An Inducible Pathway for Degradation of FLIP Protein Sensitizes
Tumor Cells to TRAIL-induced Apoptosis*
Youngsoo
Kim,
Nanjoo
Suh
,
Michael
Sporn, and
John C.
Reed§
From The Burnham Institute, La Jolla, California 92037 and the
Department of Pharmacology, Dartmouth Medical School,
Hanover, New Hampshire 03755
Received for publication, March 13, 2002, and in revised form, April 6, 2002
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ABSTRACT |
TRAIL (Apo2 ligand) is a member of the tumor
necrosis factor (TNF) family of cytokines that induces apoptosis.
Because TRAIL preferentially kills tumor cells, sparing normal tissues,
interest has emerged in applying this biological factor for cancer
therapy in humans. However, not all tumors respond to TRAIL, raising
questions about resistance mechanisms. We demonstrate here that a
variety of natural and synthetic ligands of peroxisome
proliferator-activated receptor-
(PPAR) sensitize tumor but not
normal cells to apoptosis induction by TRAIL. PPAR ligands
selectively reduce levels of FLIP, an apoptosis-suppressing protein
that blocks early events in TRAIL/TNF family death receptor signaling.
Both PPAR agonists and antagonists displayed these effects,
regardless of the levels of PPAR expression and even in the presence
of a PPAR dominant-negative mutant, indicating a PPAR-independent
mechanism. Reductions in FLIP and sensitization to TRAIL-induced
apoptosis were also not correlated with NF-
B, further suggesting a
novel mechanism. PPAR modulators induced ubiquitination and
proteasome-dependent degradation of FLIP, without
concomitant reductions in FLIP mRNA. The findings suggest the
existence of a pharmacologically regulated novel target of this class
of drugs that controls FLIP protein turnover, and raise the possibility
of combining PPAR modulators with TRAIL for more efficacious
elimination of tumor cells through apoptosis.
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INTRODUCTION |
Considerable interest has emerged in the possibility of exploiting
the apoptotic effects of
TRAIL1 for the treatment of
cancer. TRAIL is a member of the tumor necrosis factor (TNF) family of
cytokines that is capable of inducing apoptosis (1). The
apoptosis-inducing receptors for TRAIL include Trail-R1 (DR4) and
Trail-R2 (DR5), which are transmembrane type I receptors expressed on
the surface of many types of cell. However, TRAIL also binds to
non-apoptosis-inducing decoy receptors, which compete with death
receptors for ligand and suppress apoptosis, including DcR1, DcR2, and
osteoprotegerin (reviewed in Refs. 2 and 3). Empiric analysis of the
effects of TRAIL on normal and malignant cells has provided compelling
evidence that recombinant TRAIL protein preferentially induces
apoptosis of cancer cells without harming most types of untransformed
cells (reviewed in Ref. 2). When properly prepared and purified,
recombinant trimeric TRAIL also lacks significant toxicity in primate
species that possess receptors capable of binding human TRAIL (4,
5).
Preclinical studies of recombinant TRAIL (extracellular domain) in mice
have demonstrated impressive anti-tumor activity and synergy with
cytotoxic anticancer drugs (6). However, not all tumors respond to
TRAIL. This lack of response may be attributed either to unfavorable
ratios of death and decoy receptors or because of intracellular
resistance mechanisms (3, 7-10). With respect to intracellular
resistance mechanisms, the FLIP protein has been identified as a
blocker of apoptosis induced by TNF family death receptors (reviewed in
Ref. 11). FLIP binds to and neutralizes adapter proteins and
pro-caspases normally recruited to the cytosolic domains of
apoptosis-inducing TRAIL receptors upon ligand stimulation, thus
interrupting early steps in TRAIL signaling. Furthermore, overexpression of FLIP protein has been documented in cancers (12).
PPAR is a member of the steroid/retinoid superfamily of
ligand-activated transcription factors. Agonistic ligands of PPAR include modified fatty acids, cyclopentenone-containing prostaglandins, triterpenoids, and the thiazolidinediones, a class of
insulin-sensitizing drugs used in the treatment of type II diabetes
(reviewed in Ref. 13). Anti-tumor properties of PPAR agonists have
been reported. For example, thiazolidinediones have been shown to
suppress the growth of human colon and breast cancer cell lines
in vitro and in vivo in the mouse (14, 15), and a
member of a new class of PPAR agonists (tyrosine analogs) suppresses
mammary carcinogenesis in a standard rat model (16). However,
troglitazone increases incidence of colonic polyps in a mouse model in
which one allele of adenomatous polyposis coli is inactive (17,
18), suggesting complex effects on neoplasia. Moreover, the
concentrations of thiazolidinediones required for some apoptotic
effects are beyond clinically attainable ranges (15).
Here we explored the effects of PPAR ligands on TRAIL-induced
apoptosis in epithelial cancers cell lines. Our findings demonstrate a
new PPAR-independent mechanism for these compounds, resulting in
rapid decreases in FLIP protein without concomitant reductions in
c-FLIP mRNA, and causing sensitization of tumor but not several types of normal cells to TRAIL-induced apoptosis. The mechanism invoked by these PPAR modulatory compounds involves ubiquitination and proteasome-dependent degradation of FLIP, thus revealing the existence of an inducible pathway for reducing FLIP expression that
might be exploited for promoting death receptor-induced apoptosis of
neoplastic cells.
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MATERIALS AND METHODS |
Cell Cultures--
Human prostate cancer, PPC-1 and LNCaP,
ovarian cancer, OVCAR-3, and SK-OV-3 cells were cultured in RPMI 1640 (Irvine Scientific, Santa Ana, CA) supplemented with 10% fetal bovine
serum (HyClone, Tulare, CA), 1 mM L-glutamine,
and antibiotics. Dulbecco's modified Eagle's medium (Invitrogen)
containing the same additives was used for HT29, COLO205 colon cancer,
and HeLa cervical cancer cells. Cynomologus monkey hepatocytes were
obtained from Cedra (Austin, TX). The TRAIL receptors of this species
have been reported to be 84-99% identical in amino acid sequence to
their human counterparts, and these monkey-derived cells have been
shown to exhibit essentially identical apoptotic responses to
recombinant preparations of TRAIL as their human counterparts (5).
Human umbilical vascular endothelial cells were purchased from
Clonetics (Walkersville, MD). Prostaglandins and ciglitazone were
obtained from Cayman Chemical (Ann Arbor, MI) and Biomol (Plymouth
Meeting, PA), respectively, and dissolved in Me2SO
before use. Troglitazone was provided from Sankyo (Tokyo, Japan). CDDO
and CDDO-Me were synthesized and employed in cultures as described
previously (19, 20). Human recombinant TNF and TRAIL were purchased
from R & D Systems (Minneapolis, MN) and Biomol, respectively. The
protease inhibitors Z-VAD-fmk, lactacystin, MG132, TLCK, and calpeptin
were obtained from Calbiochem. MG-115 and epoxomicin were purchased
from Sigma and Alexis (San Diego, CA), each.
