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Carcinogenesis Advance Access originally published online on January 3, 2008
Carcinogenesis 2008 29(3):544-551; doi:10.1093/carcin/bgm294
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© The Author 2008. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Activation of the JNK pathway promotes phosphorylation and degradation of BimEL—a novel mechanism of chemoresistance in T-cell acute lymphoblastic leukemia

Kam Tong Leung, Karen Kwai-Har Li1, Samuel Sai-Ming Sun, Paul Kay Sheung Chan2, Vincent Eng-Choon Ooi and Lawrence Chi-Ming Chiu*

Department of Biology, The Chinese University of Hong Kong, Hong Kong SAR, China
1 Department of Pediatrics
2 Department of Microbiology, Prince of Wales Hospital, Hong Kong SAR, China

* To whom correspondence should be addressed. Tel: +852-3163-4466; Fax: +852-2603-5745; Email: chimingchiu{at}graduate.hku.hk


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 Funding
 References
 
T-cell acute lymphoblastic leukemias (T-ALLs) are highly malignant tumors with 20% of patients continues to fail therapy, in part due to chemoresistance of T-ALL cells via largely unknown mechanisms. Here, we showed that lack of Bcl-2-interacting mediator of cell death (Bim)EL protein expression, a BH3-only member of the Bcl-2 family proteins, conferred resistance of a T-ALL cell line, Sup-T1, to etoposide-induced apoptosis. Overexpression of BimEL significantly restored its sensitivity to etoposide-induced caspase activation and poly(ADP-ribose) polymerase cleavage. Surprisingly, we found that constitutive activation of the c-Jun N-terminal kinase (JNK) pathway in Sup-T1 cells promoted phosphorylation and degradation of BimEL via the proteosome. Blocking with a proteosome inhibitor yielded an elevated level of BimEL and accumulation of BimEL species phosphorylated at Ser69. Pretreatment of Sup-T1 cells with a specific JNK inhibitor, SP600125, also increased the BimEL level and resensitized the cells to etoposide-induced apoptosis. Together, our findings suggest that the JNK activation status may correlate with the BimEL level and in turn can control the sensitivity of T-ALL cells to chemotherapeutic agents.

Abbreviations: Bim, Bcl-2-interacting mediator of cell death; DEVD-AFC, N-acetyl-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin; ERK, extracellular signal-regulated kinase; JNK, c-Jun N-terminal kinase; LEHD-AFC, N-acetyl-Leu-Glu-His-Asp-7-amino-4-trifluoromethyl coumarin; PARP, poly(ADP-ribose) polymerase; PCR, polymerase chain reaction; T-ALL, T-cell acute lymphoblastic leukemia; z-VAD-fmk, benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone


   Introduction
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 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 Funding
 References
 
Despite the adverse risk associated with T-cell acute lymphoblastic leukemia (T-ALL) has progressively been overwhelmed by intensive chemotherapeutic regimens, ~20% of patients with T-ALL continues to fail therapy (1). Higher rates of induction failure and early relapse in these patients suggest that their blast cells are more inherently resistant to conventional chemotherapeutic agents. Unfortunately, the precise mechanism of chemoresistance in T-ALL patients is still largely unknown. Thus, identification of additional prognostic factors and chemoresistance mechanisms that can be used to tailor therapy more precisely remains a top priority.

Most chemotherapeutic drugs activate the intrinsic apoptosis pathway, which is tightly regulated by proteins of the Bcl-2 family. Abnormal expression of these proteins has been implicated in the development of chemoresistance in tumor cells (2,3). The Bcl-2-interacting mediator of cell death (Bim) is a BH3-only proapoptotic Bcl-2 family protein with three major isoforms generated by alternative splicing: BimEL, BimL and BimS (4). The involvement of Bim in apoptosis was first implicated in a physiological study using gene-knockout mice (5). Bim-deficient mice had an elevated level of lymphoid and myeloid cells. Moreover, Bim–/– lymphocytes were resistant to apoptotic stimuli such as cytokine deprivation, calcium ion flux and microtubule perturbation (5). Other than cells in the hematological system, Bim is also required for apoptosis in neurons (6) and osteoclasts (7). Recent studies have reported the roles of Bim in tumorigenesis and response to chemotherapeutic drugs. The loss of a single allele of Bim induced the development of mouse B-cell leukemia in Myc-transgenic mice (8), and homozygous deletion of Bim was found in 17% of the patients with mantle cell lymphoma (9). For the responsiveness of tumor cells to chemotherapeutic drugs, Bim is required for apoptosis induced by inhibitors of histone deacetylase and paclitaxel (10,11). Recently, Bim was also found essential to the glucocorticoids-induced apoptosis in T-ALL cells (12).

The expression and proapoptotic activity of Bim are subjected to transcriptional and posttranslational regulation (13,14). Transcriptional regulation of Bim is cell-type dependent. The growth factor withdrawal-induced Bim expression requires activation of the c-Jun N-terminal kinase (JNK) pathway in neurons (6), but involves activation of the Akt pathway in hematopoietic cells (15). In fibroblasts and breast epithelial cells, Bim expression is depended on the inactivation of the extracellular signal-regulated kinase (ERK) pathway (16,17). More recent studies have demonstrated that Bim can be regulated posttranslationally by phosphorylation, and the kinases responsible for Bim phosphorylation involve ERK, JNK, p38 and Akt (18–23).

