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[Cancer Research 66, 9235-9244, September 15, 2006]
© 2006 American Association for Cancer Research

Experimental Therapeutics, Molecular Targets, and Chemical Biology

Identification of New Compounds That Trigger Apoptosome-Independent Caspase Activation and Apoptosis

Emanuela Aleo, Clare J. Henderson, Alessandra Fontanini, Barbara Solazzo and Claudio Brancolini

Dipartimento di Scienze e Tecnologie Biomediche, Sezione di Biologia and MATI Center of Excellence, Universita' di Udine, Udine, Italy

Requests for reprints: Claudio Brancolini, Dipartimento di Scienze e Tecnologie Biomediche, Sezione di Biologia, Universita' di Udine. P.le Kolbe 4, 33100 Udine, Italy. Phone: 39-0432-494382; Fax: 39-0432-494301; E-mail: cbrancolini{at}makek.dstb.uniud.it.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Identification of alternative pathways of caspase activation is an important step to develop new antitumor treatments. We report here the result of a screening with a small chemical library, the Developmental Therapeutics Program-National Cancer Institute "challenge set," on cells expressing mutated caspase-9. We have identified two molecules capable of activating an apoptosome-independent apoptotic pathway. These compounds, named F6 and G5, target the ubiquitin-proteasome system by inhibiting the ubiquitin isopeptidases. We have shown that F6 and G5 induce a rather unique apoptotic pathway, which includes a Bcl-2-dependent but apoptosome-independent mitochondrial pathway with up-regulation of the BH3-only protein Noxa, stabilization of the inhibitor of apoptosis antagonist Smac, but also the involvement of the death receptor pathway. Noxa plays an important role in the induction of mitochondrial fragmentation and caspase activation, whereas the death receptor pathway becomes critical in the absence of a functional apoptosome. This study suggests that screening of chemical libraries on cancer cells with defined mutations in apoptotic key elements can lead to the identification of compounds that are useful to characterize alternative pathways of caspase activation. (Cancer Res 2006; 66(18): 9235-44)


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Apoptosis is the most common mechanism through which many chemotherapeutic drugs kill neoplastic cells and promote tumor eradication (1). Although it is now emerging that cells can also die by nonapoptotic mechanisms, apoptosis is regarded the preferred mechanism for chemotherapeutic drugs to kill tumor cells. In fact, apoptosis resistance plays a critical role in the efficiency of chemotherapy, with chemoresistance and tumor progression often arising when cancer cells accumulate mutations in the critical genes regulating the apoptotic program (1, 2).

Apoptosis relies on the timing activation of caspases, a group of cysteine proteases that cleave selected cellular substrates after aspartic residues (3). Two main apoptotic pathways keep in check caspase activation. The extrinsic pathway is triggered by the activation of a family of death receptors at the cell surface and controls the activation of caspase-8, whereas the intrinsic pathway is triggered by the release of killer proteins, such as cytochrome c and Smac/DIABLO (4–6), and induces the assembly of the apoptosome and the activation of caspase-9.

The mitochondrial or intrinsic apoptotic pathway governs cell death in response to various cellular stresses, including many of the available antitumor treatments. Genetic mutations in critical elements of the mitochondrial pathway are often observed in tumors and correlate with chemoresistance (7). To identify new compounds able to induce caspase activation and cell death by alternative mechanisms, we have used cells expressing a dominant-negative form of caspase-9 (C9DN), which interferes with the formation of a functional apoptosome (8). Fifty-seven compounds that constitute the challenge set of the Developmental Therapeutics Program (DTP) from the National Cancer Institute (NCI) United States (US), with unknown mechanism of cell killing, were evaluated for their ability to trigger caspase activation and apoptosis in apoptosome-defective cells. We have identified two compounds, named F6 and G5, which efficiently activate caspases and apoptosis in the absence of a functional apoptosome. These molecules target the ubiquitin-proteasome system by inhibiting the ubiquitin isopeptidase activities. In this work, we have characterized the apoptotic pathway activated by F6 and G5.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Culture conditions, apoptosis, and transfection. IMR90-E1A (E1A) and E1A/C9DN cells have been described previously (8). All the cell lines, with the exception of the BJ, were grown in DMEM supplemented with 10% FCS. BJ human foreskin fibroblasts expressing the different oncogenes (9, 10) were maintained in MEM with Earle's salts supplemented with nonessential amino acids and 10% fetal bovine serum at 37°C in 5% CO2. In all trypan blue exclusion assays, 160 to 400 cells from three independent samples were counted for each data point. Data were represented as arithmetic mean ± SD for at least three independent experiments.

Stealth RNA interference (RNAi) were purchased from Invitrogen (Carlsbad, CA): Noxa RNAi, GCTGGAAGTCGAGTGTGCTACTCAA (bp 233-257); Apaf-1 RNAi, GGGTACAGTACTTTCTTGTGACATT (bp 3255-3279); histone deacetylase 4 (HDAC4) RNAi, CCACCGGAATCTGAACCACTGCATT (bp 561-585); and the control RNAi, TGCGTTAGTAACGATTTCATTGGAA. Cells were transfected 24 hours after plating by adding the Opti-MEM medium containing LipofectAMINE (Invitrogen) plus the stealth RNAi oligos.

Different drug concentrations were used to determine the dose-response profile and to calculate the IC50 value, the concentration used to obtain 50% of cell death, after 24 hours of treatment. For drug screening, cells were seeded in 96-wells plates at a density of 5 x 104/mL per well in 0.1 mL of medium. After 24 hours, cells were incubated with the required concentrations of the 57 compounds that constitute the challenge set obtained from the DTP-NCI-US.1 After 24 or 48 hours of incubation, cell death was evaluated by trypan blue staining.

Subcellular fractionation. Cells were plated (5 x 104/mL) into 58 cm2 Petri dishes, and 24 hours later, they were treated with the different compounds. After 20 hours, apoptotic and adherent cells were collected in the lysis buffer (1 mmol/L MgCl2, 1 mmol/L EDTA, 1 mmol/L EGTA, 20 mmol/L HEPES, 250 mm/L sucrose, 75 µg/mL digitonin) in the presence of a cocktail of protease inhibitors. Lysates were centrifuged at 13,000 rpm to separate the pellet containing mitochondria from the cytosolic fraction.

