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TRAIL is an endogenous death receptor ligand also used therapeutically because of its selective proapoptotic activity in cancer cells. In the present study, we examined chromatin alterations induced by TRAIL and show that TRAIL induces a rapid activation of DNA damage response (DDR) pathways with histone H2AX, Chk2, ATM, and DNA-PK phosphorylations. Within 1 h of TRAIL exposure, immunofluorescence confocal microscopy revealed γ-H2AX peripheral nuclear staining (γ-H2AX ring) colocalizing with phosphorylated/activated Chk2, ATM, and DNA-PK inside heterochromatin regions. The marginal distribution of DDR proteins in early apoptotic cells is remarkably different from the focal staining seen after DNA damage. TRAIL-induced DDR was suppressed upon caspase inhibition or Bax inactivation, demonstrating that the DDR activated by TRAIL is downstream from the mitochondrial death pathway. H2AX phosphorylation was dependent on DNA-PK, while Chk2 phosphorylation was dependent on both ATM and DNA-PK. Downregulation of Chk2 decreased TRAIL-induced cell detachment; delayed the activation of caspases 2, 3, 8, and 9; and reduced TRAIL-induced cell killing. Together, our findings suggest that nuclear activation of Chk2 by TRAIL acts as a positive feedback loop involving the mitochondrion-dependent activation of caspases, independently of p53.
Programmed cell death, apoptosis, is a normal physiological process in which damaged or harmful cells are eliminated. It is essential for tissue homeostasis, providing a balance between survival and cellular destruction. Deregulation of apoptosis is a common characteristic of many diseases, including cancers and autoimmune disorders (66). Apoptosis consists of intrinsic and extrinsic pathways. The intrinsic pathway can be initiated by DNA or tubulin damage and engages the mitochondria. The extrinsic pathway is initiated by members of the TNF (tumor necrosis factor) superfamily and engages the death-inducing signaling complex (DISC) at the cell surface (14).
TNF-related apoptosis-inducing ligand (TRAIL), a member of the TNF/death receptor (DR) gene superfamily, is a physiological endogenous ligand and a promising cancer therapy because it induces apoptosis preferentially in cancer cells (29, 37). TRAIL can bind to four plasma membrane receptors and one soluble receptor, i.e., TRAIL-R1 (DR4), TRAIL-R2 (DR5/KILLER), TRAIL-R3 (DcR1), TRAIL-R4 (DcR2), and osteoprotegerin, respectively (26). DR4 and DR5 contain a common conserved death domain motif. After binding TRAIL, DR4 and DR5 form trimeric complexes and generate the DISC. In the DISC, the adaptor protein FADD (Fas-associated death domain) binds to the DR and to procaspase 8, promoting the autoactivation of this caspase. DISC-activated caspase 8 engages the downstream apoptotic death machinery in two ways. In type I cells, DISC assembly activates sufficient amounts of caspase 8 to cleave and activate downstream effector caspases. In type II cells, DISC assembly activates smaller amounts of caspase 8 and requires amplification of the apoptotic signal through the mitochondrial apoptotic pathway. Activation of this mitochondrial amplification loop is achieved through cleavage of Bid, a proapoptotic member of the Bcl-2 family (33). Cleaved Bid binds to and activates Bax and Bak, which causes the release of apoptogenic factors such as cytochrome c from mitochondria. In turn, cytochrome c activates effector caspases via Apaf-1 and caspase 9 in the apoptosome (33).
Preclinical evaluations demonstrated that recombinant TRAIL inhibits tumor growth and induces the regression of a broad range of leukemia and solid malignancies (26). Clinical trials with TRAIL (www.gene.com/gene/pipeline/status/oncology/apo2l/) and DR antibodies are ongoing (53). HGS-ETR1 (a human agonistic monoclonal antibody that targets DR4) is in clinical trial in combination with bortezomib against advanced multiple myeloma and in combination with carboplatin/paclitaxel and cisplatin/gemcitabine against various advanced solid malignancies. HGS-ETR2 (a human agonistic monoclonal antibody that targets DR5) is in phase I trial with patients with advanced solid malignancies. In contrast to monoclonal antibodies to TRAIL receptors, TRAIL interacts with both DR4 and DR5, as well as with the decoy receptors DcR1 and DcR2. Thus, recombinant TRAIL may have a wider therapeutic spectrum than the highly specific antibodies.
One of the biochemical landmarks of apoptosis is the formation of DNA double-strand breaks (DSB; producing oligonucleosomal DNA fragments and in some cells only large fragments [50 to 300 kbp]) (55). Under conditions unrelated to apoptosis, DSB induce the rapid activation of conserved DNA damage response (DDR) pathways (56). Chk2 (checkpoint kinase 2) becomes phosphorylated at threonine 68 (T68) by several members of the phosphatidylinositol-3-kinase family, i.e., ATM (ataxia telangiectasia mutated), ATR (ATM and Rad3 homolog), and DNA-PK (DNA-dependent protein kinase) (2, 31, 43). In addition to its cell cycle and checkpoint/DNA repair functions, Chk2 has a proapoptotic function, which is mediated in part by p53 (6, 20, 23, 25, 35, 58).
One prominent chromatin modification in response to DSB is phosphorylation of histone H2AX on serine 139, which is referred to as γ-H2AX (48). Both ATM and DNA-PK catalyze H2AX phosphorylation (5, 15, 38), and endogenous activation of the DDR pathways has been observed in early tumorigenesis (3, 13). γ-H2AX has also been associated with apoptosis (38, 50, 60). The focus of the present study was to determine whether TRAIL induces the activation of DDR pathways and whether this activation could have a functional impact on apoptosis. We also describe and analyze a previously unidentified marginal and confluent (ring) staining of γ-H2AX in early apoptotic cells.