Transfections--
PPC-1 or HeLa cells were transiently
transfected using LipofectAMINE Plus (Invitrogen) following the
recommended protocol. For luciferase reporter assays, Tk-PPRE3-Luc
plasmid (0.5 µg/35-mm well) was transfected into cells along with
pCMV-
-galactosidase plasmid (0.2 µg/well). Cells were then
cultured for 16 h in complete media and treated with various
ligands for the indicated times. To reduce background activity, the
usual medium was exchanged with medium containing 0.1% fetal bovine
serum prior to ligand treatment. Luciferase activity was assayed with
the luciferase assay system from Promega (Madison, WI) and measured
with a Luminometer (EG & G Berthold). The activity of -galactosidase
was determined by incubating lysates with 0.7 mg/ml
o-nitrophenyl--D-galactopyranoside (Sigma) at
37 °C for 10 min and measuring absorbance at 405 nm. For
apoptosis experiments, cells in 35- or 60-mm dishes were co-transfected with 1 µg of pEGFP-N2 (CLONTECH, Palo Alto, CA)
and 4 µg of plasmid DNA encoding IKK, c-FLIP, CrmA, Bcl-2,
PPAR, and dominant-negative PPAR (21, 22). The total amount of
DNA per transfection was normalized by addition of empty plasmid DNA.
For antisense experiments, FLIP antisense, ISIS 23296 (ACTTGTCCCTGCTCCTTGAA), or control oligonucleotide, ISIS 132383 (AGTTCTCTCTGCCCCTAGAT), was delivered into cells by lipofection at a
final concentration of 300 nM, as described (23). Antisense
oligonucleotides were chimeric molecules in which the first and last 5 nucleotides in the sequence were 2'-O-methoxyethyl-modified,
with the center 10 nucleotides composed of natural ribose moieties to
support RNase H activity (24). The oligonucleotides linkages contained
phosphorothioate modifications to enhance nuclease resistance.
Caspase Assays--
Cell extracts were prepared, normalized for
total protein content, and incubated with 100 µM
Z-DEVD-AFC (Enzyme Systems Products, Livermore, CA) in caspase assay
buffer for measuring caspase-mediated release of AFC using a
fluorimeter, as described (25).
Cytotoxicity and Apoptosis Assays--
MTT assays were performed
using a kit from Roche Molecular Biochemicals (MTT-based Cell
Proliferation Kit 1) following the suggested protocol. In brief, 5 × 104 cells were seeded in 96-well plates and cultured for
48 h prior to treatment. Cells were then treated with various
concentrations of PPAR modulators with or without 25-250
(ng/ml) TRAIL for 24 h. MTT was added for 4 h at
37 °C, and absorbance at 550 nm was measured using a microtiter
plate reader. Apoptosis was determined by fixing cells in 3.7%
paraformaldehyde and staining with 0.1 µg/ml
4,6-diamidino-2-phenylindole (DAPI), scoring the percentage of cells
having intensely condensed chromatin and/or fragmented nuclei
by UV microscopy (n = 200).
Electrophoretic Mobility Shift Assays (EMSAs)--
Nuclear
extracts were prepared from cells, and EMSAs were carried out as
described previously (26). In brief, oligonucleotides containing a
consensus NF-B-binding site, 5'-AGTTGAGGGGACTTTCCCAGGC-3' (Promega)
were end-labeled with [-32P]ATP (PerkinElmer Life
Sciences) using T4 polynucleotide kinase (Amersham Biosciences). After
purification with MicroSpin G-25 columns (Amersham Biosciences), the
labeled probe (15 fmol) was incubated with 3 µg of nuclear extract
for 25 min at room temperature and separated by electrophoresis in
nondenaturing 5% polyacrylamide gels in 0.25× TBE (22.5 mM Tris borate, 0.5 mM EDTA) at 4 °C. After
drying, gels were exposed to x-ray film at 
70 °C.
Immunoblotting and Immunoprecipitations--
Whole cell lysates
prepared with RIPA buffer or cytosolic extracts prepared as above were
subjected to SDS-PAGE and transferred to nitrocellulose membranes
(Bio-Rad). Immunoblotting was performed with the following antibodies:
anti-caspase 8 at 1:3000 (v/v) (21) or at 1:1000 from Alexis; anti-PARP
(BD PharMingen, La Jolla, CA) at 1:1000; anti-FLIP (NF-6) at 1:20 (27);
anti-DR5 (Alexis) at 1:500; anti-DR4 (Millennium, Boston, MA) at 1:500; anti-RIP (BD PharMingen) at 1:200; anti-PPAR (Santa Cruz
Biotechnology Inc.) at 1:200; anti-TRADD (Santa Cruz Biotechnology
Inc.) at 1:250; anti-FADD (BD PharMingen) 1:1000; anti-DcR1 (ProSci
Inc., Poway, CA) at 1:500; anti-DcR2 (Calbiochem) at 1:1000; anti-DAP3 (Transduction Laboratories, San Diego, CA) at 1:500; anti-ubiquitin (Santa Cruz Biotechnology Inc.) at 1:200, and anti-
-tubulin (Sigma) at 1:1000 in 0.05% Tween/Tris-buffered saline (T-TBS) blocking buffer
containing 5% nonfat skim milk for 1-3 h at room temperature, followed by washing with T-TBS for 30 min. Goat anti-rabbit or anti-mouse IgGs coupled with horseradish peroxidase (Bio-Rad) were used
as secondary antibodies at 1:3000 (v/v). Immunospecific bands were
detected by using an enhanced chemiluminescence (ECL) detection system
(Amersham Biosciences).
For immunoprecipitation experiments, OVCAR-3 cells were treated with
either CDDO (5 µM) or TRAIL (100 ng/ml) alone or both for
2 h. Cells were washed with ice-cold phosphate-buffered saline once and lysed in 1 ml of lysis buffer (1% Triton X-100, 150 mM NaCl, 10% glycerol, 20 mM Tris-HCl (pH
7.5), 2 mM EDTA, 0.5 mM sodium orthovanadate,
10 mM -glycerophosphate, and protease inhibitor mixture)
at 4 °C for 30 min. In parallel, 10% of cell pellets were lysed in
RIPA buffer for immunoblot analysis. After centrifugation at
13,000 × g for 15 min, the lysates were mixed with 20 µl of anti-DR5 antibody (Alexis) and 50 µl of protein-A-Sepharose
at 4 °C for 6 h. The immune complexes (immunoprecipitations)
were washed four times with the lysis buffer, and the samples from immunoprecipitations and total lysates were subjected to
SDS-PAGE/immunoblot analysis using antibodies specific for caspase 8 at
1:1000 (Alexis), FADD at 1:1000 (BD PharMingen), and c-FLIP (NF-6) at
1:20 (v/v).
In Vitro Kinase Assays--
Cells were pretreated with PPAR
ligands for 15 min and then treated with TNF (20 ng/ml) for 20 min.
Cytosolic extracts (200 µg) prepared as above were incubated with 2 µg of anti-IKK (BD PharMingen) for 2 h, and 30 µl of
protein A-Sepharose (Sigma) was added overnight at 4 °C.