JNK and its associated signaling pathways have been implicated in stress-induced apoptosis (24). Previous data suggest that JNK functions as a proapoptotic kinase. For example, jnk-null fibroblasts were shown to be defective in cytochrome c-driven apoptosis (25). Similarly, immature thymocytes from jnk2-null mice were resistant to anti-CD3 antibody-mediated apoptosis (26). However, evidence has also been reported regarding to the antiapoptotic function of JNK. In mice lacking both jnk1 and jnk2, increased apoptosis and caspase activation were found in their forebrain (27). Moreover, skin tumorigenesis was suppressed in jnk2–/– mice (28). Although substantial evidence supports a functional role for the JNK-signaling pathway in cell survival, the molecular mechanism is incompletely understood. In this report, we investigated the chemoresistance mechanisms in a T-ALL cell line, Sup-T1. We demonstrate that the expression level of Bim is an important determinant for the sensitivity of T-ALL cells to drug-induced apoptosis. We also show that BimEL is phosphorylated and targeted for proteosomal degradation when the JNK pathway is activated, thereby rendering Sup-T1 cells resistant to drug-induced apoptosis. Thus, our findings define a novel mechanism of chemoresistance in T-ALL cells and provide new insights into the regulation of cellular Bim level and the mechanisms of JNK-mediated cell survival.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 Funding
 References
 
Cell culture
The human T-ALL cell lines, HSB-2, MOLT-3 and Sup-T1, as well as the Burkitt lymphoma-derived cell line, Raji, were purchased from the American Type Culture Collection (Rockville, MD). All cell lines were grown in RPMI-1640 (Sigma, St Louis, MO) supplemented with 10% heat-inactivated fetal bovine serum (Gibco, Rockville, MD), 1.5 g/l sodium bicarbonate, 1 mM sodium pyruvate, 10 mM N-(2-hydroxyethyl)piperazine-N'-(2-ethanesulfonic acid), 100 U/ml penicillin and 100 µg/ml streptomycin. The cells were maintained in 5% CO2 at 37oC. Where indicated, Sup-T1 cells were incubated with different inhibitors including MG-132 (50 µM), U0126 (10 µM), PD98509 (10 µM), SP600125 (10 µM), SB203580 (15 µM), LY294002 (50 µM) or wortmannin (100 nM) for the specified time points. All inhibitors were obtained from Calbiochem, La Jolla, CA.

Induction and quantification of apoptosis
To induce apoptosis, cells were treated with or without 1–10 µM etoposide (Sigma) for the indicated time points. In some experiments, cells were preincubated with 50 µM benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (z-VAD-fmk) for 1 h prior to etoposide treatment. After treatment, cells were washed with phosphate-buffered saline and fixed in 70% ethanol at –20°C for 24 h. The cells were next incubated in dark for 30 min with 1 mg/ml ribonuclease A and 10 µg/ml propidium iodide (Sigma). Samples were analyzed for DNA content by measuring propidium iodide fluorescence on the Epics XL-MCL flow cytometer (Beckman Coulter, Miami, FL). Cells displaying hypodiploid DNA content were quantified and regarded as the apoptotic population.

Determination of caspase activities
Caspase-3 and caspase-9 activities were determined by using the fluorogenic peptide substrates, N-acetyl-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin (DEVD-AFC) (50 µM) and N-acetyl-Leu-Glu-His-Asp-7-amino-4-trifluoromethyl coumarin (LEHD-AFC) (100 µM) (Calbiochem), respectively. Cell lysates and substrates were mixed in a standard reaction buffer (20 mM N-(2-hydroxyethyl)piperazine-N'-(2-ethanesulfonic acid), 10% glycerol, 2 mM dithiothreitol at pH 7.5) and incubated for 1 h at 37°C. The amount of enzyme-catalyzed AFC release was measured by a fluorescence plate reader (Tecan, Grödig, Austria) with excitation/emission wavelengths of 400/505 nm. Fluorescence values were converted into picomoles of AFC release by using a standard curve generated with free AFC.