Recombinant human soluble TRAIL was obtained from Alexis Biochemicals (Axxora, San Diego, CA). The broad-spectrum caspase inhibitor Z-VAD-fmk (z-Val-Ala-dl-Asp-fluoromethylketone) was from Bachem (Torrance, CA). The DNA-PK kinase inhibitor (DNA-PKi; NU7441) and ATM kinase inhibitor (ATMi; KU-55933) used were from Kudos Pharmaceuticals (Cambridge, United Kingdom).
The human colon carcinoma HCT116, cervix carcinoma HeLa, and leukemic Jurkat cell lines were obtained from ATCC (Manassas, VA). HCT116 cells complemented with Mre11 (HCT116-Mre11 cells) were generated in our laboratory (59). Bax+/−, Bax−/−, Chk2−/−, p53+/+, and p53−/− HCT116 cells were kind gifts from Bert Vogelstein (Johns Hopkins Oncology Center, Baltimore, MD) (4, 24, 64). The glioma M059J-Fus1 and M059J-Fus9 cell lines were from Janice Pluth (Lawrence Berkeley National Laboratory, Berkeley, CA) (21). PrEC human prostate epithelial cells were obtained from Lonza Walkersville, Inc. (Walkersville, MD).
Cells were washed twice in PBS (phosphate-buffered saline) and lysed at 4°C in buffer containing 1% SDS (sodium dodecyl sulfate) and 10 mM Tris-HCl, pH 7.4, and supplemented with protease inhibitors (Roche Applied Science, Indianapolis, IN) and phosphatase inhibitors (Sigma Chemical Co., St. Louis, MO). The viscosity of the samples was reduced by brief sonication. Equal amounts of proteins were boiled for 5 min in Tris-glycine-SDS sample buffer (Invitrogen, Carlsbad, CA) or heated at 70°C for 10 min in lithium dodecyl sulfate sample buffer (Invitrogen), separated by Tris-glycine or Tris-acetate polyacrylamide gel electrophoresis (Invitrogen), and electroblotted onto nitrocellulose membranes (Bio-Rad, Hercules, CA). The membranes were saturated with milk, incubated overnight at 4°C with primary antibodies, washed, and then incubated for 45 min with secondary antibodies, i.e., peroxidase-conjugated goat anti-mouse or anti-rabbit immunoglobulin G (Santa Cruz Biotechnology, Santa Cruz, CA). Signals were revealed by autoradiography with the enhanced chemiluminescence detection kit (Pierce, Rockford, IL).
The primary antibodies used were anti-P-ATM S1981 (4526; Cell Signaling, Danvers, MA), anti-53BP1 (NB100-305; NOVUS Biologicals, Littleton, CO), anti-P-Cdc25A S123 (AP3045a; ABGENT, San Diego, CA), anti-Cdc25A (sc-7389; Santa Cruz Biotechnology), anti-P-Cdc25C S216 (9528; Cell Signaling), anti-P-Chk1 S345 (2341; Cell Signaling), anti-Chk1 (sc-8408; Santa Cruz Biotechnology), anti-P-Chk2 T68 (2661; Cell Signaling), anti-Chk2 (sc-17747; Santa Cruz Biotechnology), anti-P-DNA-PK S2056 (ab18192; Abcam, Cambridge, MA), anti-P-DNA-PK T2609 (ab18356; Abcam), anti-DNA-PK (NA57; Calbiochem, EMD Biosciences, San Diego, CA), anti-P-E2F1 S364 (ab5391; Abcam), anti-E2F1 (sc-251; Santa Cruz Biotechnology), anti-γ-H2AX (05-636; Upstate, Temecula, CA), anti-H2AX (07-627; Upstate), anti-histone H3 trimethyl K9 (ab8898; Abcam), anti-lamin B1 (ab16048; Abcam), anti-p21 (OP64; Calbiochem), anti-P-p53 S15 (9284; Cell Signaling), anti-P-p53 S20 (9287; Cell Signaling), anti-p53 (sc-126; Santa Cruz Biotechnology), and antitubulin (MS-581; Lab Vision, Fremont, CA).
siRNA targeting DNA-PK was obtained from Dharmacon, Chicago, IL (SMARTpool, catalogue number M-005030-01). siRNA targeting Chk2 was obtained from Qiagen, Valencia, CA (validated siRNA, catalogue number SI02224271). siRNA targeting H2AX and negative control siRNA were obtained from Ambion, Austin, TX (catalogue numbers AM16210 and 4635). Cells were seeded in six-well plates, at a density of 200,000 cells per well, 16 h before transfection. For each sample, 500 pmol of siRNA was mixed with 250 μl of Opti-MEM (mixture A; Invitrogen). Five microliters of Lipofectamine 2000 (Invitrogen) was mixed with 250 μl of Opti-MEM and incubated for 5 min at room temperature (mixture B). After mixing (mixtures A and B) and further incubation for 20 min at room temperature, the siRNA/Lipofectamine complexes were added to 2 ml of culture medium. After 5 h, the medium was replaced with regular medium and the cells were incubated for a further 48 h.