Immunoprecipitates were washed three times (1% Nonidet P-40, 0.5 M NaCl, 1 mM EDTA, 1 mM
dithiothreitol, 0.5 mM sodium orthovanadate, 5 mM NaF, and 50 mM HEPES (pH 7.4), containing
protease inhibitor mixture (Roche Molecular Biochemicals)) and washed
once more with the kinase buffer (20 mM HEPES (pH 7.4), 2 mM MgCl2, 2 mM MnCl2, 1 mM dithiothreitol, 0.5 mM sodium orthovanadate, 2 mM NaF, 10 mM -glycerophosphate,
containing protease inhibitor mixture). Kinase assays were performed in
25 µl of kinase buffer containing 10 µM ATP, 10 µCi
of [-32P]ATP, and 1 µg of GST-IB (Santa Cruz
Biotechnology Inc., Santa Cruz, CA) for 30 min at 30 °C. The samples
were subjected to 12% SDS-PAGE. Gels were dried and exposed to x-ray film.
Metal Affinity Capture--
SK-OV-3 cells were co-transfected
with plasmids encoding FLIP and His6-ubiquitin (pCW7).
After 18 h, cells were treated with 5 µM CDDO for
4 h in the absence or presence of 400 nM epoxomicin and lysed in 1 ml of GTN buffer (6 M guanidine HCl, 20 mM Tris-HCl (pH 8.0), 200 mM NaCl, 20 mM imidazole, and 0.1% Triton X-100). His6-tagged ubiquitin was recovered from lysates by
incubation for 30 min at 25 °C with 80 µl of cobalt chelate resin
(CLONTECH). Captured proteins were eluted in 20 mM Tris-HCl (pH 8.0), 200 mM NaCl, 200 mM imidazole, and 0.1% Triton X-100 after 3 washes of GTN
buffer containing 10 mM imidazole, and analyzed by
SDS-PAGE/immunoblotting using anti-FLIP antibody.
RNase Protection Assays--
PPC-1 or OVCAR-3 cells were treated
with the ligands for 1 and 6 h, and then total RNA was isolated
with TRIzol Reagent (Invitrogen) according to the manufacturer's
protocol. The probe for c-FLIP was synthesized by using the human Apo3b
Multi-Probe template set (BD PharMingen) with
[-32P]UTP and hybridized with 10 µg of RNA at
56 °C overnight. After RNase treatment, the protected transcripts
were separated by denaturing PAGE (5%). Gels were dried and exposed to
x-ray film at 70 °C.
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RESULTS |
PPAR Modulators Increase Sensitivity of Tumor Cell Lines to
TRAIL--
To explore preliminarily the effects of PPAR modulators
on tumor cell responses to TRAIL, we contrasted the effects of three different classes of PPAR ligands as follows: (a)
15d-PGJ2, a natural cyclopentenone prostaglandin having
PPAR agonist activity; (b) ciglitazone (Cig) and
troglitazone (Tro), representing synthetic thiazolidinediones PPAR
agonists; and (c) the triterpenoids CDDO and CDDO-Me, which
function as a weak agonist and as an antagonist of PPAR,
respectively (28). When tested against a variety of solid tumor cell
lines (PPC-1, PC3, OVCAR-3, SK-OV-3, HT29, COLO205, LNCaP, HeLa, HEY3,
and HT1080) at concentrations of 
1 µM, most of these
PPAR modulators had little effect on cell viability, as measured by
MTT dye reduction assays (Fig. 1 and data
not shown). For example, relative numbers of viable cells were within
80% of control following 1 day of treatment with 15d-PGJ2,
Cig, Tro, CDDO, or CDDO-Me at 1 µM, although
cytotoxic activity was observed at higher concentrations (5-25
µM). An exception was CDDO-Me, which exhibited anti-tumor
activity at 1 µM in occasional tumor lines (data not
shown). The sensitivity of these tumor cell lines to TRAIL was also
tested. When treated with TRAIL, 
80% of the tumor cells remained
viable after 1 day. In fact, of 10 tumor lines treated with 100 ng/ml
TRAIL, only one was significantly inhibited (not shown). Thus, neither
PPAR modulators nor TRAIL by themselves was generally effective at
killing tumor cells. However, combined treatment of epithelial cancer
cell lines with TRAIL and PPAR modulators resulted in synergistic
reductions in relative numbers of viable cells. Fig. 1 shows examples
for several PPAR modulators and tumor cell lines. Synergy was
observed both when holding the concentration of TRAIL fixed (100 ng/ml) and varying the concentration of PPAR modulators and, conversely, when holding the concentration of PPAR modulators fixed and varying TRAIL (Fig. 1). In contrast to tumor cell lines, the combination of
TRAIL and PPAR modulators was not cytotoxic to normal cells, including primary cultures of hepatocytes, endothelial cells, peripheral blood leukocytes, and bone marrow (Fig. 1C and
data not shown).
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Fig. 1.
Effect of PPAR
ligands on cell viability. Cell viability was measured by
MTT assay and expressed as % relative to control cultures. Data
represent mean ± S.D. of triplicate cultures and are
representative of 2-5 independent experiments. A, human
cancer cells OVCAR-3, COLO205, HT29, SK-OV-3, PPC-1, and LNCaP were
plated in 96-well plates 48 h prior to treatment (5-6 × 104/well) and then treated with the indicated
concentrations of CDDO for 24 h with (closed circles)
or without TRAIL (open circles) at either 25 (COLO205 and
PPC-1), 100 (OVCAR-3, HT29, and SK-OV-3), or 250 ng/ml (LNCaP).
B, SK-OV-3 cells were treated with the indicated
concentrations of ligands for 24 h with (closed
circles) or without TRAIL (open circles) (100 ng/ml).
C, SK-OV-3 cells, hepatocytes from cynomologus monkeys and
human umbilical vascular endothelial cells (HUVEC) were
treated with CDDO (0.5 µM) or troglitazone (20 µM) along with the increasing amounts of TRAIL.
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To explore whether these results were attributed to induction of
apoptosis, tumor cell lines were treated with TRAIL in combination with
PPAR modulators, and apoptosis was measured by DAPI staining (counting percentages of cells with apoptotic nuclear morphology, as
determined by chromatin condensation and nuclear fragmentation). Also,
caspase activity was measured in cell lysates (based on cleavage of the
fluorigenic caspase substrate, Ac-DEVD-AFC), and cleavage of the
caspase substrate poly(ADP-ribose) polymerase (PARP) was monitored (by
immunoblotting). Fig. 2 shows some
representation data. Note that tumor lines such as prostate cancer
PPC-1, ovarian cancer OCVAR-3, and cervical cancer HeLa are triggered
by the combination of PPAR modulators and TRAIL to undergo apoptosis (Fig. 2, A and B), activate caspases (Fig.
2C), and cleave the caspase substrate PARP (Fig.
2D). In contrast, culturing these cells with either TRAIL or
PPAR modulators alone had little effect. The broad spectrum
caspase inhibitor, Z-VAD-fmk, potently suppressed TRAIL-induced cell
death (not shown), further supporting an apoptotic mechanism. The
synergistic effect of TRAIL and PPAR modulators was not attributable
to a mere shift in the kinetics of apoptosis induction, as demonstrated
by time course analysis (Fig. 2E).
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Fig. 2.
Synergistic induction of apoptosis of
tumor cells treated with combination of TRAIL and
PPAR modulators. PPC-1 (A) and
OVCAR3 (B) cells were treated with 15d-PGJ2 (10 µM) or CDDO (5 µM) for 6 h in the
absence or presence of TRAIL (100 ng/ml). Apoptotic cells were counted
by DAPI staining (%) (mean ± S.D.; n = 3). Data
are representative of 2-3 experiments. C, lysates from
OVCAR-3 cells treated as above were prepared and assayed for caspase
activity, measuring release of the AFC fluorophore from the caspase
substrate DEVD-AFC. Activity is shown as the fold induction over
control, based on measurements of enzyme rates as described (54).