Western blotting. Proteins obtained after a SDS denaturating lysis and sonication were transferred to a 0.2-µm pore size nitrocellulose membrane and incubated with the following antibodies: anti-caspase-2, anti-Smac/DIABLO (9), anti–green fluorescent protein (GFP; ref. 11), anti–poly(ADP-ribose) polymerase (PARP; Cell Signaling, Beverly, MA), anti-p85 PARP fragment (Promega, Madison, WI), anti-tubulin, anti–cytochrome c (Cell Signaling), anti-HDAC4 (11), anti-Apaf-1, anti-hemagglutinin (HA; Sigma, St. Louis, MO), anti-F1-ATP synthase, anti-Mcl-1 (Cell Signaling), anti-Bim (Cell Signaling), anti-Noxa (Calbiochem, San Diego, CA), and anti-Bcl-2 (Sigma).

In vitro proteolytic assays. For proteasome and ubiquitin isopeptidase activity assays, two fluorescent substrates were used: Suc-LLVY-AMC and z-LRGG-AMC, respectively (Bachem, Bubendorf, Switzerland). Cells were lysed in 25 mmol/L HEPES (pH 7.5), 5 mmol/L EDTA, and 0.1% CHAPS for 30 minutes at 4°C. The protein concentration was adjusted to 1 mg/mL. For the isopeptidase activity assay, cell lysates were preincubated for 30 minutes at 4°C with 10 µmol/L MG-132 to minimize the background proteolytic activity of the proteasome (12). Next, the different compounds were added for 30 minutes at 4°C. Finally, the assay mixture was incubated with the substrates at 37°C and the AMC fluorescence was quantified by exciting the samples at {lambda} = 360 nm and monitoring the emission at {lambda} = 460 nm. Fluorogenic caspase-3/caspase-7 peptide substrate assays (Promega) were done as described previously (9).

Immunofluorescence microscopy. Mitochondria were labeled in vivo for 1 hour with 25 nmol/L MitoTracker Red CMXRos (Molecular Probes, Eugene, OR). After 12 hours of treatment with the different compounds, cells were fixed with 3% paraformaldehyde in PBS for 20 minutes at room temperature. Fixed cells were washed with PBS/0.1 mol/L glycine (pH 7.5) and then permeabilized with 0.1% Triton X-100 in PBS for 5 minutes. The coverslips were treated with the anti-Smac antibodies diluted in PBS for 30 minutes in a moist chamber at 37°C. They were then washed twice with PBS and incubated with Alexa Fluor 488–conjugated secondary antibodies (Molecular Probes) for 30 minutes at 37°C. Cells were examined by epifluorescence with a Zeiss (Oberkochen, Germany) Axiovert 35 microscope or with a Leica (Wetzlar, Germany) SP laser scan microscope equipped with a 488-nm argon laser and a 543-nm helium neon laser.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Identification of two compounds capable of triggering caspase activation in the presence of mutated caspase-9. To identify new compounds capable of triggering caspase activation and apoptosis with high efficiency even in the presence of a nonfunctional apoptosome, the challenge set (DTP-NCI), consisting of 57 molecules with unknown mechanisms of cell killing, was tested for the ability to induce apoptosis in E1A cells. All the different compounds were initially tested to define their IC50 (50% of apoptosis) in E1A cells. The range of concentration used varied from 0.01 to 40 µmol/L depending on the compound; the appearance of apoptosis was scored up to 48 hours from treatment. Once the IC50 was defined for each compound, we tested the caspase activity in E1A and in E1A/C9DN cells, which express C9DN (8). The activity of caspase-3/caspase-7 was measured with the z-DEVD-R110 fluorogenic substrate. After this screening (Fig. 1A ), two compounds were identified: 4H-thiopyran-4-one, tetrahydro-3,5-bis[(4-nitrophenyl)methylene]-,1,1-dioxide (NSC 144303) and 4-piperidione,3,5-bis[(4-methylphenyl)methylene]-hydrochloride (NSC 632839), both capable of sustaining caspase-3/caspase-7 activity in the absence of a functional caspase-9. For simplicity, NSC 144303 and NSC 632839 have been nominated G5 and F6, respectively. As summarized in Fig. 1B, F6 and G5 were able to activate caspase-3/caspase-7 with similar intensity in E1A and E1A/C9DN cells; we used the proteasome inhibitor MG-132 as a caspase-9-independent positive control (9) and etoposide as a caspase-9-independent negative control (8). In accordance with the data from the fluorogenic assays, immunoblots showed similar amounts of processed caspase-2 and PARP in E1A and E1A/C9DN cells after treatment with F6, G5, or MG-132. For etoposide, both caspase-2 and PARP processing were drastically reduced in E1A/C9DN cells (Fig. 1C). Similarly to MG-132, F6 and G5 induced cell death at comparable levels in both cell lines, whereas etoposide, as expected, triggered apoptosis less efficiently in E1A/C9DN cells (Fig. 1D). Finally, the induction of apoptosis was confirmed by Annexin V/propidium iodide staining and flow cytofluorimetric detection (Supplementary Fig. S1).


Figure 1
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  		Figure 1. Identification of two compounds capable of triggering caspase activation and apoptosis with high efficiency in caspase-9-mutated cells. A, chemical structures and NSC numbers for F6 and G5. The flow chart summarizes the screening procedure adopted to evaluate the 57 compounds of the challenge set (DPT-NCI) for the ability to activate caspases in caspase-9-defective cells. B, DEVDase activity in E1A and E1A/C9DN cells treated for 20 hours with the indicated agent. Columns, mean (n = 3); bars, SD. C, processing of caspase-2 and PARP in E1A and E1A/C9DN cells treated with the indicated drugs for 20 hours. Equal amounts of cell lysates were subjected to SDS-PAGE. Immunoblots were done using the indicated antibodies. Tubulin was used as loading control. D, E1A and E1A/C9DN cells were treated with the indicated apoptotic insults for 20 hours, and the appearances of apoptosis were scored by trypan blue staining. Columns, mean (n = 3); bars, SD. Concentrations used were as follows: F6 (10 µmol/L), G5 (1.25 µmol/L), MG-132 (5 µmol/L), and etoposide (ETO; 50 µmol/L).

 
The IC50 (the concentration required to obtain 50% of cell death) for F6 was 15.65 and 16.23 µmol/L in E1A and E1A/C9DN cells, respectively, whereas G5 is a more potent apoptotic inducer showing a IC50 of 1.76 and 1.6 µmol/L in E1A and E1A/C9DN cells, respectively (Supplementary Fig. S2). We also confirmed that F6 and G5 induce apoptosis with undistinguishable kinetics in E1A and E1A/C9DN cells (Supplementary Fig. S3).