Cells (105) were washed in PBS, collected by centrifugation at 500 × g for 10 min, and resuspended in 500 μl of PBS containing Hoechst 33342 (final concentration, ~2 to 10 μg/ml). After a 30-min incubation in the dark at 37°C, the cells were centrifuged at 500 × g for 5 min, resuspended in 20 μl of PBS, and analyzed by fluorescence microscopy.
Cells were washed with PBS, fixed with 2% formaldehyde in PBS for 20 min, washed with PBS, postfixed and permeabilized with cold (−20°C) 70% ethanol for 20 min, washed with PBS, blocked with 8% bovine serum albumin (BSA) in PBS for 1 h, washed with PBS, incubated with the first antibody (P-Chk2 T68, 1/500 dilution; γ-H2AX, 1/800 dilution; P-ATM S1981, 1/250 dilution; P-DNA-PK T2609, 1/250 dilution; H3K9me3, 1/250 dilution; lamin B1, 1/500 dilution; 53BP1, 1/500 dilution) in 1% BSA in PBS for 2 h, washed with PBS, incubated with the secondary antibody conjugated with Alexa Fluor 488 or 568 for 1 h at room temperature, washed with PBS, and mounted with Vectashield mounting medium with DAPI (4′,6′-diamidino-2-phenylindole) or propidium iodide to counterstain the DNA (Vector Laboratories). Incubation with 0.5 mg/ml RNase A for 15 min was performed before the addition of propidium iodide-containing Vectashield. Confocal images were sequentially acquired with Zeiss AIM software on a Zeiss LSM 510 NLO confocal system (Carl Zeiss Inc., Thornwood, NY). Three-dimensional (3D) pictures were realized with the Bitplane (Zurich, Switzerland) Imaris software v6.0. The line intensity profiles were realized with the Zeiss AIM software v3.2.
Three hundred thousands cells were fixed and permeabilized with cold (−20°C) 70% ethanol overnight, washed with 1% BSA-PBS, and further permeabilized with 0.25% Triton in 1% BSA-PBS for 5 min on ice, incubated with anti-γ-H2AX antibody (dilution, 1/37.5) in 10% normal goat serum-1% BSA-PBS, washed with 1% BSA-PBS, and incubated with a secondary antibody conjugated with Alexa Fluor 488. Finally, propidium iodide (final concentration, 0.05 mg/ml) and RNase A (final concentration, 0.5 mg/ml) were added. Fluorescence intensities were determined with a FACScan flow cytometer (Becton Dickinson) and quantified with CellQuest software (Becton Dickinson).
Following 1 h of treatment with TRAIL (0.1 μg/ml), cells were trypsinized, washed in PBS, and seeded in triplicate at three densities, 100, 1,000, and 10,000 cells per well, in six-well plates. Colonies were allowed to grow for 10 days and visualized following washing with PBS, fixation in methanol for 30 min, washing again with PBS, and staining with 0.05% methylene blue for 30 min. Percent survival was normalized to the observed number of colonies generated from untreated cells.
Cells were washed twice in PBS, lysed in 150 mM NaCl-50 mM Tris-HCl (pH 8.0)-0.1% SDS-1% Nonidet P-40-0.5% sodium deoxycholate for 30 min at 4°C, and centrifuged (10,000 × g at 4°C). Fifteen micrograms of proteins from the resulting supernatant was incubated in 100 mM HEPES (pH 7.0)-1 mM EDTA-0.1% 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS)-10% glycerol-20 mM dithiothreitol in the presence of the fluorogenic peptide substrate Z-VDVAD-AFC (Z-Val-Asp-Val-Ala-Asp-AFC, caspase 2), Ac-DEVD-AFC (Ac-Asp-Glu-Val-Asp-AFC, caspase 3), Z-IETD-AFC (Z-Ile-Glu-Thr-Asp-AFC, caspase 8), or Ac-LEHD-AFC (Ac-Leu-Glu-His-Asp-AFC, caspase 9) (Calbiochem) at 100 μM. The 7-amino-4-trifluoromethylcoumarin (AFC) released from the substrate was excited at 400 nm to measure emission at 505 nm. Fluorescence was monitored continuously at 37°C for 30 min in a dual-luminescence fluorimeter (SpectraMax Gemini XS; Molecular Devices). Caspase activities were determined as initial velocities expressed as relative intensity per minute per milligram of protein.
The assay has been described previously (57). Briefly, cells were incubated with [2-14C]thymidine (0.02 μCi/ml) for 2 days and chased overnight in radioisotope-free medium. After drug treatment, cells were loaded onto a protein-adsorbing filter (Metricel membrane filter, 0.8-μm pore size, 25-mm diameter; PALL Corp., Cortland, NY), washed with PBS, and lysed in 0.2% sodium Sarkosyl-2 M NaCl-0.04 M EDTA, pH 10. The filters were then washed with 0.02 M EDTA, pH 10. DNA was depurinated by incubation of the filters in 1 M HCl at 65°C and then released from the filters with 0.4 M NaOH at room temperature. Radioactivity was counted by liquid scintillation spectrometry in each fraction (wash, lysis, EDTA wash, and filter). DNA fragmentation was measured as the fraction of disintegrations per minute in the wash plus lysis fraction plus EDTA wash relative to the total intracellular disintegrations per minute.
Cells were washed with PBS, fixed, and permeabilized with cold (−20°C) 70% ethanol overnight. The next day, cell pellets were washed again with PBS, resuspended in PBS buffer containing 0.2% NP-40 and 0.5 mg/ml RNase A, incubated at room temperature for 15 min, and put on ice 10 min prior the addition of 50 μg/ml propidium iodide. DNA content was determined with a FACScan flow cytometer (Becton Dickinson) and quantified with CellQuest software (Becton Dickinson).