Western blotting
For preparation of whole-cell lysates, cells were harvested and suspended for 30 min on ice in an extraction buffer (20 mM Tris–HCl at pH 8.0, 150 mM NaCl, 5 mM ethylenediaminetetraacetic acid, 0.2% bovine serum albumin and 1% Triton X-100) supplemented with protease and phosphatase inhibitors (BD Biosciences, San Jose, CA). Lysates were clarified by centrifugation for 15 min at 15 000g. Aliquots of the supernatants were used for protein concentration determination by bicinchoninic acid assay (Pierce, Rockford, IL). To obtain mitochondrial and cytosolic fractions, cell homogenates were fractionated by the Cytosol–Mitochondria Fractionation Kit (Calbiochem) according to the manufacturer’s instructions. Samples (30–100 µg protein) were denatured in Laemmli buffer and resolved on 10 or 13% sodium docecyl sulfate–polyacrylamide gel electrophoresis minigels, followed by electrophoretic transfer to nitrocellulose membranes (Hybond ECL; Amersham, Buckinghamshire, UK). The membranes were blocked for 1 h in Tris-buffered saline with 0.1% Tween-20 containing 5% non-fat dry milk. Immunodetection was performed by overnight incubation at 4°C with primary antibodies diluted in the same blocking buffer. Primary antibodies to poly(ADP-ribose) polymerase (PARP), caspase-3, caspase-9, Bcl-xL, Bak, Bad, Bim, Bid, phospho-ERK (Thr-202/Tyr-204), ERK, phospho-JNK (Thr-183/Tyr-185), JNK, phospho-p38 (Thr-180/Tyr-182), p38, phospho-Akt (Ser-473), Akt, phospho-c-Jun (Ser-63), c-Jun and cytochrome c were obtained from Cell Signaling Technology, Danvers, MA. Antibodies to Bcl-2, Mcl-1, Bax, Bik and Nip-3 were purchased from BD Biosciences. Antibodies to phospho-BimEL (Ser-55 and Ser-69) were obtained from Upstate Biotechnology (Lake Placid, NY), and β-actin antibody was from Sigma. After extensive washing, the blots were probed with horseradish peroxidase-conjugated secondary antibodies (Cell Signaling Technology) for 1 h at room temperature, and proteins were visualized by enhanced chemiluminescence (Cell Signaling Technology). Primary antibodies were used at 1:250 to 1:5000 dilutions and secondary antibodies were used at a 1:2000 dilution.

Cell-free apoptosis reactions
For preparation of S-100 cytosolic extracts, 5 x 107 cells were collected and incubated for 10 min in ice-cold S-100 buffer (20 mM N-(2-hydroxyethyl)piperazine-N'-(2-ethanesulfonic acid)–KOH at pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM ethylenediaminetetraacetic acid, 1 mM ethyleneglycol-bis(aminoethylether)-tetraacetic acid and 1 mM dithiothreitol) supplemented with protease inhibitors. Cells were disrupted using a Dounce homogenizer and centrifuged at 1000g for 10 min at 4°C. Supernatants were further centrifuged at 100 000g for 1 h, and the resulting supernatants (S-100 fraction) were stored at –70°C or used immediately. For cell-free caspase activation, S-100 cytosol (100 µg) was incubated with 1 mM deoxyadenosine triphosphate and 4 µM cytochrome c (Sigma) for 1 h at 30°C. Aliquots of reaction products were then collected for assessment of procaspase-3 processing by western blotting.

Analysis of mitochondrial membrane potential
Loss of mitochondrial membrane potential ({Delta}{Psi}m) was detected by using the potential-sensitive fluorescent probe, tetramethylrhodamine ethyl ester (Molecular Probes, Eugene, OR). Cells were incubated with 25 nM tetramethylrhodamine ethyl ester for 15 min at 37°C prior to analysis on the flow cytometer. As a positive control, cells were incubated with 100 µM potential-disrupting agent, carbonyl cyanide 3-chlorophenylhydrazone (Sigma) for 24 h. Data are displayed as the percentage of cells with reduction in mitochondrial membrane potential.

Transient transfection of Sup-T1 cells
The Bcl-2 antisense oligonucleotide, which targets the first six codons of the opening reading frame of Bcl-2 messenger RNA, was purchased from Calbiochem. The full-length, wild-type human BimEL and Bax cDNA, which are cloned into the expression vector pCMV6-XL5, were obtained from Origene, Rockville, MD. Transient transfection of Sup-T1 cells was accomplished using Lipofectin reagent (Invitrogen, Carlsbad, CA) under serum-free conditions. Briefly, 2.5 x 106 cells were transfected with either 3 µg of Bcl-2 antisense oligonucleotide or 5 µg of BimEL or Bax expression vectors. After 24 h of transfection, the medium was replaced with complete medium, and the cells were treated with or without etoposide for 24 h. Cells were then harvested for analysis of gene expression and processing of procaspase-3 and PARP by western blotting.

Reverse transcription–polymerase chain reaction
Total RNA was isolated by TRIzol reagent (Invitrogen). First-strand cDNA was synthesized by reverse transcription using the SuperScript First-Strand Synthesis System (Invitrogen) according to the manufacturer’s instructions. The first-strand cDNA was diluted and used as template in polymerase chain reactions (PCRs) containing 0.8 µM primers, 0.2 mM deoxyribonucleotide triphosphates, 1.5 mM MgCl2, 1x PCR buffer and 0.5 U Taq polymerase. All PCR reagents were purchased from Amersham. The PCR was subjected to 3 min denaturation at 96°C, followed by 25 cycles of amplification (94°C, 30 s; 55°C, 30 s and 72°C, 60 s) and a final extension at 72°C for 10 min. The gene-specific primer pairs were as follows: Bax sense (5'-CTGAGCAGATCATGAAGACAGG-3') and Bax antisense (5'-AAGTAGAAAAGGGCGACAACC-3'); Bcl-2 sense (5'-GAATTCCACTGTCAAGAAAGAGCAGT-3') and Bcl-2 antisense (5'-GCTTTCGAAATATCAACCACAGCATT-3'); Bim sense (5'-TGATGTAAGTTCTGAGTGTG-3') and Bim antisense (5'-CGCATATCTGCAGGTTCAGCC-3') and glyceraldehyde 3-phosphate dehydrogenase sense (GAPDH) (5'-AAGATCATCAGCAATGCCTCC-3') and glyceraldehyde 3-phosphate dehydrogenase antisense (5'-CCTGCTTCACCACCTTCTTGA-3'). PCR products were resolved by electrophoresis in 1% agarose gels and were visualized by ethidium bromide staining.