D, HeLa cells were treated with 15d-PGJ2 (10 µM), ciglitazone (20 µM), or CDDO (5 µM) in the absence () or presence (+) of TRAIL (100 ng/ml) for 6 h. Cell lysates were immunoblotted with antibodies
for FLIP and PARP. The arrows indicate full-length (~110
kDa) and cleaved ( 85 kDa) forms of PARP. C, control and
untreated cells. E, OVCAR-3 cells were treated with CDDO
(0.5 µM) for indicated times in the presence or absence
of TRAIL (100 ng/ml). Apoptotic cells were counted by DAPI staining
(%) (mean ± S.D.; n = 3).
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TRAIL Sensitization by PPAR Modulators Is Not Mediated by
Effects on NF-B--
In addition to their effects on PPAR
activity, several PPAR modulators have been reported to directly
inhibit the IB kinases, IKK and IKK, thereby suppressing
NF-B induction (29). Because NF-B plays important roles in
suppressing apoptosis induced by TNF, affecting expression of
apoptosis-suppressing genes (30, 31), we explored whether inhibition of
IKK activity or suppression of NF-B induction correlated with the
ability of various PPAR modulators to sensitize tumor cells to
TRAIL.
For IKK activity assays, tumor cell lines such as PPC-1 were treated
for 15 min with four different PPAR modulators which we had
demonstrated sensitize tumor cells to TRAIL, including 15d-PGJ2, ciglitazone, CDDO, and CDDO-Me. Because TRAIL did
not induce significant IKK activation (not shown), TNF was added to cultures for 20 min to stimulate IKKs. Cells were then lysed, and
IKK was recovered by immunoprecipitation, measuring its activity by
in vitro kinase assay using GST-IB as an in
vitro substrate. Loading of equal amounts of IKK protein was
confirmed by immunoblotting. Although 15d-PGJ2 was a potent
inhibitor of IKK activation, consistent with previous reports (29,
32), and CDDO reduced IKK activity by about half, the other PPAR
modulators ciglitazone and CDDO-Me had no effect on IKK activity
(Fig. 3A). Thus, suppression
of IKK activity cannot explain the ability of ciglitazone and
CDDO-Me to sensitize tumor cells to TRAIL.
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Fig. 3.
Inhibition of IKK and
NF-B activity by PPAR
ligands does not correlate with sensitization to TRAIL-induced
apoptosis. A, PPC-1 cells were pretreated with PPAR
ligands for 15 min and then treated with TNF (20 ng/ml) for 20 min. IKK
complex was immunoprecipitated from 200 µg of lysates, and kinase
activity was determined using GST-IB as a substrate
(top). Relative amounts of recovered IKK protein were
determined by immunoblotting (IB) (bottom).
B, PPC-1 cells were treated with the indicated ligands in
the presence or absence of TNF (20 ng/ml) or TRAIL (100 ng/ml) for
2 h, and assayed for NF-B DNA binding activity by EMSA. The
arrows show NF-B complexes (top and
middle). Asterisks indicate nonspecific
complexes. In parallel, cells were treated for 6 h with PPAR
ligands plus TRAIL, and PARP cleavage was assayed by immunoblotting.
Arrows indicate the full-length and cleaved forms of PARP
protein (bottom).
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Similar conclusions were reached by assessing NF-B induction using
EMSAs, where binding of NF-B to 32P-labeled
oligonucleotide probes containing NF-B-binding sites was measured
(Fig. 3B). For example, when TNF was used as a stimulus for
inducing NF-B DNA binding activity, ciglitazone and CDDO-Me had
little or no effect. In contrast, 15d-PGJ2 completely
inhibited NF-B induction and CDDO reduced levels of NF-B DNA
binding activity by about half, consistent with the kinase data (Fig.
3A). Thus, some PPAR modulators that sensitize tumor
cells to TRAIL reduce IKK activity and inhibit NF-B induction
(15d-PGJ2; CDDO), but others do not (ciglitazone;
CDDO-Me).
To probe further the relationship of NF-B to effects of
PPAR modulators on TRAIL sensitivity, levels of NF-B were also measured in PPC-1 cells following stimulation with TRAIL (instead of
TNF) and correlated with induction of apoptosis, using caspase-mediated cleavage of PARP as a surrogate marker for apoptosis (Fig.
3B). Similar to when TNF was employed, 15d-PGJ2
caused striking reductions in NF-B levels in TRAIL-stimulated cells,
and this correlated with sensitization to TRAIL-induced apoptosis, as
evidenced by PARP cleavage. In contrast, levels of NF-B in
TRAIL-stimulated cells were not reduced by ciglitazone, CDDO, and
CDDO-Me (and may have even been slightly increased), yet all of these
PPAR modulators sensitized PPC-1 cells to TRAIL-induced apoptosis.
We also tested additional prostaglandins, some of which inhibit NF-B
and others which do not, correlating their effects on NF-B with
TRAIL-induced PARP cleavage (Fig. 3B). Among the seven prostaglandins tested (PGA1, PGD2,
PGE2, PGF2
, PGJ2,
12d-PGJ2, and 15d-PGJ2), three of them
(PGA1, PGJ2, and 15d-PGJ2)
completely and one (12d-PGJ2) partially reduced levels of
NF-B to base line. Specifically, PGA1 completely
inhibited NF-B induction by either TRAIL or TNF, probably through
direct inhibition of IKK activity as reported previously (29), but it
failed to sensitize tumor cells to TRAIL-induced apoptosis (Fig.
3B). Thus, reductions in NF-B do not correlate with
sensitization of tumor cells to TRAIL.
TRAIL-sensitizing PPAR Modulators Reduce Levels of FLIP
Protein--
In searching for a mechanism that might explain why
certain PPAR modulators and prostaglandins sensitize tumor cells to
TRAIL, we examined by immunoblotting the levels of several proteins
that are known to be relevant to mechanisms of TRAIL or TNF signaling, including the following: (a) the TRAIL death receptors, DR4
and DR5; (b) the TRAIL decoy receptors, DcR1 and DcR2;
(c) the adapter proteins FADD and TRADD; (d) the
NF-B-inducing TNFR complex component, receptor-interacting
protein; (e) the TNF/Fas-modulator DAP3; and (d)
FLIP, a cellular protein that binds pro-caspase 8 and FADD and
suppresses apoptosis induction by TNF family death receptors (9). Among
these, only the levels of FLIP were affected by PPAR modulators
(Fig. 4). All PPAR modulators and
prostaglandins that had been demonstrated to sensitize tumor cells to
TRAIL-induced apoptosis caused striking reductions in the levels of
c-FLIP protein, including PGJ2, 12d-PGJ2,
15d-PGJ2, ciglitazone, troglitazone, CDDO, and CDDO-Me. In
contrast, FLIP protein levels were not reduced in cells stimulated with
prostaglandins that failed to sensitize tumor cells to TRAIL, including
PGA1, PGD2, PGE2, and
PGF2. Arachidonic acid, a PPAR agonist, also did not
cause reductions in FLIP (Fig. 4) and did not sensitize tumor cells to
TRAIL (Fig. 3B). This selective down-regulation of FLIP
levels by PPAR modulators was observed in every tumor line where
TRAIL sensitization was observed (data for PPC-1 and OVCAR-3 are
presented in Fig. 4). Furthermore, close correlation was observed
between the concentrations of PPAR modulators required to
down-regulate FLIP levels compared with the concentrations needed to
induce apoptosis (Fig. 4, D and E).