Figure Figure11 shows that treatment with TRAIL induces rapid phosphorylation of Chk2 (at threonine 68), ATM (at serine 1981), H2AX (at serine 139), and DNA-PK (at threonine 2609), starting at 1 h after TRAIL treatment. These phosphorylations peaked approximately 2 to 4 h after the addition of TRAIL, concurrently with apoptosis and the development of DNA fragmentation (Fig. (Fig.1B),1B), and then tended to decrease (Fig. (Fig.1A1A).
To determine whether the activations of ATM and Chk2 by TRAIL were part of a full DDR, we looked at the activation of other DDR targets of Chk2 and ATM. Cdc25C, which is implicated in entry into G2, was phosphorylated on serine 216, and Cdc25A, which is required for progression from G1 to the S phase of the cell cycle, was degraded in response to TRAIL (Fig. (Fig.1A).1A). p53 was also phosphorylated on serine 15 (an ATM, DNA-PK, and ATR substrate) in response to TRAIL but not on serine 20 (a Chk2 substrate), and its basal levels remained unchanged (Fig. (Fig.1A).1A). p21WAF1/CIP1, one of the p53 transcriptional targets, was not upregulated but was rapidly cleaved (into an ~14-kDa fragment) in response to TRAIL (Fig. (Fig.1A)1A) (40). In contrast, E2F1 phosphorylation was not detectable and E2F1 basal levels remained unchanged in TRAIL-treated cells (Fig. (Fig.1A).1A). Moreover, activation of the other checkpoint kinase Chk1, which also phosphorylates some of the Chk2 substrates, was not detectable as measured by lack of Chk1 phosphorylation on serine 345 (Fig. (Fig.1A).1A). Together, these results indicate that TRAIL induces rapid activation of the Chk2/ATM/DNA-PK pathways without detectable activation of the E2F1, p21WAF1/CIP1, or Chk1 pathways.
Because HCT116 cells are deficient in Mre11 and in the MRN complex (a heterotrimer composed of Mre11, Rad50, and Nbs1) (15, 17) and because MRN binds DSB sites and activates the ATM and Chk2 checkpoint kinases (30), we examined whether Mre11 complementation (which also restores Rad50 and Nbs1 protein levels) (59) could have an impact on TRAIL-induced DDR activation. Using HCT116 cells stably complemented for Mre11 (HCT116-Mre11 cells), we previously showed that Mre11 complementation restores Rad50 and Nbs1 levels and enables robust Chk2 activation by ATM (59). Figure Figure1C1C shows that phosphorylations of Chk2, ATM, H2AX, and DNA-PK were induced earlier in HCT116-Mre11 cells than in wild-type HCT116 cells in response to TRAIL (see also Fig. 1D and E). The apoptotic response was also faster in the Mre11-complemented cells (compare lower panel C to panel B in Fig. Fig.1),1), and the phosphorylations of ATM, Chk2, DNA-PK, and H2AX were observed at TRAIL concentrations producing an apoptotic response (Fig. 1A, B, and F). Thus, activation of ATM, Chk2, DNA-PK, and H2AX in response to TRAIL appears to involve MRN.
Since at early times (1 to 2 h), only a fraction of the HCT116 cells underwent apoptosis (Fig. (Fig.1B)1B) and H2AX phosphorylation, we examined whether those cells that were engaging in early apoptosis were in a specific cell cycle phase. This was done by flow cytometry with the anti-γ-H2AX antibody while measuring DNA content with propidium iodide. γ-H2AX formed at all phases of the cell cycle, although with a preference for S-phase cells (Fig. (Fig.2A).2A). The preferential sensitivity of S-phase cells to TRAIL is consistent with recent studies (47). However, the induction of apoptosis by TRAIL in S-phase cells is probably not directly related to DNA replication since this induction was not affected by pretreatment with the DNA polymerase inhibitor aphidicolin (data not shown).
Activation of the DDR pathway was not limited to HCT116 colon carcinoma cells, as it was also induced in all of the other TRAIL-sensitive human cancer cell lines examined (HeLa cervical carcinoma cells [Fig. [Fig.2B],2B], Jurkat T-cell leukemia cells [Fig. [Fig.2C],2C], and M059J glioblastoma cells [see Fig. Fig.6D]),6D]), as well as in normal prostate epithelial cells (Fig. (Fig.3C3C).
Immunofluorescence confocal microscopy revealed chromatin alterations within 1 h of TRAIL exposure (Fig. (Fig.3A).3A). γ-H2AX staining in those TRAIL-responsive cells was noticeably different from the DDR induced by DNA-damaging agents. Instead of the well-defined foci distributed throughout the nucleus and characteristic of ionizing radiation (IR) (Fig. (Fig.3B;3B; see also Fig. Fig.4F)4F) and DNA-damaging drugs (42), γ-H2AX staining tended to be confluent and followed a progression that could be subdivided into three phases (Fig. 3A and B). First, γ-H2AX appeared at the nuclear periphery (Fig. (Fig.3A,3A, middle panels; type I, ring staining in panel B). Then the nuclei appeared fully stained without alteration of their overall shape or size (type II, panstaining in Fig. Fig.3B).3B). Finally, the nuclei shrank and formed apoptotic nuclear bodies that remained fully and brightly stained with γ-H2AX (Fig. (Fig.3A,3A, lower panels; type III staining in panel B). Several control experiments were performed to demonstrate that the staining observed with the γ-H2AX antibody was attributable to γ-H2AX. H2AX downregulation with siRNA resulted in a 60% reduction in γ-H2AX-positive cells compared to cells transfected with control siRNA (data not shown). Moreover, using the same antibody, we found no γ-H2AX staining in H2AX knockout cells after IR, contrary to the foci observed in H2AX-proficient cells (data not shown). In those H2AX knockout fibroblasts, we could not use TRAIL to induce γ-H2AX because normal murine embryo fibroblasts were not responsive to TRAIL. Finally, Western blotting showed only one band with the electrophoretic migration expected for γ-H2AX.
