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80 °C overnight and subsequently
stored at
180 °C in liquid nitrogen for 1-30 days. Prior to
experiments, cells were thawed by incubation in a 38 °C water bath
and subsequently plated in culture media as described above.
Endothelial cells appear resistant to the freeze and thaw procedure
employed, since after thawing >95% of the cells were recovered, and
the viability according to the trypan blue exclusion test was
>98%.
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180 °C under liquid nitrogen for 1-30 days. The
cells were then thawed, washed, and plated in Petri dishes in 10% calf
serum in DMEM and incubated at 37 °C for 24 h. BAEC appear
resistant to freeze and thaw as more than 95% of control cells (either
untreated or those treated with non-radioactive IdURd) were recovered
after thawing, and their viability by trypan blue exclusion was >98%. Control cells cultured after thawing produced monolayers that appeared
normal by morphology and growth kinetics (data not shown).
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Fig. 1.
Effect of [125I]dURd
incorporation on apoptosis and cellular ceramide levels in BAEC.
Pre-confluent BAEC were metabolically labeled with
[125I]dURd and stored in suspension at 180 °C in
liquid nitrogen as described under "Materials and Methods." At the
indicated times, frozen cells were thawed, washed, and plated in Petri
dishes in 10% calf serum/DMEM at 37 °C. A shows the
apoptotic response at 24 h after thawing, quantified by staining
with the DNA-specific fluorochrome bisbenzimide. Control cells in this
experiment were not exposed to IdURd treatment. However, control cells
treated with non-radioactive IdURd (0.25 nM) also failed to
exhibit apoptosis (data not shown). 500 cells were scored for each data
point. Each value represents the mean of duplicate determinations from
one of three similar experiments. B shows the levels of
cellular ceramide. Ceramide was quantified by the DAG kinase method as
described under "Materials and Methods" at 16 h after thawing.
Each value represents the mean of triplicate determinations from one of
five similar experiments.
180 °C for 15 days, the concentration of
ceramide immediately after thawing was 225 ± 0.2 pmol/106 cells, a value similar to that reported in BAEC
under standard growth conditions (41). Small elevations of ceramide
were noted in [125I]dURd-treated cells as early as 4-5 h
after thawing, peaking at 16 h (data not shown). The peak level of
ceramide was proportional to the duration of [125I]dURd
treatment. At 16 h after thawing, there was a 6.5-fold increase in
the ceramide level over control in cells treated with [125I]dURd for 15 days, and for cells treated for 24 days, the fold increase was 7.7.
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Fig. 2.
De novo synthesis of ceramide and CS
activity in [125I]dURd-treated BAEC. Cells were
treated with [125I]dURd for 17 days as described under
"Materials and Methods." A, 106 cells were
plated after thawing on a 60-mm Petri dish in serum-free DMEM and
incubated at 37 °C for 12 h. [9,10-3H]Palmitic
acid was then added, and at the indicated times lipids were extracted
and subjected to mild alkaline hydrolysis as described under
"Materials and Methods." 3H-Labeled ceramide was
resolved by thin layer chromatography and quantified by liquid
scintillation counting. Nonspecific incorporation was determined at
0 °C and represented <10% of specific incorporation. Similar
results were observed in three different experiments. B,
microsomal membranes were prepared at 12 h after thawing, and CS
activity was measured as described under "Materials and Methods."
Lipids were extracted and dihydroceramide was resolved by thin layer
chromatography, detected by comigration with ceramide standard (Sigma
Type III), and quantified by liquid scintillation counting as described
under "Materials and Methods." Similar results were obtained in
five experiments. C, shows the Km and
Vmax from B by Eadie-Hofstee
analysis.
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Fig. 3.