F6 and G5 can induce caspase activation and apoptosis independently from Apaf-1. We down-regulated Apaf-1 expression in E1A cells to further confirm the ability of F6 and G5 to overcome the apoptosome. Stealth RNAi oligos for Apaf-1 and HDAC4, selected as a control (Fig. 2A ), were transfected in E1A cells. The apoptotic response to F6, G5, MG-132, and etoposide was measured by trypan blue staining. As expected, down-regulation of Apaf-1 significantly reduced the apoptotic response of E1A cells to etoposide but not to F6, G5, or MG-132 (Fig. 2B). Fluorogenic peptide cleavage assays confirmed that silencing Apaf-1 does not significantly affect the activity of caspase-3/caspase-7 when cells were challenged with F6, G5, or MG-132. By contrast, the presence of the Apaf-1 RNAi oligos clearly impaired the ability of etoposide to trigger caspase activation (Fig. 2C). The proteolytic processing of caspase-2 and PARP (Fig. 2D) was used to confirm that F6 and G5, but not etoposide, induced efficient caspase activation and apoptosis in Apaf-1-depleted cells. These data confirm that F6 and G5 can efficiently trigger caspases activation and apoptosis in the absence of a functional apoptosome.


Figure 2
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  		Figure 2. Apaf-1 is dispensable for F6- and G5-induced apoptosis. A, Western blot of E1A cell lysates treated with stealth RNAi oligos specific for Apaf-1 or HDAC4 as indicated. B, E1A cells transfected with Apaf-1 or HDAC4-specific RNAi oligos were incubated for 20 hours with the indicated apoptotic insults. Appearance of apoptosis was scored by trypan blue staining. Columns, mean (n = 4); bars, SD. C, DEVDase activity in E1A cells transfected with RNAi oligos specific for HDAC4 or Apaf-1 and treated for 20 hours with the indicated molecules. Columns, mean (n = 3); bars, SD. D, processing of caspase-2 and PARP in E1A cells transfected with RNAi oligos against HDAC4 or Apaf-1 and treated for 20 hours with the different apoptotic insults as indicated. Equal amounts of cell lysates were subjected to SDS-PAGE. Immunoblots were done using the indicated antibodies. Tubulin was used as loading control. Concentrations used were as follows: F6 (10 µmol/L), G5 (1.25 µmol/L), MG-132 (5 µmol/L), and etoposide (50 µmol/L).

 
Bcl-2 family members control F6- and G5-induced apoptosis. Transformed cells display a greater susceptibility to drug-induced apoptosis than nonmalignant cells (1). Accordingly, we have observed that primary human fibroblasts are more resistant to apoptosis induced by F6 or G5 compared with the counterpart of E1A-transformed cells (Fig. 3A ; Supplementary Fig. S1).


Figure 3
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  		Figure 3. Cellular transformation and Bcl-2 family members regulate apoptosis in response to F6 and G5. A, IMR90 and E1A-transformed fibroblasts were grown for the indicated hours in the presence of F6 (10 µmol/L) or G5 (1.25 µmol/L). Appearance of apoptosis was scored by trypan blue staining and by caspase-3/caspase-7 activities. Columns, mean (n = 3); bars, SD. B, BJ/E1A/Ras and BJ/E1A/Ras/Bcl-2 cells were treated with F6 (10 µmol/L), G5 (1.25 µmol/L), or etoposide (50 µmol/L) as a control for the indicated hours. Appearance of apoptosis was scored by trypan blue staining and caspase-3/caspase-7 activities. Columns, mean (n = 3); bars, SD. C, MEFs from WT or bax–/–bak–/– mice were treated with the indicated compounds for 20 hours, and the appearance of apoptosis was scored by trypan blue staining and caspase-3/caspase-7 activities. Columns, mean (n = 3); bars, SD. Two different concentrations of F6 (20 and 40 µmol/L) and G5 (1.25 and 2.5 µmol/L) were applied in this analysis. Etoposide (50 µmol/L) and MG-132 (5 µmol/L) were used as controls.

 
We next investigated the role of Bcl-2 in the apoptotic pathway elicited by F6 and G5. We used BJ cells transformed with E1A, Ras, and expressing Bcl-2. As shown in Fig. 3B and Supplementary Fig. S4, cells overexpressing Bcl-2 were protected by F6- and G5-induced cell death. Bcl-2 protection was also evident in a long-term survival assay (Supplementary Fig. S5). This indicates that Bcl-2 protects cells from F6- and G5-dependent apoptosis and that these cells are not damaged and can resume proliferation.

Multidomain proapoptotic proteins Bax and Bak are required for the apoptotic response to many insults (13). To unveil the role of these proteins during F6- and G5-dependent apoptosis, immortalized wild-type (WT) murine embryonic fibroblasts (MEF) and bax–/–bak–/– MEFs were treated with F6 and G5. Both F6 and G5 require Bax and Bak to induce cell death (Fig. 3C; Supplementary Fig. S6). Of note, a concentration up to 40 µmol/L of F6 was necessary to efficiently kill transformed MEFs.

The role of the extrinsic pathway in the apoptotic response to F6 and G5. To investigate the role of the extrinsic pathway in the apoptotic response to F6 and G5, we used competitors of caspase-8 activation, such as c-FLIPS or the dominant-negative form of FADD and CrmA, the natural inhibitor of caspase-8 catalytic activity. The different cDNAs were transfected in E1A and E1A/C9DN cells together with enhanced GFP (EGFP) as a reporter; after treatment with the indicated stressors, apoptosis was measured by scoring the appearance of membrane blebbing and cell collapse in the EGFP-positive cells.

Expression of CrmA, c-FLIPS, and FADD/DN does not influence cell death in response to F6, G5, MG-132, or etoposide in E1A cells, whereas, as expected, apoptosis in response to tumor necrosis factor-{alpha} (TNF-{alpha}) was impaired (Fig. 4 ). This result shows that the death receptor pathway is not critical for the apoptotic response to F6 and G5 in the presence of a functional apoptosome. By contrast, expression of CrmA, c-FLIPS, or FADD/DN in E1A/C9DN cells partially hampered the apoptotic response to F6, G5, and MG-132. These results reveal that the death receptor pathway can compensate for the absence of a functional apoptosome when apoptosis is induced by F6, G5, or MG-132 but not by etoposide.