To determine the specificity of the γ-H2AX response to TRAIL for cancer cells, we also examined normal diploid human cells. Figure Figure3C3C demonstrates a similar peripheral and confluent activation of γ-H2AX in primary human prostate epithelial cells. However, only a small fraction of those cells were activated by TRAIL. γ-H2AX staining concerned less than 10% of the epithelial cells, even upon incubation for extended times (quantitation in Fig. Fig.3D).3D). The appearance of γ-H2AX after TRAIL treatment was not detectable by Western blotting (Fig. (Fig.3D),3D), demonstrating the high sensitivity of the immunofluorescence method. Together, these results demonstrate the conservation of the peripheral nuclear γ-H2AX induction (γ-H2AX ring staining) induced by TRAIL in normal cells.
To show that the γ-H2AX pattern was inside rather than around the nucleus, additional double-staining experiments were performed. Figure 4A and B shows colocalization of γ-H2AX with the heterochromatin fraction at the periphery of the nucleus (histone H3 trimethyl K9 staining). However, γ-H2AX did not localize with the heterochromatin in the nuclear interior. Double staining with lamin B1 (Fig. 4C to E) also demonstrated that the γ-H2AX pattern was inside the nucleus (see representative tracing in Fig. Fig.4E).4E). Notably, the nuclear distribution of 53BP1, a protein known to be recruited with γ-H2AX during IR-induced DSB (Fig. (Fig.4F,4F, lower panels), was not mobilized within the γ-H2AX ring by TRAIL treatment, which is indicative of the different processes leading to γ-H2AX induction by IR and TRAIL (Fig. (Fig.4F,4F, middle panels).
Together, these results demonstrate that the induction of TRAIL-induced apoptosis leads to a rapid DDR activation starting with the accumulation of phosphorylated histone H2AX (γ-H2AX) in the peripheral heterochromatic region of the nucleus, which we refer to as γ-H2AX ring staining.
Because TRAIL initiates apoptosis at the plasma membrane rather than by inducing direct DNA damage, we determined whether blocking apoptosis downstream from the TRAIL receptor could affect the DDR. First, we examined Bax, a proapoptotic member of the Bcl-2 family known to be required for TRAIL-induced apoptosis following its translocation to mitochondria (12, 28, 52) and the release of Smac/DIABLO, which antagonizes caspase-inhibiting IAP family proteins (12). In these experiments, we compared HCT116 wild-type cells (which are Bax+/−) to their Bax knockout counterparts (Bax−/− HCT116 cells) (64). Bax−/− cells are resistant to TRAIL-induced apoptosis (52) and did not show detectable phosphorylation of the DDR proteins (Fig. 5A and B), indicating that Bax is required for the induction of those phosphorylations. Also, pretreatment with the broad-spectrum caspase inhibitor Z-VAD-fmk abolished TRAIL-induced phosphorylation of Chk2, ATM, H2AX, and DNA-PK (Fig. 5C and D). These experiments indicate that DDR activation is an integral part of the apoptotic process(es) induced by TRAIL.
Since we observed that both ATM and DNA-PK were activated in response to TRAIL, and both kinases are known to phosphorylate H2AX and Chk2 (5, 15, 31, 38), we determined whether ATM and DNA-PK had overlapping or distinct roles in H2AX and Chk2 phosphorylations. Pretreatment with Nu7441, a specific DNA-PKi (27, 65), reduced the TRAIL-induced γ-H2AX response but did not affect apoptosis (Fig. (Fig.6A).6A). In contrast, the ATMi KU-55933 (19) had no effect on either H2AX phosphorylation or cell death (Fig. 6B and F). Flow cytometry analyses confirmed the reduction of γ-H2AX-positive cells by the DNA-PKi (Fig. (Fig.6C).6C). To further demonstrate that DNA-PK is critical for H2AX phosphorylation in the TRAIL pathway, we used cells genetically altered for DNA-PK. In M059J cells complemented with DNA-PK (M059J-Fus1) (15, 21), TRAIL strongly induced γ-H2AX while having almost no effect on DNA-PK-deficient M059J cells (M059J-Fus9) (Fig. (Fig.6D).6D). A similar result was observed in HCT116 cells in which DNA-PK had been knocked down by siRNA (Fig. (Fig.6E).6E). Because it was recently reported that the JNK inhibitor SP-600125 was able to prevent γ-H2AX formation in the apoptotic response induced by UVA (32), we tested SP-600125 in TRAIL-treated cells. We found that SP-600125 (20 or 200 μM) did not affect γ-H2AX induction by TRAIL (data not shown). In addition, SP-600125 did not modify TRAIL-induced apoptosis (data not shown). Together, the results of the above experiments demonstrate that DNA-PK is the primary kinase for γ-H2AX in response to TRAIL.