Effects of cycloheximide or FB1 on
[125I]dURd-induced CS activation, ceramide elevation, and
apoptosis. A and B, effect of cycloheximide
on [125I]dURd-induced CS activation. BAEC were treated
with [125I]dURd for 17 days as described under Materials
and Methods," thawed, plated on Petri dishes, and incubated at
37 °C for 2 h to allow attachment of the cells to the culture
dish. There was no increase in base-line ceramide synthase activity
during this incubation period (data not shown). Distilled water
(diluent) (A) or 1 µg/ml (final concentration) of
cycloheximide (B) was then added. After 12 h at
37 °C, microsomal membranes were prepared and subjected to the CS
assay as in Fig. 2B. Similar data were observed when
cycloheximide was added immediately upon thawing (data not shown).
Values represent mean ± S.E. of determinations from three
separate experiments. C shows the effect of FB1 on
[125I]dURd-induced ceramide generation.
[125I]dURd-treated BAEC were prepared as described above.
FB1 (10-25 µM) was added to the cells immediately after
plating. After 16 h, cellular lipids were extracted and subjected
to the DAG kinase assay to quantify the ceramide level as described in
Fig. 1B. Each value represents the mean ± S.E. of
triplicate determinations from one of five similar experiments.
D shows the effect of FB1 on apoptosis. Apoptosis was
determined as in Fig. 1A at 24 h after thawing. Each
value represents the mean ± S.D. of duplicate determinations from
one of five separate studies.
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Fig. 4.
Measurement of CS activity in response to
external x-irradiation of BAEC. A, dose response. BAEC
were irradiated with the indicated doses of ionizing radiation. 12 h after irradiation, microsomal membranes were prepared and subjected
to the CS assay using 10 µM sphinganine as substrate, as
in Fig. 2B. Each value represents the mean ± S.E. of
triplicate determinations from three separate experiments.
B, time course. BAEC were exposed to 10 Gy of external
x-irradiation. At the indicated times after irradiation, microsomal
membranes were prepared and assayed for CS activity as described in
Fig. 2B. Similar results were obtained in five separate
experiments. C, determination of Km and
Vmax from B at 12 h utilizing
Eadie-Hofstee analysis.
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Fig. 5.
Effect of FB1 on ceramide generation and
apoptosis in BAEC treated with x-irradiation, and the effect of
x-irradiation on SMase and CS activation in different cell types.
A, effect of FB1 on ceramide elevation in BAEC. The media of
confluent BAEC were changed to fresh serum-free DMEM, and 25 µM FB1 was added to cells 1 h before irradiation.
After incubation of cells at 37 °C for the indicated times
(inset, short time course), ceramide levels were measured by
DAG kinase method as in Fig. 1B. Each value represents the
mean ± S.E. of triplicate determinations from five experiments.
B, effect of FB1 on radiation-induced apoptosis. BAEC were
exposed to ionizing radiation and treated with FB1 as in A
and morphological changes of nuclear apoptosis quantified as in Fig.
1A. Each value represents the mean ± S.E. of duplicate
determinations from seven independent experiments. C, survey
of radiation-induced SMase and CS activation in different cell types.
Cells were cultured and irradiated (10 Gy, except 20 Gy for lung
endothelium in vivo) as described under "Materials and
Methods." Activation of SMase was defined by a reduction in
sphingomyelin content coupled with quantitative elevation of ceramide.
CS assays were performed as under "Materials and Methods." LNCaP
cells were treated with 10 ng/ml TPA for 45 min prior to
irradiation.
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Fig. 6.