Alkaline phosphatase digestion of Bim
Cell lysates (50 µg) were digested with 50 U of calf intestinal alkaline phosphatase (Calbiochem) for 1 h at 37°C in a digestion buffer (50 mM Tris–HCl at pH 7.9, 100 mM NaCl, 10 mM MgCl2 and 1 mM dithiothreitol) prior to western blotting.

Statistical analysis
All values are reported as mean ± SD. Results were analyzed with Student’s t-test. P values were considered statistically significant at <0.05.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 Funding
 References
 
The T-ALL cell line Sup-T1 is resistant to etoposide-induced apoptosis
We first tested the response of a panel of established T-ALL cell lines to the commonly used anticancer drug, etoposide. Raji, a Burkitt lymphoma-derived cell line, which had been shown previously to be resistant to etoposide (29), was included as a control cell line. Cells treated with 1 µM of etoposide for 24 h were subjected to flow cytometric evaluation of DNA content using propidium iodide staining. Less than 10% apoptosis was detected in Raji and Sup-T1 cells. In contrast, >80% apoptosis was detected in HSB-2 and MOLT-3 cells (Figure 1A). To confirm these results, cell lysates were immunoblotted with specific antibody that recognizes full-length PARP and its cleaved form. As shown in Figure 1B, etoposide-triggered processing of PARP to its 89 kDa fragment was evident in HSB-2 and MOLT-3 cells but could not be detected in Raji and Sup-T1 cells. Neither lengthening the incubation period (48 h) nor increasing the dose of etoposide (5- and 10-fold) induced apoptosis in Raji and Sup-T1 cells (data not shown). Etoposide exerts its anticancer effect by inducing DNA damage and activating the mitochondrial-dependent apoptosis pathway (30). Thus, we believed that the resistance to etoposide-induced apoptosis in Sup-T1 cells is attributed to impairment of the effectors or regulators involved in the mitochondrial pathway. Activation of caspases is crucial to etoposide-induced apoptosis in Jurkat T cells (30). To test whether the same can be applied to our panel of T-ALL cells, cells were preincubated with z-VAD-fmk (a pan-caspase inhibitor) prior to etoposide treatment. Results from flow cytometry showed that apoptosis in the etoposide-sensitive HSB-2 and MOLT-3 cells was significantly blocked by z-VAD-fmk, from >80 to <15% (Figure 1A), suggesting that caspases are playing a critical role in etoposide-mediated apoptosis in T-ALL cells. In an independent experiment, cell lysates were immunoblotted with specific antibodies against caspase-3 and caspase-9. Etoposide-triggered processing of procaspase-3 into the p17 active subunit was readily detected in HSB-2 and MOLT-3 cells (Figure 1C, top panel). Concomitant processing of procaspase-9 into its p35 and p37 active fragments was also evident in HSB-2 and MOLT-3 extracts (Figure 1C, middle panel). However, processing of procaspase-3 and procaspase-9 were absent in etoposide-treated Raji and Sup-T1 cells. To verify these data, cells treated with etoposide were subjected to measurement of caspase-3 and caspase-9 activities according to specific cleavage of the fluorescent substrates Ac-DEVD-AFC and Ac-LEHD-AFC, respectively. The results showed that neither Ac-DEVD-AFC (Figure 1D, top panel) nor Ac-LEHD-AFC cleavage (Figure 1D, bottom panel) could be detected in etoposide-treated Raji and Sup-T1 cells. Taken together, these data demonstrate that Sup-T1 cells are resistant to etoposide-induced apoptosis and caspase activation.