To identify the kinase(s) involved in T68-Chk2 phosphorylation, we again tested the DNA-PKi and ATMi. Neither was able to block T68-Chk2 phosphorylation by itself (Fig. 6A, B, and F). Pretreatment with SP-600125 (a JNK inhibitor) also had no effect (data not shown). In contrast, combination of the ATMi and DNA-PKi suppressed T68-Chk2 phosphorylation (Fig. (Fig.6F).6F). Therefore, we conclude that both ATM and DNA-PK are implicated in the phosphorylation of Chk2 at T68 in response to TRAIL whereas DNA-PK, but not ATM, is involved in H2AX phosphorylation at S139.
ATM has recently been shown to phosphorylate DNA-PK at T2609 after IR (10), indicative of cross talk between ATM and DNA-PK. After TRAIL treatment, we observed that phosphorylation of ATM on S1981 was reduced by KU-55933 (ATMi), which was expected since S1981 is a known ATM autophosphorylation site (Fig. (Fig.7A)7A) (1). However, S1981-ATM phosphorylation was also completely suppressed by Nu7441 (DNA-PKi) (Fig. (Fig.7A).7A). Conversely, phosphorylation of DNA-PK on T2609 was suppressed strongly by ATMi and completely by DNA-PKi. In contrast, phosphorylation of DNA-PK on S2056 was unmodified by the ATMi and suppressed only by the DNA-PKi (Fig. (Fig.7A).7A). To gain further evidence for the DNA-PK-dependent phosphorylation of ATM, we used siRNA against DNA-PK. Decreasing the levels of DNA-PK (Fig. (Fig.7B,7B, lower panel) strongly reduced the phosphorylation of ATM on S1981 after TRAIL treatment (Fig. (Fig.7B,7B, upper panel). From these results we conclude that, after TRAIL treatment, ATM phosphorylation on S1981 is under the control of DNA-PK. We also conclude that DNA-PK phosphorylation on T2609 is partially dependent on ATM, while DNA-PK phosphorylation on S2056 is an autophosphorylation. Thus, TRAIL induces cross phosphorylations between ATM and DNA-PK.
We then performed immunofluorescence microscopy experiments to determine whether the three phosphorylated/activated kinases (ATM, DNA-PK, and Chk2) and γ-H2AX were colocalized in cells responding to TRAIL. Figure 7C and D show that cells positive for P-Chk2-T68 were also positive for γ-H2AX, P-ATM-S1981, and P-DNA-PK-T2609. Moreover, these proteins colocalized at the periphery of the nucleus (ring staining) as described above for γ-H2AX (Fig. 3A and B). Control experiments demonstrated that the staining observed with the P-Chk2-T68 antibody was attributable to P-Chk2-T68. Indeed, Chk2 downregulation by siRNA resulted in 80% fewer P-Chk2-T68-positive cells compared to cells transfected with a control siRNA (data not shown). Moreover, using the same antibody, we found no P-Chk2-T68 staining in Chk2-deficient HCT15 cells after IR, contrary to the foci observed in Chk2-proficient cells (data not shown). The absence of P-Chk2-T68 staining (with the same antibody) after camptothecin treatment in Chk2-deficient HCT15 cells has also been published recently (44). Together, our findings demonstrate that TRAIL induces the coincident activation of ATM, DNA-PK, Chk2, and histone H2AX as individual cells induce DDR and that these phosphorylated proteins are colocalized in similar chromatin regions that define the ring staining that occurs in the early phase of apoptosis.
Because of our finding that Chk2 was robustly activated by TRAIL, and of the previously known effect of Chk2 in promoting apoptosis (20, 58), we investigated Chk2's functional involvement in TRAIL-induced apoptosis.
Using Chk2 siRNA (Fig. (Fig.8A),8A), we first observed that Chk2 downregulation had a negative impact on TRAIL-induced cell detachment, one of the early events in cells undergoing programmed cell death. As shown in Fig. Fig.8B8B (left panels with representative images and quantitations under images), the fraction of cells that remained adherent after TRAIL treatment was significantly greater in cells whose Chk2 had been downregulated by siRNA. Two hours after TRAIL addition, 49% of the Chk2 siRNA cells remained attached while only 16% of the control siRNA cells were attached. That difference persisted for up to 4 h during TRAIL treatment (Fig. (Fig.8B,8B, right panel, quantitation).
Next we determined the impact of Chk2 downregulation by siRNA on cell survival measured by clonogenic assays. Figure Figure8C8C shows that Chk2 knockdown significantly increased cell survival (~50%).
We also studied the impact of Chk2 on TRAIL-induced γ-H2AX. Figure Figure8D8D shows that downregulation of Chk2 delayed the progression of γ-H2AX staining (from type I to type III). These results suggest that Chk2 activation tends to facilitate the γ-H2AX response to TRAIL.
Finally, to determine the molecular impact(s) of Chk2 knockdown on TRAIL-induced apoptosis, we measured the activation of several caspases. Activation of caspases 2, 3, 8, and 9 was delayed in the Chk2 knockdown cells (Fig. (Fig.8E8E).
Together, the results obtained with siRNA against Chk2 demonstrate that Chk2 affects TRAIL-induced programmed cell death at several levels: kinetics of caspase activation (Fig. (Fig.8E),8E), γ-H2AX response (Fig. (Fig.8D),8D), cell detachment (Fig. (Fig.8B),8B), and ultimately cell survival (Fig. (Fig.8C8C).