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Fig. 4.
Down-regulation of c-FLIP protein by
PPAR ligands. A, PPC-1 cells were treated with 50 µM arachidonic acid, PGA1,
PGD2, PGE2, and PGF2,
PGJ2 or 10 µM 12d-PGJ2 and
15d-PGJ2 for 6 h. Cell lysates were subjected to
immunoblot analysis using c-FLIP, RIP, and -tubulin antibodies.
Control, treatment with vehicle; , lysates from time 0. B, human ovarian cancer OVCAR-3 cells were treated with
15d-PGJ2 (10 µM) or CDDO (5 µM)
for 6 h. Immunoblot analysis was performed using antibodies for
c-FLIP, DR4, DR5, and -tubulin. C, control and untreated
cells. C, PPC-1 cells were treated with 15d-PGJ2
(10 µM), ciglitazone (20 µM), CDDO (5 µM), CDDO-Me (0.5 µM), or troglitazone (10 µM) for 6 h. Cell lysates were normalized for total
protein content and analyzed by immunoblotting using antibodies
specific for FLIP, RIP, TRADD, caspase 8, FADD, DAP3, DcR1, and DcR2.
D, OVCAR-3 cells were cultured for 24 h with various
concentrations of CDDO with or without (control) TRAIL
(top). The percentage of apoptotic cells was determined by
DAPI staining (mean ± S.D.; n = 3). Lysates were
also prepared from cells treated with CDDO alone for 6 h,
normalizing for total protein content (35 µg), and analyzed by
SDS-PAGE/immunoblotting using antibodies specific for FLIP or
-tubulin (bottom). E, OVCAR-3 cells were
cultured and analyzed as in D except that troglitazone was
added to cultures instead of CDDO.
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|
All tumor cell lines tested expressed the longer isoform of FLIP
(FLIPL), whereas the shorter FLIPS protein was
only found in occasional lines. Regardless, PPAR modulators reduced
the levels of both FLIPL and FLIPS (not shown).
PPAR Modulators Enhance Assembly of TRAIL Receptor Signaling
Complexes--
The FLIP protein inhibits recruitment and activation of
pro-caspase 8 at ligand-activated TNF-family death receptor complexes (33, 34). Thus, if PPAR modulators sensitize cells to TRAIL by
reducing FLIP levels, then we would expect to observe enhanced recruitment of pro-caspase 8 to TRAIL receptors and increased caspase 8 activation. To explore this hypothesis, OVCAR-3 cells were
cultured with or without TRAIL and PPAR modulator CDDO individually and in combination, and then the TRAIL receptor DR5 was
immunoprecipitated and associated caspase 8, FLIP, and FADD (a
pro-caspase 8-binding adapter protein) were examined by
SDS-PAGE/immunoblotting (Fig. 5). Control
experiments confirmed successful immunoprecipitation of DR5 by the
anti-DR5 but not by control antibodies (not shown).
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Fig. 5.
CDDO enhances TRAIL-induced caspase 8 activation. OVCAR-3 cells were cultured with 100 ng/ml TRAIL, 5 µM CDDO, both, or neither of these agents, and then cells
were lysed 2 h later, and DR5 was immunoprecipitated. Immune
complexes and lysates were analyzed by SDS-PAGE using antibodies
specific for caspase 8 (top), FLIP (middle), and
FADD (bottom). The positions of unprocessed and fully
processed caspase 8 are indicated. The p43 FLIP band arises from
caspase 8-mediated cleavage (34, 35). The asterisk indicates
contaminating immunoglobulin heavy chain which co-migrates with
pro-caspase 8 and uncleaved p55 FLIP (33, 34) in
co-immunoprecipitations (1st 4 lanes), thus obscuring these
proteins from view.
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When treated with TRAIL or CDDO individually, little pro-caspase 8 was
associated with anti-DR5 immune complexes. However, DR5 immune
complexes recovered from TRAIL-treated cells did contain cleaved
p43-FLIP protein, as expected from prior studies of
FLIPL-expressing cells (33, 34), showing that
FLIPL can become cleaved by caspase 8 when recruited to the
CD95 death-inducing signaling complex (34, 35). The full-length,
uncleaved p55 FLIP protein was not visible in these experiments because
of its co-migration with the immunoglobulin heavy chain band (see Fig.
5 for details). In contrast, treating OVCAR-3 cells with the
combination of CDDO and TRAIL resulted in disappearance of FLIP and
increased association of pro-caspase 8 with DR5 complexes. Moreover, a
partially processed 41-43-kDa form of caspase 8 was found at the
receptor complex (Fig. 5). Analysis of lysates from the same cells by
immunoblotting demonstrated detectable levels of fully processed
18-kDa caspase 8 (catalytic large subunit) and partially processed
41-43-kDa caspase 8 only in cells treated with the combination of CDDO
and TRAIL but not in cells treated with either agent alone. Levels of
FADD were unchanged in cell lysates, thus providing an internal control
for equal loading. Interestingly, recruitment of FADD to DR5 also was
apparently enhanced by CDDO treatment, based on these
co-immunoprecipitation experiments (Fig. 5). Thus, CDDO enhances proper
assembly of the TRAIL-mediated death-inducing signaling complex and
activation of caspase 8.
Genetic Manipulation of FLIP Levels Correlates with Sensitivity to
TRAIL--
To explore the functional significance of the observed
reductions of FLIP protein in tumor cell lines treated with PPAR
modulators, we performed gene transfer experiments, asking what effect
overexpression of FLIP has on the ability of PPAR modulators to
sensitize cells to TRAIL. Transient transfection of PPC-1 cells with an
expression plasmid encoding FLIP abrogated the ability of
15d-PGJ2 (Fig. 6A)
and CDDO (Fig. 6C) to sensitize tumor cells to TRAIL, as
measured by reductions in apoptosis of the transfected cells.
Immunoblotting confirmed that FLIP protein levels were restored in the
transfected cells which received the FLIP expression plasmid (not
shown). These data provide correlative evidence that the sensitization of tumor cells to TRAIL by PPAR modulators could be due to decreases in FLIP levels. Overexpression of FLIP also inhibited TNF-mediated apoptosis (Fig. 6A), consistent with previous reports
(36-38).
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Fig. 6.
Genetic modulation of FLIP expression
correlates with sensitivity to TRAIL-induced apoptosis. PPC-1
cells were transiently transfected with pEGFP (1 µg) and 4 µg of
plasmids encoding c-FLIP (A) or IKK (B). After
20 h, cells were treated with TRAIL (100 ng/ml) or TNF (20 ng/ml)
for 6 h in the presence of 15d-PGJ2 (10 µM). C, cells were transiently transfected
with 1 µg of pEGFP and 4 µg of plasmids encoding c-FLIP, IKK,
CrmA, and Bcl-2 proteins. The next day, cells were treated with CDDO (5 µM) in the presence of TRAIL (100 ng/ml) for 6 h.