Figure 1
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  		Fig. 1. Sup-T1 cells are resistant to etoposide-induced apoptosis and caspase activation. (A) Leukemic cells preincubated for 1 h in the absence (black bars) or presence (gray bars) of 50 µM the pan-caspase inhibitor, z-VAD-fmk, were treated with 1 µM etoposide for 24 h and subjected to flow cytometric analysis of DNA content using propidium iodide staining. White bars indicate control cells. Cells with hypodiploid DNA content (sub-G1population) were counted, and their numbers are expressed as a percentage of total population. Results are shown as mean values ± SDs (n = 2). *Etoposide treatment caused significant increases in the percentage of apoptotic cells compared with untreated control; #pretreatment with caspase inhibitor caused significant decreases in the percentage of apoptotic cells compared with cells treated with etoposide alone (P < 0.05). (B) Lysates from leukemic cells treated for 24 h with or without 1 µM etoposide were analyzed by western blotting for the processing of PARP. β-Actin was used as a loading control. Results are representative of three independent experiments. (C) Lysates from leukemic cells treated for 24 h with or without 1 µM etoposide were analyzed by western blotting for the processing of caspase-3 (top panel) and caspase-9 (middle panel). (D) Cells treated for 24 h with (black bars) or without (white bars) 1 µM etoposide were harvested for measurement of caspase-3 (top panel) and caspase-9 activity (bottom panel) according to specific cleavage of the fluorescent substrates DEVD-AFC and LEHD-AFC, respectively. Data are shown as mean ± SD (n = 2) and depicted as {rho}mol AFC release per minute. *P values <0.05.

 
Sup-T1 cells are insusceptible to etoposide-induced mitochondrial alterations
Etoposide triggers mitochondrial release of cytochrome c in cancer cells, resulting in activation of the caspase cascade and induction of apoptosis (30). To investigate whether etoposide could induce mitochondrial release of cytochrome c, cytosolic and mitochondrial fractions of T-ALL cells were subjected to western blotting analysis. As expected, etoposide triggered mitochondrial release of cytochrome c in HSB-2 and MOLT-3 cells, but not in Sup-T1 cells (Figure 2A). Therefore, defective caspase-3 and caspase-9 activation in Sup-T1 cells is probably attributed to the absence of mitochondrial cytochrome c release. We then used a cell-free system to further evaluate whether Sup-T1 cells are capable of promoting cytochrome c-dependent caspase activation. To this end, exogenous cytochrome c was added to S-100 cytosolic extracts derived from T-ALL cell lines. Processing of procaspase-3 into the p17 active fragment was readily detected in etoposide-sensitive HSB-2 and MOLT-3 cells, as well as in etoposide-resistant Sup-T1 cells (Figure 2B). Thus, defective caspase activation in Sup-T1 cells was due to the absence of mitochondrial cytochrome c release. These data also indicate that the death signaling downstream of mitochondria in Sup-T1 cells is robust. In Raji cells, despite etoposide could induce mitochondrial release of cytochrome c (Figure 2A), addition of exogenous cytochrome c could not restore caspase activation (Figure 2B). These observations are consistent with those published in a recent report, which showed that the resistance to etoposide-induced apoptosis in Raji cells is blocked downstream of cytochrome c release, yet upstream of caspase-3 activation. In fact, the resistance mechanism in Raji cells is due to the sequestration of apoptotic protease-activating factor-1 to the plasma membrane (29). To further delineate the apoptosis defect, we tested whether etoposide could dissipate the mitochondrial membrane potential ({Delta}{Psi}m) of T-ALL cells by flow cytometry using a potential-sensitive fluorescent probe, tetramethylrhodamine ethyl ester. Etoposide-triggered mitochondrial depolarization was evident in HSB-2 and MOLT-3, but not in Sup-T1 cells (Figure 2C). Treatment with an uncoupling agent, carbonyl cyanide 3-chlorophenylhydrazone, however, dissipated the mitochondrial membrane potential in Sup-T1 cells (Figure 2C). Altogether, these data imply that defective caspase activation in the Sup-T1 cell line could be ascribed to its insusceptibility to etoposide-induced mitochondrial alterations.


Figure 2
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  		Fig. 2. Apoptosis in Sup-T1 cells is blocked upstream of mitochondrial alterations. (A) Homogenates from leukemic cells treated for 24 h with or without 1 µM etoposide were separated into cytosolic and mitochondrial fractions. The release of cytochrome c (cyt c) was evaluated by immunoblotting of cytosolic versus mitochondrial extracts. β-Actin was used as a loading control for the cytosolic fraction. (B) S-100 cytosols were prepared from the leukemic cells, and exogenous deoxyadenosine triphosphate and cytochrome c (dATP/cyt c) were introduced to initiate caspase activation. Processing of procaspase-3 was determined by western blotting. (C) Leukemic cells treated for 24 h with or without 1 µM etoposide were subjected to flow cytometric analysis of mitochondrial membrane potential by tetramethylrhodamine ethyl ester (TMRE) staining. Carbonyl cyanide 3-chlorophenylhydrazone (CCCP) was used as a positive control for membrane depolarization. The percentages of cells (mean ± SD, n = 3) with reduction in mitochondrial membrane potential are shown.