To further examine the functional role of Chk2 in apoptosis, we also performed experiments with Chk2 knockout cells (24). Figure Figure9A9A shows that apoptosis measured by sub-G1 response was significantly reduced in Chk2−/− cells.
Finally, based on our finding that TRAIL-induced activation of the (Chk2/ATM/DNA-PK) DDR pathway invoked a partial p53 response, as determined by phosphorylation of p53 at S15 (Fig. (Fig.1A),1A), the role of p53 in TRAIL-induced cell killing was examined with isogenic p53−/− and p53+/+ HCT116 cells (4). Caspase activation was comparable in p53−/− and p53+/+ cells (data not shown), and attenuation of this caspase response by Chk2 downregulation was similar in both cell types (Fig. (Fig.9B),9B), indicating that the proapoptotic effect of Chk2 is p53 independent.
Our study focused on the involvement of Chk2 and histone H2AX in response to DR activation by TRAIL. We show that Chk2, following its activation by DNA-PK and ATM, can amplify the apoptotic response elicited by TRAIL by a feedback loop independently of p53. The Chk2-mediated positive feedback extends outside the nucleus, as it involves upstream caspases 8 and 9. Caspase 8 forms a complex with the TRAIL membrane receptors (DR4 and DR5) within the DISC, and caspase 9 is a key effector of the apoptosome. Another TRAIL-induced feedback loop implicating caspase 8 has recently been described, which involves the nuclear translocation of a self-cleaved death effector domain segment of caspase 8 and p53-dependent upregulation of procaspase 8 gene expression (62). Our feedback mechanism would act more directly on caspases than the p53-dependent transcriptional activation of procaspase 8 (62). Figure Figure1010 gives an outline of the TRAIL-induced DDR pathways and schematizes the feedback loops that are revealed in our study.
The functional impact of Chk2 on TRAIL-induced apoptosis was detectable as an enhancement of cell detachment, an increased cell death measured by clonogenic assays, an increased γ-H2AX induction, and a faster activation of caspases (Fig. (Fig.8).8). In fact, TRAIL was remarkably potent in HCT116 cells. Even after only 1 h of TRAIL exposure, only ~1% of the cells were able to form colonies (Fig. (Fig.8C).8C). Therefore, the cells progress to apoptosis once they have been committed upon TRAIL exposure. Inactivation of Chk2 increased the number of surviving colonies by 40 to 60% (Fig. (Fig.8C),8C), demonstrating the importance of Chk2 in eliminating residual cells. The functional impact of Chk2 may be quite significant because of the potentially severe physiological consequences of abortive apoptosis. Apoptosis has been viewed as an important mechanism to eliminate oncogenes from dying tumor cells or virus-infected cells. Also, even when cells die, it is important that apoptosis proceed to completion to avoid autoimmune diseases, which have been linked to circulating DNA fragments (39, 63). Thus, it is possible that Chk2 promotes the completion of the apoptosis program, thereby avoiding the survival of abnormal cells and the release of toxic metabolic intermediates.
To our knowledge, our study is the first to implicate Chk2 in TRAIL-induced apoptosis independently of p53. Prior studies had provided functional evidence for the proapoptotic role of Chk2 in the context of DNA damage produced by IR, radiomimetic agents, and arsenic trioxide (20, 23, 25, 58). In those models, the proapoptotic function of Chk2 was linked to p53 activation. In contrast, in our study, the proapoptotic function of Chk2 is p53 independent. p53 was phosphorylated on serine 15 but neither phosphorylated on serine 20 nor stabilized as determined by lack of p53 protein accumulation in TRAIL-treated cells (Fig. (Fig.1A).1A). Also, p21WAF1/CIP1 was not upregulated by TRAIL (Fig. (Fig.1A).1A). These results are in accordance with the literature showing that phosphorylation on serine 20 is required for stabilization of p53 (9) and enhancement of the transcriptional activation of p21WAF1/CIP1 by p53 (22). p21WAF1/CIP1 was rapidly cleaved in response to TRAIL (Fig. (Fig.1A),1A), probably due to caspase 3 activity (40). The degradation of p21WAF1/CIP1 could facilitate apoptosis (36). We also found that the proapoptotic role of Chk2 was unaffected in the absence of p53 (Fig. (Fig.9).9). Thus, our findings are consistent with other studies showing that p53 is not essential for TRAIL-induced apoptosis (45, 46).
A main focus of our study is the H2AX response to TRAIL. We show that histone H2AX is rapidly phosphorylated on serine 139 (γ-H2AX) and that γ-H2AX tends to form a confluent staining that initiates in peripheral nuclear regions (γ-H2AX ring staining) before gross changes in nuclear morphology that characterize apoptosis (Fig. (Fig.3,3, ,4,4, ,5,5, ,7,7, and and8).8). We had previously shown γ-H2AX formation in response to Fas antibody, staurosporine, and DNA-damaging agents (50) soon after the discovery of γ-H2AX (51). But at that time, we did not investigate the nuclear distribution of γ-H2AX by immunofluorescence microscopy. In the present study, several novel points are noteworthy regarding TRAIL-induced γ-H2AX activation. It is remarkably rapid (starting within 1 h), while the cells retain an overall normal morphology. Only a fraction of the cells initially show γ-H2AX staining, and those cells tend to be in S phase, although not exclusively (Fig. (Fig.2A).2A). The initial γ-H2AX staining initiates as a confluent pattern at the periphery of the nucleus (γ-H2AX ring staining) before diffusing to form a panstaining pattern covering the entire nucleus and, later on, the nuclear bodies (Fig. (Fig.3,3, ,4,4, and and8).8). This confluent pattern was also observed in normal epithelial cells treated with TRAIL, although in much fewer cells than in the cancer cells examined here (Fig. 3C and D), which suggests conservation of the γ-H2AX induction process in both cancer and normal cells. The γ-H2AX peripheral distribution (γ-H2AX ring staining) occurred within the peripheral heterochromatic regions at the contact of the lamin B1 nuclear boundary (Fig. (Fig.4).4). The TRAIL-induced γ-H2AX patterns never formed detectable nuclear foci and were not accompanied by the recruitment/colocalization of 53BP1, another ubiquitous DDR factor, which sets apart the TRAIL-induced γ-H2AX response from the focal response(s) characteristic of DNA-damaging agents (49). The ring staining for γ-H2AX also contained activated ATM phosphorylated on S1981, activated DNA-PK phosphorylated on T2609, and activated Chk2 phosphorylated on T68 (Fig. (Fig.7C).7C). Further studies are under way to determine whether this novel γ-H2AX response is induced during other apoptotic processes besides TRAIL-induced DR pathway activation.