The percentage of apoptotic GFP-positive cells was determined by DAPI
staining (mean ± S.D.; n = 3). D,
cells were transfected with FLIP or control antisense oligonucleotides
for 14 h and then treated with TRAIL (25 ng/ml) for 7 h.
Apoptotic cells were counted by DAPI staining. Insets show
the levels of long and short forms of FLIP proteins in cells
transfected with control (C) and FLIP antisense
(AS) oligonucleotides, respectively. Levels of -tubulin
are shown as a control.
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A gene transfer approach was also used to explore other aspects of the
apoptotic mechanism induced by the combination of TRAIL and PPAR
modulators. To address further the issue of NF-B, PPC-1 cells were
transiently transfected with a plasmid producing IKK, which caused
marked increases in NF-B activity (not shown). When apoptosis was
induced by the combination of TRAIL and 15d-PGJ2, IKK
overexpression failed to provide protection (Fig. 6B). In contrast, when apoptosis was induced by the combination of TNF and
15d-PGJ2, then IKK overexpression potently suppressed
apoptosis. These data are consistent with prior reports demonstrating
an important role for NF-B in regulating apoptosis induction by TNF
(31) but indicate that NF-B is not protective against TRAIL-induced apoptosis (39). Consistent with the documented role for caspases in
TRAIL induced apoptosis, overexpression of the CrmA (a viral inhibitor
of caspase 8 (40, 41) suppressed apoptosis induced by the
combination of TRAIL and PPAR modulators such as CDDO (Fig.
6C). In contrast, overexpression of Bcl-2 did not suppress apoptosis (Fig. 6C), consistent with evidence that TRAIL and
other death receptors can trigger apoptosis through Bcl-2-independent mechanisms in many types of cells.
Finally, antisense oligonucleotides targeting FLIP were used to explore
whether down-regulation of FLIP protein levels is sufficient to
sensitize tumor cells to TRAIL. Treatment of PPC-1 cells with FLIP
antisense oligonucleotides resulted in a pronounced reduction in the
levels of both the long and short isoforms of FLIP protein, compared
with control oligonucleotide-treated cells (Fig. 6D). This
antisense-mediated reduction in FLIP protein levels was correlated with
enhanced sensitivity of PPC-1 cells to TRAIL-induced apoptosis (Fig.
6D). Therefore, we conclude that reducing FLIP expression is
indeed sufficient to sensitize at least some tumor cell lines to
TRAIL-induced apoptosis.
PPAR Levels and Activity Do Not Correlate with TRAIL
Responses--
Although the agents used here are known to modulate
PPAR, we suspected a PPAR-independent mechanism accounted for
their ability to sensitize tumor cells to TRAIL, given that PPAR
agonists (15d-PGJ2, ciglitazone, and troglitazone), weak
agonists (CDDO), and antagonists (CDDO-Me) were effective. We therefore
explored the effects of overexpressing PPAR on TRAIL-induced
apoptosis, reasoning that if PPAR modulators were functioning
through a PPAR-dependent process then they should be
more potent when PPAR levels are higher. HeLa cells were employed
for these studies because of their relatively low endogenous levels of
PPAR protein (32). Overexpressing PPAR had no effect on apoptosis
induction by the combination of TRAIL and CDDO, despite successful
transfection of >80% of cells as determined by a marker plasmid
encoding green fluorescent protein (GFP). To confirm that these
transfections resulted in higher levels of functional, bioactive
PPAR protein, reporter gene assays were performed in parallel,
confirming that transfection of HeLa cells with PPAR-encoding
plasmids resulted in dose-dependent increases in
ligand-activated expression of a luciferase reporter gene plasmid
containing PPAR-response elements (PPREs) (Fig.
7B). We conclude therefore
that although 15d-PGJ2 and CDDO are capable of activating
PPAR, their ability to sensitize tumor cells to TRAIL does not
correlate with effects on PPAR.
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Fig. 7.
Synergistic apoptosis induced by TRAIL and
PPAR ligands is independent of PPAR levels. A, HeLa
cells were transiently transfected with pCMX-mPPAR and pEGFP
plasmids and then treated 1 day later with 15d-PGJ2 (10 µM) or CDDO (5 µM) in the presence of TRAIL
(50 ng/ml) for 8 h. Apoptosis was determined by DAPI staining
among GFP-positive cells (mean ± S.D.; n = 3).
Inset shows the levels of PPAR protein detected by
immunoblotting after normalization of cell lysates for total protein
content. B, HeLa cells were transiently transfected with
various amounts of pCMX-mPPAR together with a fixed amount of
Tk-PPRE3-luciferase vector. After 16 h, transfected cells were
treated with 15d-PGJ2 (10 µM) or CDDO (5 µM) for 8 h, and then luciferase activity was
determined. Data are presented as fold induction relative to luciferase
activity in cells transfected with the empty vector and cultured
without ligand (mean ± S.D.; n = 3).
C, PPC-1 cells were transiently transfected with plasmids
encoding dominant-negative PPAR (4 µg) and pEGFP (1 µg) in 60-mm
dishes. After 16 h of incubation, cells were treated with TRAIL
(100 ng/ml) plus CDDO (5 µM) for 6 h. Apoptotic
cells were determined by DAPI staining (mean ± S.D.;
n = 3). D, PPC-1 cells were transfected with
Tk-PPRE3-luciferase vector along with dominant-negative PPAR
plasmids for 16 h and then treated with 15d-PGJ2 (5 µM) for 24 h, and luciferase activity was measured.
Data are shown as fold induction relative to luciferase activity in
cells transfected with the empty vector and cultured without the ligand
(mean ± S.D.; n = 3).
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As an alternative approach, we employed a mutant of PPAR that
functions as a dominant-negative inhibitor of the endogenous protein
(22). Overexpression of dominant-negative PPAR failed to abrogate
CDDO-mediated sensitization of PPC-1 cells to TRAIL (Fig.
7C), despite its ability to block completely
15d-PGJ2-mediated induction of PPAR transcriptional
activity (as determined by reporter gene assays using a
PPAR-inducible luciferase reporter gene) (Fig. 7D). We
conclude therefore that the TRAIL-sensitizing effects of CDDO can be
dissociated from its effects on PPAR.
PPAR Modulators Reduce FLIP Protein Levels through a
Non-transcriptional Mechanism Involving Protein
Ubiquitination--
RNase protection assays were performed to measure
relative levels of c-FLIP mRNA in PPC-1 and
OVCAR-3 cells treated with PPAR modulators. Although causing a
profound decrease in FLIP protein levels (Fig. 4), corresponding
decreases were not observed in c-FLIP mRNA levels or in
levels of several other mRNAs that serve as comparison controls
(Fig. 8A and not shown). Thus,
PPAR modulators appear to cause FLIP protein reductions through a
transcription-independent process, lending further support for a
PPAR-independent mechanism.
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Fig. 8.
Down-regulation of FLIP protein by PPAR
modulators is mediated through a ubiquitin-proteasome pathway.