 
BimEL is required for etoposide-induced apoptosis in Sup-T1 cells
Apoptosis that proceeds via the mitochondrial pathway involves permeabilization of the outer mitochondrial membrane, which is responsible for the release of cytochrome c and other apoptogenic molecules. This essential step is regulated by proteins of the Bcl-2 family (31). To explore whether chemoresistance in Sup-T1 cells could be explained by altered expression of Bcl-2 family members, we first determined the protein levels of 10 better-established Bcl-2 family members in T-ALL cells by western blotting. When their basal expression levels in Sup-T1 cells were compared with those in etoposide-sensitive cells, a moderate elevation of the antiapoptotic Bcl-2 was observed (Figure 3A). Moreover, expression levels of the proapoptotic members Bax and BimEL were apparently reduced in Sup-T1 cells (Figure 3A). We next considered if their altered expression levels render Sup-T1 cells resistant to etoposide-induced apoptosis. To address this question, we investigated whether silencing of Bcl-2 as well as the enforced expression of Bax and BimEL could restore sensitivity of Sup-T1 cells to etoposide-induced apoptosis. Results from gene-silencing experiments showed that transfection of a Bcl-2 antisense oligonucleotide into Sup-T1 cells significantly reduced the protein level of Bcl-2, but not in cells transfected with the corresponding, missense oligonucleotide (Figure 3B). However, the reduced level of Bcl-2 could not resensitize Sup-T1 cells to etoposide-induced caspase-3 activation and the consequent apoptosis, as judged by PARP cleavage (Figure 3B). Transient transfection with a wild-type Bax expression vector into Sup-T1 cells elevated the protein level of Bax (Figure 3C). Despite slight increases in the levels of active caspase-3 and cleaved PARP could be detected in Bax-overexpressing cells upon etoposide treatment, the levels of these fragments, however, were comparable with cells transfected with an empty vector or a Bax expression vector (Figure 3C), suggesting that an enhanced expression of Bax still could not restore etoposide-induced apoptosis. Transfection with a BimEL expression vector resulted in an elevation of BimEL protein level (Figure 3D). Indeed, overproduction of BimEL in Sup-T1 cells resulted in significant level of cell death, as indicated by increased levels of procaspase-3 and PARP processing (Figure 3D). The p17 active subunit of caspase-3 and cleaved PARP were most readily detected in BimEL-overexpressing cells following etoposide treatment (Figure 3D), indicating that enhanced level of BimEL could resensitize Sup-T1 cells to etoposide-induced apoptosis. Taken together, these data clearly reveal a critical role of BimEL in determining the sensitivity of Sup-T1 cells to etoposide-induced apoptosis.


Figure 3
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  		Fig. 3. Lack of BimEL expression renders Sup-T1 cells resistant to etoposide-induced apoptosis. (A) Leukemic cell lysates were analyzed for protein levels of the Bcl-2 family members by western blotting. (B) Sup-T1 cells were either mock transfected (lane 1) or transfected with Bcl-2-specific antisense oligonucleotide (Bcl-2 ASO, lane 3) and treated with 1 µM etoposide for 24 h (lanes 4 and 5). Cells transfected with a control oligonucleotide (lane 2) that minimally reduces Bcl-2 expression were included as a negative control. (C and D) Sup-T1 cells were either mock transfected (lane 1), transfected with an empty vector (lane 2) or wild-type Bax or BimEL expression vectors (lane 3) and treated with 1 µM etoposide for 24 h (lanes 4–6). After treatment, cell lysates were analyzed for levels of the proteins of interest as well as processing of procaspase-3 and PARP by western blotting. β-Actin was used as a loading control.