Figure 4
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  		Figure 4. Role of the extrinsic pathway in the apoptotic response to F6 and G5. The different cDNAs (LacZ, CrmA, FADD/DN, and c-FLIPS) were transiently expressed in E1A or E1A/C9DN cells together with EGFP as a reporter. Cells were treated with MG-132, F6, G5 (12 hours), etoposide (20 hours), or CHX and TNF-{alpha} (40 hours), and green cells showing collapsed morphology and presenting extensive membrane blebbing were scored as apoptotic. Columns, mean (n = 3); bars, SD. Concentrations used were as follows: F6 (10 µmol/L), G5 (1.25 µmol/L), MG-132 (5 µmol/L), etoposide (50 µmol/L), CHX (2 µg/mL), and TNF-{alpha} (80 ng/mL).

 
F6 and G5 interfere with the ubiquitin-proteasome system. Careful inspection for the presence of structural motifs that could predict the cellular targets of F6 and G5 allowed us to identify a recent characterized molecular determinant that confers the ability to inhibit the ubiquitin-proteasome system. F6 and G5 are typified by a cross-conjugated {alpha},ß-unsaturated dienone with two sterically accessible electrophilic ß-carbons, a determinant that confers isopeptidase inhibitory activity (Fig. 5A ; refs. 12, 14–16).


Figure 5
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  		Figure 5. Effect of F6 and G5 on the ubiquitin-dependent proteolysis. A, chemical structure of F6 and G5 that evidences the sterically accessible ß-carbons. B, accumulation of polyubiquitins in E1A cells expressing HA-tagged ubiquitin and treated for 12 hours with the indicated molecules. EGFP was used to evaluate the transfection efficiency. Concentrations used were as follows: F6 (10 µmol/L), G5 (1.25 µmol/L), MG-132 (5 µmol/L), DPTP (30 µmol/L), and etoposide (50 µmol/L). C, cellular lysates from E1A cells (1 µg/µL) were incubated with DMSO (UNT) or with the indicated concentrations of F6, G5, DBA, DPTP, or MG-132. After 30 minutes, the substrate (Suc-LLYY-AMC) for the catalytic particle of the proteasome was added. Columns, mean (n = 3); bars, SD. D, cellular lysates obtained from E1A cells (1 µg/µL) were incubated with DMSO (UNT) or with the indicated concentrations of F6, G5, DBA, and DPTP. After 30 minutes, the isopeptidase substrate z-LRGG-AMC was added. To prevent proteasome-mediated degradation of the isopeptidase substrate, cell lysates were preincubated for 30 minutes with 10 µmol/L MG-132. Columns, mean (n = 3); bars, SD. *, P < 0.013, statistically significant differences between cellular lysates incubated with F6 and DPTP (60 µmol/L).

 
To confirm that F6 and G5 can inhibit the ubiquitin-proteasome system, we tested whether an accumulation of polyubiquitins could be observed in cells after treatment with these compounds. E1A cells transfected with HA-tagged ubiquitin were treated with F6, G5, DBA, a previously characterized isopeptidase inhibitor, DPTP, a compound with sterically hindered ß-carbons that does not inhibit isopeptidase activity (12), MG-132, and etoposide. EGFP was coexpressed as a transfection control efficiency. Figure 5B shows that only F6, G5, and DBA, similarly to the proteasome inhibitor MG-132, provoke the accumulation of polyubiquitins, whereas the other compounds do not.

To confirm the ability of F6 and G5 to target the isopeptidases, we first excluded that F6 and G5 could inhibit the catalytic activity of the proteasome. We examined hydrolysis of a substrate for the chymotrypsin-like (Suc-LLVY-AMC) sites of the 20S catalytic subunit of the proteasome, which play a pivotal role in protein cleavage (17, 18). Addition of F6 or G5 to cell extracts obtained from E1A cells did not inhibit the proteolytic activity of the proteasome, whereas MG-132, as expected, potently inhibited the cleavage of the LLVY-AMC substrate (Fig. 5C). Next, we measured inhibition of isopeptidase activity by using a previously described assay (14), which relies on the fluorogenic peptide z-LRGG-AMC that mimics the ubiquitin COOH-terminal isopeptide linkage. In this case, a concentration-dependent inhibition of the isopeptidase substrate cleavage was observed when cell extracts were incubated with F6, G5, or DBA (Fig. 5D). As expected, the sterically hindered compound DPTP was ineffective as an isopeptidase inhibitor (12).

F6 and G5 increase the level of cytosolic Smac/DIABLO and of the Bcl-2 proteins regulated by the ubiquitin-proteasome system. The expression of various elements of the apoptotic machinery is under the tight control of the ubiquitin-proteasome system (19). Among them, various data indicate that the inhibitor of apoptosis (IAP) antagonist Smac/DIABLO could be targeted for degradation by the action of the RING finger IAPs after release in the cytosol (20). Hence, we analyzed the levels of cytosolic Smac when apoptosis was induced by F6 and G5 in comparison with etoposide. Subcellular fractionations were done to isolate the mitochondrial and the cytosolic fractions from E1A cells treated with F6, G5, MG-132, or etoposide. We used doses of the different compounds that had previously been tested to trigger comparable apoptosis. Trypan blue scoring, HDAC4, and PARP processing as detected by immunoblot confirmed that apoptosis was similarly induced by the different insults (Fig. 6A ).


Figure 6
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  		Figure 6. F6 and G5 regulate the levels of different elements of the apoptotic machinery. A, subcellular fractionation of E1A cells treated or not (Untr) for 20 hours with the indicated compounds by separating soluble cytosolic fractions (SNs) from digitonin insoluble fractions, including mitochondria (Pellets). Western blots were done using the indicated antibodies. Bottom, apoptosis measured by trypan blue assay. B, time course analysis of Mcl-1, Bcl-2, Noxa, and Bim expression. Lysates from E1A cells treated with the different compounds for the indicated times were prepared and subjected to immunoblot analysis using the specific antibodies. C, Western blot of E1A/C9DN cell lysates either treated or not for 20 hours with F6 and G5 shows the effect of Noxa-specific stealth RNAi oligos or control (C) oligos on Noxa and Smac expression. D, E1A/C9DN cells transfected with Noxa RNAi oligos or with control ones were incubated for 20 hours with F6, G5, and MG-132 or maintained untreated. Appearance of apoptosis was measured by trypan blue staining. Columns, mean (n = 3); bars, SD. E, DEVDase activity in E1A/C9DN cells transfected with Noxa RNAi oligos or with control ones. Cells were incubated for 20 hours with F6, G5, and MG-132 or maintained untreated. Columns, mean (n = 3); bars, SD. Concentrations used were as follows: F6 (10 µmol/L), G5 (1.25 µmol/L), MG-132 (5 µmol/L).