Effect of radiation and FB1 on CS activation,
ceramide generation, and apoptosis in normal and homozygous AT
lymphoblastoid cells. A and B show dose and
time effects of ionizing radiation on the induction of apoptosis in
normal (C3ABR) (A) and AT (AT1ABR) (B)
lymphoblasts. One hour before irradiation, the medium of the cultures
was changed to 0.1% human albumin/RPMI (5 × 105
cells/ml), and apoptosis was quantified as in Fig. 1A. Each
value represents the mean of duplicate determinations from one of three
similar experiments. C and D show the effect of
10 Gy on CS activity as described in the Fig. 2 at 20 h
post-irradiation in C3ABR (C) and AT1ABR (D)
lymphoblasts. Similar results were obtained in three independent
experiments. E shows effect of FB1 on radiation-induced
ceramide elevation in C3ABR and AT1ABR lymphoblasts. Cells were
pretreated with 50 µM FB1 for 1 h and irradiated
with 10 Gy, and after 24 h ceramide was quantified by the DAG
kinase assay as in Fig. 1B. Each value represents mean ± S.E. of triplicate determinations from one of three similar
experiments. F shows effect of FB1 on radiation-induced
apoptosis in normal and AT lymphoblasts. Cells were handled, treated
with 50 µM FB1, and irradiated with 10 Gy as in
E. 36 h after irradiation, cells were collected and
assessed for apoptosis as in A. Each value represents the
mean ± S.D. of duplicate determinations from one of three similar
studies.
CS activation and apoptosis induced by ionizing radiation in AT cell
lines
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Fig. 7.
Effects of sense or antisense ATM
cDNA expression on radiation-induced CS activation.
A, prevention of radiation-induced CS activation in AT1ABR
lymphoblasts by restoration of ATM activity. ATM was expressed in
AT1ABR cells stably expressing pMAT1 (containing ATM under
control of a metallothionein promoter) by stimulation with 5 µM CdCl2. After 9 h, the medium was
changed to 0.1% human albumin/RPMI (5 × 105
cells/ml), and cells were irradiated (10 Gy) as described under
"Materials and Methods." 16 h post-irradiation, microsomal
membranes were prepared and assayed for CS activity using 10 µM sphinganine as in Fig. 2B. Values represent
mean ± S.D. of duplicate determinations from one of three
independent experiments. Inset, ATM levels were detected by
Western blot 9 h after CdCl2 treatment as described
(51). B, antisense inactivation of ATM confers
radiation-induced CS activation onto C3ABR lymphoblasts. Endogenous ATM
was inactivated in normal lymphoblasts stably transfected with the
pMAT2 vector (containing an antisense ATM cDNA construct
under control of a metallothionein promoter) by stimulation with 5 µM CdCl2 for 9 h prior to irradiation.
16 h after irradiation, microsomal membrane was prepared and
assayed for CS activity as in A. Values represent mean ± S.D. of duplicate determinations from one of three independent
experiments. Inset, ATM levels were detected by Western blot
9 h after CdCl2 treatment as described (51).
DISCUSSION
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ABSTRACT
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FOOTNOTES |
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* This work was supported by Grant CA52462 from the National Institutes of Health.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.
§ Both authors contributed equally to this work.
** To whom correspondence should be addressed: Laboratory of Signal Transduction, Memorial Sloan-Kettering Cancer Center, 1275 York Ave., New York, NY 10021. Tel.: 212-639-8573; Fax: 212-639-2767; E-mail: R-Kolesnick{at}mskcc.org.
2 K. K. Khanna and M. Lavin, unpublished data.
3 R. Kolesnick and A. Morales, unpublished observations.
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ABBREVIATIONS |
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The abbreviations used are: dsb, double-stranded breaks; dURd, 5-iodo-2'-deoxyuridine; CS, ceramide synthase; PI3-K, phosphatidylinositol 3-kinases; BAEC, bovine aortic endothelial cell; SMase, sphingomyelinase; FB1, fumonisin B1; EBV, Epstein-Barr virus; ATM, ataxia telangiectasia-mutated gene product; DMEM, Dulbecco's modified Eagle's medium; PBS, phosphate-buffered saline; Gy, gray; AT, ataxia telangiectasia; TPA, 12-O-tetradecanyolphorbol-13-acetate DAG, diacylglycerol; DNA-PK, DNA-dependent protein kinase; Z, benzyloxycarbonyl; AFC, 7-amino-4-trifluoromethylcoumarin, ASMase, acid sphingomyelinase.
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