 
The reduced levels of BimEL in Sup-T1 cells are due to the presence of constitutively active JNK
Bim expression can be regulated at either transcriptional or posttranslational level (6,13–23). To investigate whether the low level of Bim protein in Sup-T1 cells could be explained by reduced transcription of the Bim gene, messenger RNA level corresponding to the BimEL isoform was determined by reverse transcription–PCR. We found that BimEL was highly expressed in Sup-T1 cells, and its expression level was even higher than those of HSB-2 and MOLT-3 cells (Figure 4A), suggesting that BimEL expression in Sup-T1 cells could be partially regulated by posttranslational mechanisms. The messenger RNA levels of Bcl2 and Bax in Sup-T1 cells, on the contrary, were found similar to those of their respective protein products (Figures 3A and 4A). The proteosome pathway has been shown previously to be involved in the degradation of Bim following its phosphorylation (7,18,23). Thus, the apparent loss of BimEL protein in Sup-T1 cells may also be resulted from the same pathway. Western blot analysis revealed that incubation with MG-132, a proteosome inhibitor, significantly increased the protein level of BimEL in Sup-T1 cells (Figure 4B). In addition, treatment with MG-132 resulted in the mobility shift of BimEL protein, as evidenced by the presence of immunoreactive species with higher molecular masses (Figure 4B). To prove that the observed mobility shift of BimEL species was due to phosphorylation, cell lysate from Sup-T1 cells treated with proteosome inhibitor was digested with calf intestinal alkaline phosphatase. Digestion with calf intestinal alkaline phosphatase resulted in a diminished mobility of BimEL species, when compared with the undigested samples (Figure 4B). By using site-specific antibodies, we found that at least two BimEL species targeted to the proteosome was preferentially phosphorylated at Ser69, but not at Ser55 (Figure 4C). This suggests that BimEL could be phosphorylated at sites other than Ser69 before targeting to the proteosome. Taken together, these data suggested that some of the newly synthesized BimEL in Sup-T1 cells was phosphorylated, serving as a signal for degradation through the proteosome pathway, and thereby resulting in the loss of its expression. Bim has been reported as substrates for ERK (18,19), JNK (20,21), p38 (22) and Akt phosphorylation (23). To investigate whether these protein kinases are also responsible for modulation of BimEL expression in Sup-T1 cells, we first determined their activities by using antibodies that recognize only their phosphorylated, active forms. Western blot analysis showed that all these kinases were highly expressed in Sup-T1 cells (Figure 4D). However, only Akt and JNK were found constitutively active in Sup-T1 cells (Figure 4D). To explore the potential roles of these protein kinases in BimEL degradation, Sup-T1 cells treated with specific inhibitors of the ERK (PD98059, U0126), JNK (SP600125), p38 (SB203580) and Akt (LY294002, wortmannin) pathways were subjected to detection of endogenous BimEL by western blotting. We found that the expression of BimEL could not be restored by the Akt pathway inhibitors, LY294002 and wortmannin. Similar results were obtained from cells treated with the ERK pathway inhibitors, PD98059 or U0126, or the p38 pathway inhibitor, SB203580 (Figure 4E). On the contrary, treatment of Sup-T1 cells with the JNK pathway inhibitor, SP600125, significantly elevated the BimEL protein level (Figure 4E), suggesting that the degradation of BimEL in Sup-T1 cells was mediated by JNK-dependent events. BimEL was found crucial for etoposide-induced apoptosis in Sup-T1 cells (Figure 3D). Then, suppression of its expression by JNK-mediated events is very likely to be the reason that renders this cell line resistant to etoposide-induced apoptosis. To address this possibility, we investigated whether the JNK inhibitor, SP600125, could resensitize Sup-T1 cells to etoposide therapy. The results revealed that the JNK activity was apparently reduced by treatment of Sup-T1 cells with SP600125, as indicated by the decreased level of phosphorylated c-Jun (Figure 4F). Again, BimEL expression was significantly elevated when the JNK activity was inhibited, but the increased level of BimEL did not result in caspase-3 activation and PARP cleavage (Figure 4F). However, processing of procaspase-3 and PARP was readily detected in cells preincubated with SP600125 followed by etoposide treatment (Figure 4F), indicating that suppression of JNK activity resensitized Sup-T1 cells to etoposide-induced apoptosis.


Figure 4
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  		Fig. 4. Activation of the JNK pathway induces phosphorylation and consequential degradation of BimEL via the proteosome. (A) Messenger RNA levels of Bcl-2, Bax and BimEL in leukemic cells were assessed by reverse transcription–PCR. Glyceraldehyde 3-phosphate dehydrogenase was used as a loading control. (B) Sup-T1 cells were cultured for 12 h in the absence (lane 1) or presence of 50 µM proteosome inhibitor, MG-132 (lane 2). After treatment, cells were harvested, and lysates were analyzed for BimEL protein levels by western blotting. Aliquots of the lysate from cells treated with MG-132 were digested with calf intestinal alkaline phosphatase (CIAP, lane 3) for 1 h, prior to western blot analysis. (C) MG-132-treated cells were detected for site-specific phospho-BimEL by western blotting. (D) Basal levels of ERK, JNK, p-38 and Akt phosphorylation in leukemic cells were determined using phospho-specific antibodies. Blots were also analyzed for the total protein levels of the kinases. (E) Sup-T1 cells were incubated for 24 h with either dimethyl sulfoxide (control), 10 µM U0126, 10 µM PD98509, 10 µM SP600125, 15 µM SB203580, 50 µM LY294002 or 100 nM wortmannin. Cell lysates were analyzed for BimEL protein levels by western blotting. (F) Sup-T1 cells were preincubated in the absence (lanes 1 and 3) or presence (lanes 2 and 4) of 10 µM JNK inhibitor, SP600125, for 1 h, and then treated with (lanes 3 and 4) or without (lanes 1 and 2) 1 µM etoposide for 24 h. The inhibitor remained present throughout etoposide treatment. Cell lysates were analyzed for p-c-Jun, c-Jun and BimEL levels, as well as processing of procaspase-3 and PARP. β-Actin was used as a loading control.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
Funding
 References
 
The cause of treatment failure in T-ALL patients is still largely unknown. Only a limited number of genes have been identified as sensitivity determinants of T-ALL cells to chemotherapeutic agents (12,32–34). In this study, we used Sup-T1 as a model T-ALL cell line to study the resistance mechanisms of chemotherapy-induced apoptosis. We showed that Sup-T1 cells were deficient in etoposide-induced activation of caspases and mitochondrial cytochrome c release. However, addition of exogenous cytochrome c in cell-free apoptosis reactions induced prominent caspase-3 activation. These findings suggest that chemoresistance observed in Sup-T1 cells is due to its insusceptibility to drug-induced mitochondrial alterations. Indeed, our data obtained in the Sup-T1 model showed that the defective apoptosis and activation of caspases are linked to the insufficient level of Bim. Transient transfection of BimEL cDNA into resistant Sup-T1 cells significantly restored their sensitivity to etoposide-induced apoptosis, thus providing direct evidence showing that the expression level of Bim is an important determinant for the sensitivity of Sup-T1 cells to drug-induced apoptosis. By using a comparative whole-genome expression profiling approach, Schmidt et al. (34) recently showed that the Bim gene is induced in childhood ALL patients who are sensitive to glucocorticoid treatment. Moreover, the induction of Bim was found essential for glucocorticoid-induced apoptosis in another T-ALL cell line, CCRF-CEM (12). A recent report also showed that dexamethasone resistance in pediatric ALL patients was associated with failure to induce the Bim gene (35). Thus, the present study provides additional evidence supporting the crucial role of Bim in drug-induced apoptosis in T-ALL cells. Since the expression of Bim is uniformly low in blast cells from patients with chronic myelogenous leukemia (36), it will be interesting to determine whether the same situation exists in chemoresistant T-ALL blasts and to establish its physiological relevance.