 
Surprisingly, we noted a second form of Smac/DIABLO with reduced mobility in SDS-PAGE in the pellet fraction obtained from cells treated with the inhibitors of the ubiquitin-proteasome system (Fig. 6A, asterisk). This form displays the predicted size of the pro-Smac, which includes the NH2-terminal presequence for mitochondrial targeting. It is interesting to note that the ability of proteasome inhibitors to alter the import of mitochondrial preproteins has already been documented (21). Further studies are required to clarify if this additional form of Smac represents its precursor.

Most importantly, the levels of Smac in the cytosolic fractions were higher when apoptosis was elicited by F6, G5, or MG-132 compared with etoposide. The levels of cytochrome c and the rate of HDAC4 processing were similar under the different apoptotic insults. This suggests that differences in the amounts of cytosolic Smac cannot be attributed to differences in the permeability of the mitochondrial outer membrane.

The expression of some Bcl-2 family members is modulated by the proteasome (19). To further confirm the action of F6 and G5 on the ubiquitin-proteasome system, we investigated whether these compounds could modulate the levels of the Bcl-2 family members proven to be under the proteasome control. We focused our attention on Mcl-1, Noxa, Bcl-2, and Bim. F6 and G5 dramatically increased the levels of Noxa and Mcl-1 in E1A cells during a time course analysis. As expected, also MG-132 induced higher levels of Mcl-1 and Noxa, whereas etoposide did not. In our cell line, Bim was not up-regulated in response to alterations of the ubiquitin-proteasome system (Fig. 6B). This probably reflects its tissue-specific modulation by the proteasome (22). Accordingly to recent data, Bcl-2 was not modulated by proteasome inhibitors during the time of the analysis (23, 24).

To determine the role of Noxa during F6- and G5-induced apoptosis, its expression was down-regulated by RNAi in E1A/C9DN cells. As shown in Fig. 6C, Noxa induction by F6 and G5 was efficiently inhibited by the specific stealth RNAi oligos but not by the control oligos. Down-regulation of Noxa has a minimal effect on the accumulation of the putative Smac precursor (Fig. 6C). When apoptosis was scored by trypan blue exclusion and caspase activity, it was evident that down-regulation of Noxa inhibited but did not completely suppress apoptosis and caspase activation in response to F6, G5, or MG-132 (Fig. 6D-E). These results indicate that Noxa plays an important role in the apoptotic pathway activated by F6 and G5 and that alternative mechanisms are operating in the absence of Noxa.

Mitochondrial fragmentation is an early response to proteasome alterations and depends on induction of Noxa. A delicate balance of fission and fusion normally controls the morphology of mitochondria. During apoptosis, mitochondria undergo morphologic changes resulting in the generation of small round organelles (25). This process has been described as mitochondrial fragmentation. We noted that in F6- and G5-treated cells, mitochondria presented altered morphologies; hence, we decided to explore in detail the effect of these compounds on mitochondrial morphology. Because there is a temporal relationship between mitochondrial fragmentation and the release of cytochrome c (26), we scored mitochondrial fragmentation in relation to mitochondrial outer membrane permeabilization (MOMP). The release of Smac into the cytosol was used as an indicator of MOMP. Double immunofluorescence experiments were done to visualize mitochondrial morphology (MitoTracker labeling) and Smac subcellular localization. As illustrated in Fig. 7A , three phenotypes can be distinguished in cells treated with apoptotic stressors in terms of mitochondrial fragmentation and MOMP: cells that present an extended mitochondrial tubular network, cells with fragmented mitochondria but with intact outer mitochondrial membrane, and, finally, cells with fragmented mitochondria and MOMP.


Figure 7
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  		Figure 7. Mitochondrial fission in response to the inhibition of the ubiquitin-proteasome system. A, immunofluorescence analysis to visualize mitochondrial morphology and Smac localization. Mitochondria were evidenced with MitoTracker as described in Materials and Methods. B, E1A and E1A/C9DN cells treated or not for 12 hours with the indicated compounds were subjected to immunofluorescence analysis to visualize mitochondria and Smac. Appearance of mitochondrial fission and MOMP was scored as described in (A). Columns, mean (n = 3); bars, SD. C, E1A cells transfected with Noxa RNAi oligos or with control (contr) oligos were incubated for 12 hours with the indicated compounds or maintained untreated. Mitochondrial fragmentation was scored in relation to Smac localization by immunofluorescence as described in (A). Columns, mean (n = 3); bars, SD. Concentrations used were as follows: F6 (10 µmol/L), G5 (1.25 µmol/L), MG-132 (5 µmol/L), and etoposide (50 µmol/L).

 
The different apoptotic inducers were characterized by their ability to induce mitochondrial fragmentation (Fig. 7B). F6, G5, and MG-132 provoke mitochondrial fragmentation in a higher percentage of cells with an intact outer mitochondrial membrane compared with etoposide. This suggests that proteasome inhibitors can alter mitochondrial morphology before MOMP. Caspase-9 mutation does not affect mitochondrial fragmentation but instead augmented the percentage of cells showing MOMP as a consequence of the delayed cellular dismantling.

Under certain conditions, BH3-only proteins can regulate mitochondrial fission (27). Hence, we decided to investigate whether Noxa stabilization plays any role in the early mitochondrial fission induced by F6 and G5.

Noxa induction was suppressed by the specific small interfering RNA, and cells were analyzed for mitochondrial fragmentation and MOMP as described above. Figure 7C illustrates that Noxa induction plays an important role in mitochondrial fragmentation in response to alterations of the ubiquitin-proteasome system. Nevertheless, because Noxa silencing reduces but does not completely suppress mitochondrial fragmentation, alternative mechanisms may exist that synergize with Noxa.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this report, we have characterized the apoptotic pathway activated by two compounds, 4-piperidione,3,5-bis[(4-methylphenyl)methylene]-hydrochloride and 4H-thiopyran-4-one, tetrahydro-3,5-bis[(4-nitrophenyl)methylene]-,1,1-dioxide, which belong to the challenge set of the DTP-NCI. For simplicity, these compounds have been named F6 and G5 throughout the work.

F6 and G5 have been identified for their ability to initiate a Bcl-2-dependent but apoptosome-independent pathway of caspase activation. This pathway seems to be extremely peculiar given that (a) it was not induced by the other 55 compounds that characterize the challenge set and (b) a large variety of anticancer molecules failed to activate this pathway (9). A similar Bcl-2-dependent but apoptosome-independent pathway of caspase activation is stimulated by proteasome inhibitors (9). Hence, it should not be surprising that also F6 and G5 act as inhibitors of the ubiquitin-proteasome system. As a matter of fact, the presence of a pharmacophore that confers inhibitory activity toward ubiquitin isopeptidases (12, 14–16) and experimental data here reported has confirmed that these new compounds can impede ubiquitin-dependent protein degradation.