We found that Chk2 has an impact on the progressive induction of γ-H2AX staining in response to TRAIL. Chk2 siRNA slowed down the formation of γ-H2AX-positive cells with nuclear segmentation (type III staining) (Fig. (Fig.8D).8D). This reduction could result from delayed caspase activation (Fig. (Fig.8E).8E). However, it is also plausible that Chk2 acts directly on chromatin. In fact, histone H1 is a substrate for Chk2 (67). Thus, H1 phosphorylation by Chk2 might be part of the chromatin modifications controlled by Chk2. This possibility is attractive in the context of recent studies showing that Chk2 activation by DNA damage induces the release of H1.2 from chromatin and its translocation to the mitochondria, where H1.2 can act as a positive regulator of apoptosome formation (11, 54).
Our experiments demonstrate that the kinase responsible for the induction of γ-H2AX by TRAIL is DNA-PK (Fig. (Fig.6).6). In contrast, Chk2 is phosphorylated by both ATM and DNA-PK in response to TRAIL. Thus, the ATM and DNA-PK pathways appear to branch out upstream from Chk2 and H2AX (Fig. (Fig.10).10). Our finding that DNA-PK is the primary γ-H2AX kinase during TRAIL-induced apoptosis is consistent with another study showing that DNA-PK is responsible for γ-H2AX formation during staurosporine-induced apoptosis (38). However, contrary to a study of UVA-induced apoptosis (32), we found that JNK kinase is not involved in γ-H2AX induction by TRAIL and that knocking out H2AX by siRNA had no effect on TRAIL-induced apoptosis (data not shown). These differences suggest the existence of various nuclear pathways activated during apoptosis. The general dispensability of H2AX for the execution of apoptosis is actually consistent with the viability and normal embryonic development of H2AX knockout mice (7).
To investigate the potential involvement of ATR in the TRAIL pathway, we looked at the activation of Chk1, which is a preferential ATR substrate. Activation of Chk1 following its cleavage by caspases has been reported to contribute to Chk1-dependent apoptosis in response to DNA-damaging agents (34). Our present study provides no evidence for Chk1 implication in TRAIL-induced apoptosis (Fig. (Fig.1A).1A). Also, inhibition of ATM and DNA-PK was sufficient to suppress completely the activation of Chk2 and H2AX (Fig. (Fig.6F).6F). Together, these data suggest that ATM, DNA-PK, and Chk2 are the main kinases involved in the TRAIL-induced response, contrary to ATR and Chk1. Nevertheless, we cannot exclude a potential involvement of ATR. Even if Chk1 was not phosphorylated, other proteins could be targeted by ATR.
The observed cross talk between ATM and DNA-PK following TRAIL treatment is worth noting because it appears to be mutual rather than limited to only one direction in which DNA-PK is phosphorylated by ATM (10). Our present finding that ATM can be phosphorylated on S1981 in a DNA-PK-dependent manner (Fig. (Fig.7A)7A) is novel. An effect of DNA-PK on ATM has nevertheless been suggested by some prior studies. In glioblastoma cells deficient for DNA-PK (M059J), the ATM protein levels are low (8, 16). In addition, the V3 radiosensitive CHO cell line presents low ATM protein levels, and when the amount of DNA-PK is restored by transfection, the levels of ATM protein are also restored (41). In murine cells with different degrees of DNA-PK deficiency, ATM protein is reduced (41).
From a therapeutic standpoint, our findings suggest that the levels of Chk2 and activated P-Chk2-T68 in tumor tissues could have a prognostic value for predicting the efficiency of a TRAIL therapy. Moreover, our results provide a rationale for combining TRAIL with DNA-damaging agents. Indeed, DNA damage (IR or DNA-targeted drugs) might sensitize tumor cells to TRAIL by preactivating Chk2 and the positive feedback loop that amplifies TRAIL-induced apoptosis. Such a mechanism may partly account for the known synergism between TRAIL and DNA-damaging therapies (47, 61). Chk2 activation in some human tumors and precancerous lesions (3, 13, 18) could also delay or prevent cancer development, reinforcing the importance of an efficient DDR.
This research was supported in part by the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research.
We thank Susan Garfield (Laboratory of Experimental Carcinogenesis, Center for Cancer Research, NCI) for outstanding technical assistance for the generation of 3D microscopy pictures. We also thank Giovanni Capranico for editorial comments.
Published ahead of print on 27 October 2008.