Expression levels of BH3-only proteins are key determinants of cellular survival and are therefore under stringent control (13). Ubiquitin-mediated degradation of Bim through the proteasomal pathway has recently emerged as a key posttranslational mechanism controlling cellular Bim level. In a number of cellular models, the loss of Bim protein can be blocked by the proteosome inhibitor MG-132 (7,18,23). Furthermore, multiple ubiquitinated forms of Bim protein could be demonstrated in the presence of proteosome inhibitors (7,18). In agreement with these studies, we showed that the level of BimEL in Sup-T1 cells was significantly restored after incubation with MG-132. Thus, the newly synthesized BimEL in Sup-T1 cells is rapidly targeted to the proteosome and degraded, therefore resulting in the lack of its protein expression. Phosphorylation at Ser69 has been established as an important signal for BimEL turnover since mutants at this site are defective in proteosomal degradation (18,19). In line with these studies, we demonstrated that at least two BimEL species targeted to the proteosome were phosphorylated at Ser69. The presence of more than one BimEL species in MG-132-treated cells suggests that BimEL was phosphorylated at other sites together with phosphorylation at Ser69. This may provide a specific pattern of phosphorylation that specifically regulates the expression of Bim in Sup-T1 cells. Previous studies showed that targeting of Bim to the proteosome for degradation requires its phosphorylation by ERK (18) or Akt (23). Interestingly, we showed that the JNK pathway may be necessary to promote BimEL phosphorylation and its turnover in Sup-T1 cells. Blockage of the JNK pathway by a specific inhibitor, SP600125, yielded elevated level of BimEL, suggesting that proteosomal degradation of BimEL requires, at least in part, an activated JNK pathway. Furthermore, BimEL species with higher molecular masses were no longer detected in SP600125-treated cells. Thus, phosphorylation of BimEL is exclusively depended on the JNK pathway. JNK phosphorylation of Bim has been demonstrated in fibroblasts (20) and neurons (21). In fibroblasts, JNK phosphorylation of BimL causes its release from the dynein motor complex, thereby allowing it to induce Bax-dependent apoptosis (20). In neurons, the mechanism by which JNK phosphorylation affects the proapoptotic activity of BimEL is still unclear. Nevertheless, evidence of Bim phosphorylation by JNK that has been documented so far is linked to the potentiation of apoptosis. In contrast, we showed for the first time that BimEL phosphorylation by the JNK pathway could has an antiapoptotic role by targeting BimEL for proteosomal degradation. The different biological consequences of JNK phosphorylation of BimEL may indicate cell type-specific regulation of Bim.

Substantial evidence suggests that the JNK-signaling pathway can contribute to cell survival. Recent data implied that the molecular mechanisms involved in JNK-mediated cell survival are highly dependent on the specific cellular context and the nature of stimulus. In a hematopoietic cell line, JNK mediates cell survival through phosphorylation and inactivation of the proapoptotic Bad (37). Moreover, endogenous JNK activity was shown to promote resistance of prostate carcinoma cells to Fas-induced apoptosis by increasing the expression of the Fas/fas-associated death domain-interacting kinase homeodomain interacting protein kinase 3 (38). In this study, we have characterized a novel mechanism by which the JNK pathway mediates cell survival in a T-ALL cell line. Of note, despite no known kinase downstream of JNK has been identified, it is not yet clear whether JNK itself is the kinase responsible for phosphorylating BimEL in our system. Further studies should be directed to define the identity of the kinase by in vitro kinase assay.


   Funding
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Funding
References
 
Endowment Fund Research Grant Scheme, United College, The Chinese University of Hong Kong (project no. CA11099); Research Fund for the Control of Infectious Diseases from the Health, Welfare and Food Bureau of the Hong Kong Special Administrative Region Government (project no. 01030582).


   Acknowledgments
 
We are grateful to Dr Hon-Ming Lam (Department of Biology, The Chinese University of Hong Kong, Hong Kong) for propagation of the constructs. We thank Dr Kathy Chan for provision of cDNA and Ms Ruby Chiu for preparation of figures.

Conflict of Interest Statement: None declared.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Funding
 References
 

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Received February 27, 2007; revised December 14, 2007; accepted December 15, 2007.


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