Isopeptidases belong to a heterogeneous family of deubiquitinating enzymes (DUB), which can revert the conjugation of ubiquitin to specific substrates and can process inactive ubiquitin. DUBs have been divided into five distinct subfamilies based on structural similarities and mechanisms of action (28, 29). These subfamilies include a large group of enzymes of which the vast majority are cysteine proteases; only one class includes metalloenzymes. The pharmacophore found in F6 and G5 should inhibit cysteine proteases (30); however, which among the different DUBs are the real targets is still unknown. Interestingly, the prostaglandin {Delta}12-PGJ2 holds the same pharmacophore of F6 and G5 and similarly induces apoptosis via accumulation of polyubiquitinated proteins (12, 14); {Delta}12-PGJ2 has been recently suggested to inhibit UCH-L1 and UCH-L3 at micromolar concentrations (15). It is clear that further studies are required to define in detail which DUBs can be inhibited by F6 and G5.

More in general, our studies suggest that inhibition of the ubiquitin-proteasome system induces apoptosis by a variety of different mechanisms, including up-regulation of the BH3-only protein Noxa, stabilization of the IAP antagonist Smac, and activation of the death receptor pathway. We have noted and confirmed that the levels of Noxa and Smac are modulated by F6, G5, and proteasome inhibitors. The effect on Smac is evident only on the cytosolic fraction where these compounds could counteract the E3 ligase activity of the IAPs. Noxa levels are dramatically induced in response to F6 and G5. The promoter of Noxa is under the control of p53, which can be activated by proteasome inhibitors (31, 32). Nevertheless, in cancer cells, up-regulation of Noxa transcription in response to bortezomib can take place in p53-dependent and p53-independent manners (23, 24, 33).

RNAi experiments have shown that Noxa induction is important for apoptosis in response to F6 and G5; however, it should be noted that additional proapoptotic proteins are likely to be involved because only a partial protection was guaranteed by the Noxa RNAi. A possible candidate is NBK/Bik, another BH3-only protein under the proteasome control (34, 35).

Mitochondrial morphology is maintained by cycles of fission and fusion through the activities of dynamin-like proteins. Optic atrophy 1, mitofusin 1, and mitofusin 2 modulate fusion, whereas Drp1 and FIS1, the "Drp1 receptor" on the outer mitochondrial membrane, control fission (25, 26). Reduced fusion and improved fission can contribute to mitochondrial fragmentation. Studies aimed to deciphering the timing of mitochondrial fragmentation during apoptosis have shown that it coincides with the activation of Bax and Bak as proved by their colocalization in foci on the outer mitochondrial membrane and hence is concurrent with MOMP and cytochrome c release (36, 37).

A further characteristic feature of the apoptotic response triggered by F6 and G5 is the appearance of mitochondrial fragmentation before MOMP, measured as the release of Smac into the cytosol. Although Noxa induction has a role in this early fragmentation, the effect of its silencing is partial. Again, this should be expected because it is not the solely BH3-only protein capable of regulating mitochondrial fission. Recently, it has been shown that also NBK/Bik can modulate mitochondrial fusion before MOMP through endoplasmic reticulum–mediated Ca2+ release and Drp1 activation (27, 38). Therefore, Noxa and NBK/Bik could synergize under the influence of the degradative block induced by F6 and G5.

Although Noxa plays an important role in the apoptotic response to proteasome inhibitors, it is unclear if it can confer apoptosome independence. An unexpected contribution to this independence arises from the role of the death receptor pathway. By expressing CrmA, FADD/DN, and c-FLIPS in E1A/C9DN cells, we have noted that the death receptor pathway can compensate for the absence of a functional apoptosome when apoptosis is induced by inhibitors of the ubiquitin-proteasome system.

How could the inhibitors of the ubiquitin-proteasome system activate the death receptor pathway? Many evidences indicate that elements of the death receptor pathway are under the control of the proteasome. Up-regulation of DR4/TNF-related apoptosis-inducing ligand (TRAIL) receptor (TRAIL-R) 1, DR5/TRAIL-R2, and DcR2/TRAIL-R4 on the cell surface of hepatocarcinoma cells was observed after incubation with proteasome inhibitors (39). Moreover, DR5 is regulated by proteasome inhibitors at multiple levels (40), including ubiquitination (41) and increased transcription (42). Proteasome inhibitors can also reduce the levels c-FLIPS and hence synergize with TRAIL to promote apoptosis (43). It is important to remember that also Smac, which is preserved at higher levels in the cytosol of F6- and G5-treated cells, can synergize with the death receptor pathway (44–49). Moreover, artificial expression of cytosolic Smac can bypass the protection afforded by Bcl-2 and potentiate death receptor–induced apoptosis by antagonizing X-linked IAP–mediated inhibition of active caspase-3 and caspase-7 (44, 49). These authors, similarly to us, hypothesize the existence of a Bcl-2-dependent but apoptosome-independent pathway of caspase activation, which can be activated by the enrollment of the death receptors in concert with cytosolic Smac (49).

Based on these evidences, we propose the model illustrated in Fig. 8 . F6, G5, and proteasome inhibitors can induce Noxa expression, which plays a part in the release of Smac into the cytosol. The block operated by these compounds on the E3-IAP activities allows the accumulation of higher levels of released cytosolic Smac, which allows in turn to free the effector caspases from the IAP inhibition. Within this scenario, activation of the extrinsic pathway, possibly by modulating the expression and the localization of its various components, can induce activation of caspase-3 and caspase-7 no longer restrained by the IAPs in the absence of the apoptosome.


Figure 8
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  		Figure 8. Model for the mechanism of apoptosis induced by F6 and G5. Inhibition of the ubiquitin-proteasome system by F6 and G5 results in the accumulation of Noxa, which contributes to MOMP and Smac release. In the presence of F6 and G5, Smac degradation is impaired and therefore high levels of it accumulate in the cytosol. F6 and G5, in a still undefined manner, could activate the extrinsic pathway, which, with the help of Smac, keeping in check the IAPs, can turn on the effector caspases in the absence of the apoptosome.

 
In conclusion, three considerations can be drawn from these data. First, the inhibition of the ubiquitin-proteasome pathway elicits a rather unique apoptotic response as documented by the apoptosome-independent caspase activation. Second, that not only the proteolytic chamber but also the DUBs are promising targets for new anticancer treatments with a selected mechanism of cytotoxicity (50). Third, screening of chemical libraries on cancer cells containing defined mutations in elements of the apoptotic program can lead to identification of compounds that are useful to characterize alternative pathways of caspase activation and that can be further developed for clinical applications.