A, PPC-1 cells were treated with indicated PPAR ligands
(10 µM 15d-PGJ2, 20 µM
ciglitazone, 5 µM CDDO, 0.5 µM CDDO-Me, and
10 µM troglitazone) for 1 h (lanes
2, 4, 6, 8, and 10) or
6 h (lanes 3, 5, 7, 9,
and 11). Total RNAs were isolated, and 10 µg of each RNA
was analyzed using a multiplex RNase protection assay which includes
probes for c-FLIP and several other genes. The bands
for L32 and glyceraldehyde-3-phosphate dehydrogenase (GAPDH)
(loading control) were exposed for much a shorter time due to high
intensity. B, OVCAR-3 cells were cultured with 5 mM CDDO for various times before lysing cells and analyzing
levels of FLIP protein by immunoblotting. Lysates were normalized for
total protein content (35 µg), and equivalent loading was confirmed
by reprobing the same blot with anti--tubulin. C, PPC-1
cells were pretreated for 30 min with Z-VAD-fmk (50 µM),
MG-132 (25 µM), lactacystin (20 µM), MG115
(50 µM), TLCK (50 µM), or calpeptin (20 µM) and then treated with CDDO (5 µM) for
6 h. Cell lysates were analyzed by immunoblotting using anti-FLIP
antibodies. The blot was reprobed with an antibody specific for
-tubulin (loading control). D, SK-OV-3 cells were treated
with 5 µM CDDO for the indicated times in the absence or
presence of 400 nM epoxomicin. Lysates (60 µg) were
subjected to immunoblotting analysis by FLIP antibody and exposed to
x-ray film for either 30 (top) or 1 min (middle).
In parallel, lysates were immunoprecipitated with FLIP antibody and
analyzed by SDS-PAGE/immunoblotting using anti-ubiquitin antibody
(bottom). E, SK-OV-3 cells were transfected with
plasmids encoding FLIP and/or His6-ubiquitin. After 1 day,
cells were treated with CDDO (5 µM) and/or epoxomicin
(400 nM). His6-tagged ubiquitinated proteins
were purified with cobalt chelate resin, and multiubiquitinated FLIP
products were detected by anti-FLIP antibody (top). FLIP
expression was analyzed by direct anti-FLIP immunoblotting of cell
lysates (bottom).
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Analysis of the kinetics of FLIP protein reductions in cells treated
with CDDO (Fig. 8B) and other PPAR modulators (not shown) revealed a rapid process, with decreases evident within 20 min and
complete clearance of FLIP protein from cells by 2 h. Pulse-chase experiments using L-[35S]methionine-labeled
cells confirmed that CDDO induced increases in the rate of FLIP protein
degradation (not shown).
To explore the potential role of proteases in the reduction of FLIP
protein induced by PPAR modulators, tumor cells were cultured with
inhibitors of caspases (Z-VAD-fmk), the 26 S proteosome (MG132,
lactacystin, and MG115), and lysosomal proteases (TLCK and calpeptin).
MG132, lactacystin, and MG115 partially prevented FLIP down-regulation
by CDDO, suggesting a potential role for the proteosome in this
mechanism. In contrast, inhibitors of caspases and lysosomal proteases
had no effect (Fig. 8C).
Because the 26 S proteasome degrades ubiquitinated proteins, we
investigated whether PPAR modulators induce ubiquitination of the
FLIP protein. Long exposures of immunoblots probed with anti-FLIP
antibody demonstrated the presence of higher molecular weight
conjugates of FLIP after treatment with CDDO (Fig. 8D, top). The appearance of these higher molecular weight forms
of FLIP was evident within 1 h after CDDO treatment, coinciding
with the approximate time when FLIPL and FLIPS
protein levels began to decline. This higher molecular weight material
was confirmed to represent poly-ubiquitinated FLIP, based on
experiments where lysates from CDDO-treated cells were
immunoprecipitated using anti-FLIP antibody, and the resulting immune
complexes were analyzed by SDS-PAGE/immunoblotting using anti-ubiquitin
antibodies (Fig. 8D, bottom). Treating cells with
the proteasome inhibitors, such as epoxomicin, also induced a slight
increase in ubiquitinated FLIP, indicative of a basal low rate of FLIP
ubiquitation, but CDDO massively increased the abundance of ubiquitin
conjugates of FLIP (Fig. 8D). In contrast to FLIP, the
extent of ubiquitination and steady-state levels of p53 and -catenin
(proteins known to be regulated by ubiquitin-proteasome pathways (42,
43)) were not altered by CDDO treatment of cells (not shown).
To provide further evidence of CDDO-inducible ubiquitination of FLIP,
cells were transiently co-transfected with plasmids encoding FLIP and
histidine-tagged (His6) ubiquitin. After culturing cells
for 4 h with CDDO, epoxomicin, or the combination of these reagents, lysates were subjected to cobalt-chelation chromatography to
recover His6-tagged proteins, followed by
SDS-PAGE/immunoblotting using anti-FLIP antibody (Fig. 8E).
As shown, His6-ubiquitin-conjugated FLIP products
accumulated in cells treated with either CDDO or epoxomicin but not in
untreated cells (Fig. 8E). Furthermore, the combination of
CDDO and epoximicin led to even higher increases in
His6-ubiquitin-conjugated FLIP products, consistent with
the hypothesis that CDDO induces increases in FLIP ubiquitination, whereas epoxomicin prevents degradation of the ubiquitinated FLIP proteins. Taken together, these results suggest that CDDO enhances ubiquitination of FLIP, thus accelerating the degradation of FLIP protein by the proteasome.
| |
DISCUSSION |
In this report, we reveal the existence of an inducible pathway
that triggers ubiquitination and degradation of the FLIP protein and is
capable of sensitizing at least some types of the transformed cells to
death receptor-mediated apoptosis in vitro. Interestingly, T-cell receptor stimulation of T-lymphocytes has also been reported to
diminish levels of FLIP protein without concomitant changes in mRNA
levels (44). In addition, reductions in FLIP protein levels induced by
p53 are reported negated by proteasome inhibitors (45). Thus, precedent
exists for post-transcriptional regulation of FLIP expression. For many
proteins whose levels are conditionally regulated by ubiquitination and
proteasome-dependent degradation, ubiquitination is induced
upon binding of E3 ubiquitin ligases to the target protein (reviewed in
Ref. 46). In this regard, FLIP has been shown to interact with TRAF2,
which contains a RING finger domain known to possess E3 ligase activity
(33). It is conceivable therefore that PPAR modulatory drugs
influence the expression of TRAF2 or other types of FLIP-binding E3
ubiquitin ligases. Several other mechanisms could also be envisioned.
A recent report (47) demonstrated that troglitazone can sensitize two
human tumor cell lines to TRAIL; however, the responsible mechanism was
not addressed here. We extended this observation to multiple solid
tumor cell lines and to several classes of PPAR modulators. Our data
indicate that compounds previously recognized for their ability to bind
and modulate the function of PPAR have an additional
PPAR-independent mechanism, allowing them to reduce FLIP protein
levels and sensitize cancer cells to apoptosis induction by TRAIL. The
evidence arguing against a PPAR-dependent mechanism for
these compounds includes the following: (a) efficacy of both PPAR agonists (15d-PGJ2, ciglitazone, troglitazone, and
CDDO) and antagonists (CDDO-Me) (28); (b) failure of PPAR
overexpression and dominant-negative PPAR to alter effects of
compounds on TRAIL-induced apoptosis; and (c) decreases in
FLIP protein levels without concomitant reductions in mRNA,
suggesting a non-transcriptional mechanism uncharacteristic of PPAR.