   Acknowledgments
 
Grant support: Associazione Italiana Ricerca sul Cancro, Ministero dell'Istruzione, Università e Ricerca, and Consorzio Interuniversitario Biotecnologie.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank G. Lippe (Dipartimento di Scienze e Tecnologie Biomediche, Università degli Studi di Udine, Udine, Italy), R. Maestro (MMNP-Unit Experimental Oncology CRO-IRCCS, National Cancer Institute, Aviano, Italy), and Y. Lazebnik (Cold Spring Harbor Laboratory, Cold Spring Harbor, NY) for anti-ATP synthase and anti-Apaf-1 antibodies and cell lines and the DTP of the NCI for the challenge set.


   Footnotes
 
Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

1 http://dtp.nci.nih.gov/. Back

Received 2/21/06. Revised 6/14/06. Accepted 7/17/06.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Johnstone RW, Ruefli AA, Lowe SW. Apoptosis: a link between cancer genetics and chemotherapy. Cell 2002;108:153–64.[CrossRef][Medline]
  2. Igney FH, Krammer PH. Death and anti-death: tumour resistance to apoptosis. Nat Rev Cancer 2002;2:277–88.[CrossRef][Medline]
  3. Fischer U, Janicke RU, Schulze-Osthoff K. Many cuts to ruin: a comprehensive update of caspase substrates. Cell Death Differ 2003;10:76–100.[CrossRef][Medline]
  4. Boatright KM, Salvesen GS. Mechanisms of caspase activation. Curr Opin Cell Biol 2003;15:725–31.[CrossRef][Medline]
  5. Jiang X, Wang X. Cytochrome c-mediated apoptosis. Annu Rev Biochem 2004;73:87–106.[CrossRef][Medline]
  6. Lavrik I, Golks A, Krammer PH. Death receptor signaling. J Cell Sci 2005;2:265–7.
  7. Debatin KM, Poncet D, Kroemer G. Chemotherapy: targeting the mitochondrial cell death pathway. Oncogene 2002;21:8786–803.[CrossRef][Medline]
  8. Fearnhead HO, Rodriguez J, Govek EE, et al. Oncogene-dependent apoptosis is mediated by caspase-9. Proc Natl Acad Sci U S A 1998;95:13664–9.[Abstract/Free Full Text]
  9. Henderson CJ, Aleo E, Fontanini A, Maestro R, Paroni G, Brancolini C. Caspase activation and apoptosis in response to proteasome inhibitors. Cell Death Differ 2005;12:1240–54.[CrossRef][Medline]
  10. Henderson C, Mizzau M, Paroni G, Maestro R, Schneider C, Brancolini C. Role of caspases, Bid, and p53 in the apoptotic response triggered by histone deacetylase inhibitors trichostatin-A (TSA) and suberoylanilide hydroxamic acid (SAHA). J Biol Chem 2003;278:12579–89.[Abstract/Free Full Text]
  11. Paroni G, Mizzau M, Henderson C, Del Sal G, Schneider C, Brancolini C. Caspase-dependent regulation of histone deacetylase 4 nuclear-cytoplasmic shuttling promotes apoptosis. Mol Biol Cell 2004;15:2804–18.[Abstract/Free Full Text]
  12. Mullally JE, Fitzpatrick FA. Pharmacophore model for novel inhibitors of ubiquitin isopeptidases that induce p53-independent cell death. Mol Pharmacol 2002;62:351–8.[Abstract/Free Full Text]
  13. Wei MC, Zong WX, Cheng EH, et al. Proapoptotic BAX and BAK: a requisite gateway to mitochondrial dysfunction and death. Science 2001;292:727–30.[Abstract/Free Full Text]
  14. Mullally JE, Moos PJ, Edes K, Fitzpatrick FA. Cyclopentenone prostaglandins of the J series inhibit the ubiquitin isopeptidase activity of the proteasome pathway. J Biol Chem 2001;276:30366–73.[Abstract/Free Full Text]
  15. Zongmin L, Melandri F, Berdo I, et al. {Delta}12-Prostaglandin J2 inhibits the ubiquitin hydrolase UCH-L1 and elicits ubiquitin-protein aggregation without proteasome inhibition. Biochem Biophys Res Commun 2004;319:1171–80.[CrossRef][Medline]
  16. Verbitski SM, Mullally JE, Fitzpatrick FA, Ireland CM. Punaglandins, chlorinated prostaglandins, function as potent Michael receptors to inhibit ubiquitin isopeptidase activity. J Med Chem 2004;47:2062–70.[CrossRef][Medline]
  17. Hershko A, Ciechanover A. The ubiquitin system. Annu Rev Biochem 1998;67:425–79.[CrossRef][Medline]
  18. Kisselev AF, Goldberg AL. Proteasome inhibitors: from research tools to drugs candidates. Chem Biol 2001;8:739–58.[CrossRef][Medline]
  19. Demarchi F, Brancolini C. Altering protein turnover in tumor cells: new opportunities for anti-cancer therapies. Drug Resist Updat 2005;8:359–68.[CrossRef][Medline]
  20. Vaux DL, Silke J. IAPs—the ubiquitin connection. Cell Death Differ 2005;12:1205–7.[CrossRef][Medline]
  21. Wright G, Terada K, Yano M, Sergeev I, Mori M. Oxidative stress inhibits the mitochondrial import of preproteins and leads to their degradation. Exp Cell Res 2001;263:107–17.[CrossRef][Medline]
  22. Tan TT, Degenhardt K, Nelson DA, et al. Key roles of BIM-driven apoptosis in epithelial tumors and rational chemotherapy. Cancer Cell 2005;7:227–38.[CrossRef][Medline]
  23. Fernandez Y, Verhaegen M, Miller TP, et al. Differential regulation of Noxa in normal melanocytes and melanoma cells by proteasome inhibition: therapeutic implications. Cancer Res 2005;65:6294–300.[Abstract/Free Full Text]
  24. Qin JZ, Ziffra J, Stennett L, et al. Proteasome inhibitors trigger NOXA-mediated apoptosis in melanoma and myeloma cells. Cancer Res 2005;65:6282–93.[Abstract/Free Full Text]
  25. Perfettini JL, Roumier T, Kroemer G. Mitochondrial fusion and fission in the control of apoptosis. Trends Cell Biol 2005;15:179–83.[CrossRef][Medline]
  26. Youle RJ, Karbowski M. Mitochondrial fission in apoptosis. Nat Rev Mol Cell Biol 2005;6:657–63.[Medline]