Prior studies of effects of thiazolidinediones and 15d-PGJ2
on cells from PPAR knock-out mice have demonstrated PPAR-independent inhibition of cytokine production by activated macrophages (48), suggesting an alternative target of these agents.
Moreover, it has been reported recently that the growth-suppressive effects of thiazolidinediones on cells in vitro and
in vivo are independent of this receptor, based on
experiments using PPAR(/) cells (49). In this regard, the IB
kinases that control NF-B activity have been identified as direct
targets of 15d-PGJ2 and some other types of PPAR
modulators. However, IKK and NF-B also are not the relevant targets
of the TRAIL-sensitizing compounds studied here, because overexpression
of IKK failed to restore TRAIL resistance and because some
TRAIL-sensitizing compounds (CDDO-Me; ciglitazone; troglitazone) did
not inhibit IKK or reduce NF-B DNA binding activity.
We speculate that natural PPAR agonists (PGJ2,
12d-PGJ2, and 15d-PGJ2) and synthetic PPAR
modulators, including thiazolidinediones (troglitazone and ciglitazone)
and triterpenoids (CDDO and CDDO-Me), interact with an unidentified
cellular target, resulting in post-transcriptional reductions in FLIP
protein levels, inducing FLIP protein degradation through a
ubiquitin-proteasome pathway. If PPAR modulators interact with a
novel target protein, then medicinal chemistry efforts potentially
could be used for identifying analogues of these compounds that retain
the ability to reduce FLIP levels without affecting PPAR or IKK
activity. Specific analogues of this type might prove useful for
identifying the relevant molecular target that controls FLIP
ubiquitination. Given that triterpenoids such as CDDO are roughly
1 log more potent than the thiazolidinediones examined here at inducing
FLIP degradation and sensitizing tumor cells to TRAIL, this class of
chemical compounds should be given particular attention in the efforts
to design selective agonists of the FLIP degradation pathway that lack
effects on PPAR. Interestingly, when used at higher concentrations
(~5 µM), CDDO has recently been reported to trigger
apoptosis of established leukemia and osteosarcoma cell lines through a
pathway involving activation of caspase 8 but not caspase 9 and through
a mechanisms that is suppressible by CrmA but not by Bcl-XL
(50, 51). These observations are consistent with activation of the
"extrinsic" apoptosis pathway, which is commonly invoked by TNF
family death receptors, as opposed to the "intrinsic" apoptosis
pathway where mitochondria play a critical role (reviewed in Ref. 52).
Thus, CDDO and related triterpenoids may possess the ability to
activate the extrinsic pathway as single agents, without accompanying
application of TRAIL or other death ligands, provided sufficiently high
concentrations are employed. In such cases, it remains to be determined
whether these compounds induce autocrine expression of TNF family death ligands or receptors, thus accounting for these observations. Regardless, such findings hint that CDDO and related molecules possess
additional activities, besides induction of FLIP degradation, which
promote activation of the extrinsic pathway. Moreover, because FLIP has
been reported to regulate NF-B and extracellular signal-regulated kinase signal transduction pathways (33) (in addition to suppressing caspase 8 activation), it is also possible that simply ablating FLIP
expression can affect gene expression, thereby modulating apoptosis
pathways in a cell context-dependent manner.
It has been reported that some preparations of recombinant TRAIL induce
apoptosis of normal human hepatocytes, thus raising concerns about
potential hepatotoxicity (reviewed in Ref. 53). Subsequent studies,
however, demonstrated that toxicity to hepatocytes is associated with
aggregated TRAIL and is not seen with soluble trimeric TRAIL (5).
Although we were unable to obtain suitable preparations of human
hepatocytes to assess the effects of combined treatment with TRAIL and
PPAR modulators, we did perform testing using (as an alternative)
hepatocytes from cynomologus monkeys, but we failed to see induction of
apoptosis. In this regard, the TRAIL receptors of cynomologus monkey
are 84-99% identical to their human counterparts (5). Moreover, it
has been demonstrated that hepatocytes from these primates bind human
TRAIL with high affinity (5). More importantly, these monkey and human
hepatocytes are equivalent in their apoptotic responses to "good"
(trimeric) and "bad" (aggregated) preparations of TRAIL, where
trimeric TRAIL fails to induce apoptosis of both human and monkey
cells, whereas aggregated TRAIL kills hepatocytes from both species
(5). Thus, hepatocytes from cynomologus monkeys are a valid surrogate
for human hepatocytes where TRAIL-induced apoptosis is concerned. The
differential effects on normal versus malignant cells of
combination treatment with TRAIL plus PPAR modulators in
vitro suggest that it could be possible to exploit these agents
for the treatment of cancer, particularly because thiazolidinedione
class PPAR modulators are already in clinical use as insulin
sensitizers for treatment of type II diabetes. Thus, although in
vivo data are presently lacking, we speculate that perhaps these
or other PPAR modulators might be used for an alternative purpose
such as sensitizers of cancer cells to TRAIL and other activators of TNF family death receptors.
| |
ACKNOWLEDGEMENTS |
We thank M. Peter, R. Evans, D. Hwang, Z-L.
Chu, and R. R. Kopito for antibodies and plasmids; S. Takayama and
S. Matsuzawa for helpful discussion; F. Bennett and W. Rickets of ISIS
Pharmaceuticals for antisense oligonucleotides; and R. Cornell for
manuscript preparation. We also thank Frank Stenner-Liewen and Ivo
Meinhold-Heerlein for providing unpublished data regarding the TRAIL
responsiveness and TRAIL receptor status of ovarian cancer cell lines.
| |
FOOTNOTES |
*
This work was supported by National Institutes of Health
Grants CA 69381, CA55164, and CA78040 and by Susan G. Komen Foundation California Breast Cancer Research Program Grant 5FB-0170.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.
§
To whom correspondence should be addressed: The Burnham Institute,
10901 N. Torrey Pines Rd., La Jolla, CA 92037. Tel.: 858-646-3100; Fax:
858-646-3194; E-mail: jreed@burnham.org.
Published, JBC Papers in Press, April 8, 2002, DOI 10.1074/jbc.M202458200
| |
ABBREVIATIONS |
The abbreviations used are:
TRAIL, TNF-related
apoptosis inducing ligand;
TNF, tumor necrosis factor;
CDDO, 2-cyano-3,12-dioxooleane-1,9-dien-28-oic acid;
15d-PGJ2, 15-deoxy-
12,14-prostaglandin J2;
FLIP, FLICE-inhibitory protein;
IKK, IB kinase;
PPAR, peroxisome
proliferator-activated receptor;
TLCK, 1-chloro-3-tosylamido-7-amino-2-heptanone;
MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide;
Z, benzyloxycarbonyl;
fmk, fluoromethyl ketone;
GST, glutathione
S-transferase;
DAPI, 4,6-diamidino-2-phenylindole;
EMSA, electrophoretic mobility shift assays;
GFP, green fluorescent protein;
PARP, poly(ADP-ribose) polymerase;
PPREs, PPAR-response
elements;
E3, ubiquitin-protein isopeptide ligase.
| |
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TOP
ABSTRACT
INTRODUCTION