  27. Mathai JP, Germain M, Shore GC. BH3-only BIK regulates BAX,BAK-dependent release of Ca2+ from endoplasmic reticulum stores and mitochondrial apoptosis during stress-induced cell death. J Biol Chem 2005;280:23829–36.[Abstract/Free Full Text]
  28. Amerik AY, Hochstrasser M. Mechanism and function of deubiquitinating enzymes. Biochim Biophys Acta 2004;1695:189–207.[Medline]
  29. Nijman SM, Luna-Vargas MP, Velds A, et al. A genomic and functional inventory of deubiquitinating enzymes. Cell 2005;123:773–86.[CrossRef][Medline]
  30. Straus DS, Glass CK. Cyclopentenone prostaglandins: new insights on biological activities and cellular targets. Med Res Rev 2001;21:185–210.[CrossRef][Medline]
  31. Oda E, Ohki R, Murasawa H, et al. Noxa, a BH3-only member of the Bcl-2 family and candidate mediator of p53-induced apoptosis. Science 2000;288:1053–8.[Abstract/Free Full Text]
  32. Michael D, Oren M. The p53-Mdm2 module and the ubiquitin system. Semin Cancer Biol 2003;13:49–58.[CrossRef][Medline]
  33. Perez-Galan P, Roue G, Villamor N, Montserrat E, Campo E, Colomer D. The proteasome inhibitor bortezomib induces apoptosis in mantle-cell lymphoma through generation of ROS and Noxa activation independent of p53 status. Blood 2006;107:257–64.[Abstract/Free Full Text]
  34. Nikrad M, Johnson T, Puthalalath H, Coultas L, Adams J, Kraft AS. The proteasome inhibitor bortezomib sensitizes cells to killing by death receptor ligand TRAIL via BH3-only proteins Bik and Bim. Mol Cancer Ther 2005;4:443–9.[Abstract/Free Full Text]
  35. Zhu H, Zhang L, Dong F, et al. Bik/NBK accumulation correlates with apoptosis-induction by bortezomib (PS-341, Velcade) and other proteasome inhibitors. Oncogene 2005;24:4993–9.[CrossRef][Medline]
  36. Karbowski M, Lee YJ, Gaume B, et al. Spatial and temporal association of Bax with mitochondrial fission sites, Drp1, and Mfn2 during apoptosis. J Cell Biol 2002;159:931–8.[Abstract/Free Full Text]
  37. Karbowski M, Arnoult D, Chen H, Chan DC, Smith CL, Youle RJ. Quantitation of mitochondrial dynamics by photolabeling of individual organelles shows that mitochondrial fusion is blocked during the Bax activation phase of apoptosis. J Cell Biol 2004;164:493–9.[Abstract/Free Full Text]
  38. Germain M, Mathai JP, McBride HM, Shore GC. Endoplasmic reticulum BIK initiates DRP1-regulated remodelling of mitochondrial cristae during apoptosis. EMBO J 2005;24:1546–56.[CrossRef][Medline]
  39. Ganten TM, Koschny R, Haas TL, et al. Proteasome inhibition sensitizes hepatocellular carcinoma cells, but not human hepatocytes, to TRAIL. Hepatology 2005;42:588–97.[CrossRef][Medline]
  40. He Q, Huang Y, Sheikh MS. Proteasome inhibitor MG132 upregulates death receptor 5 and cooperates with Apo2L/TRAIL to induce apoptosis in Bax-proficient and -deficient cells. Oncogene 2004;23:2554–8.[CrossRef][Medline]
  41. Johnson TR, Stone K, Nikrad M, et al. The proteasome inhibitor PS-341 overcomes TRAIL resistance in Bax and caspase 9-negative or Bcl-xL overexpressing cells. Oncogene 2003;22:4953–63.[CrossRef][Medline]
  42. Yoshida T, Shiraishi T, Nakata S, et al. Proteasome inhibitor MG132 induces death receptor 5 through CCAAT/enhancer-binding protein homologous protein. Cancer Res 2005;65:5662–7.[Abstract/Free Full Text]
  43. Sayers TJ, Brooks AD, Koh CY, et al. The proteasome inhibitor PS-341 sensitizes neoplastic cells to TRAIL-mediated apoptosis by reducing levels of c-FLIP. Blood 2003;102:303–10.[Abstract/Free Full Text]
  44. Srinivasula SM, Datta P, Fan XJ, Fernandes-Alnemri T, Huang Z, Alnemri ES. Molecular determinants of the caspase-promoting activity of Smac/DIABLO and its role in the death receptor pathway. J Biol Chem 2000;75:36152–7.
  45. Zhang XD, Zhang XY, Gray CP, Nguyen T, Hersey P. Tumor necrosis factor-related apoptosis-inducing ligand-induced apoptosis of human melanoma is regulated by Smac/DIABLO release from mitochondria. Cancer Res 2001;61:7339–48.[Abstract/Free Full Text]
  46. Guo F, Nimmanapalli R, Paranawithana S, et al. Ectopic overexpression of second mitochondria-derived activator of caspases (Smac/DIABLO) or cotreatment with N-terminus of Smac/DIABLO peptide potentiates epothilone B derivative-(BMS 247550) and Apo-2L/TRAIL-induced apoptosis. Blood 2002;99:3419–26.[Abstract/Free Full Text]
  47. Fulda S, Wick W, Weller M, Debatin KM. Smac agonists sensitize for Apo2L/TRAIL- or anticancer drug-induced apoptosis and induce regression of malignant glioma in vivo. Nat Med 2002;8:808–15.[Medline]
  48. Deng Y, Lin Y, Wu X. TRAIL-induced apoptosis requires Bax-dependent mitochondrial release of Smac/DIABLO. Genes Dev 2002;16:33–45.[Abstract/Free Full Text]
  49. Sun XM, Bratton SB, Butterworth M, MacFarlane M, Cohen GM. Bcl-2 and Bcl-xL inhibit CD95-mediated apoptosis by preventing mitochondrial release of Smac/DIABLO and subsequent inactivation of X-linked inhibitor-of-apoptosis protein. J Biol Chem 2002;277:11345–51.[Abstract/Free Full Text]
  50. Adams J, Palombella VJ, Sausville EA, et al. Proteasome inhibitors: a novel class of potent and effective antitumor agents. Cancer Res 1999;59:2615–22.[Abstract/Free Full Text]



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