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. Author manuscript; available in PMC: 2014 Jun 30.
Published in final edited form as: Cancer Res. 2012 Jun 11;72(16):4193–4203. doi: 10.1158/0008-5472.CAN-11-4204

BCL2 suppresses PARP1 function and non-apoptotic cell death

Chaitali Dutta 1, Tovah Day 1, Nadja Kopp 1, Diederik van Bodegom 1, Matthew S Davids 1, Jeremy Ryan 1, Liat Bird 1, Naveen Kommajosyula 1, Oliver Weigert 1, Akinori Yoda 1, Hua Fung 1, Jennifer R Brown 1, Geoffrey I Shapiro 1, Anthony Letai 1, David M Weinstock 1,2
PMCID: PMC4075432  NIHMSID: NIHMS581586  PMID: 22689920

Abstract

BCL2 suppresses apoptosis by binding the BH3 domain of pro-apoptotic factors and thereby regulating outer mitochondrial membrane permeabilization. Many tumor types, including B-cell lymphomas and chronic lymphocytic leukemia, are dependent on BCL2 for survival, but become resistant to apoptosis after treatment. Here we identified a direct interaction between the anti-apoptotic protein BCL2 and the enzyme poly(ADP) ribose polymerase 1 (PARP1), which suppresses PARP1 enzymatic activity and inhibits PARP1-dependent DNA repair in diffuse large B cell lymphoma cells. The BH3 mimetic ABT-737 displaced PARP1 from BCL2 in a dose-dependent manner, re-establishing PARP1 activity and DNA repair and promoting non-apoptotic cell death. This form of cell death was unaffected by resistance to single-agent ABT-737 that results from upregulation of anti-apoptotic BCL2 family members. Based on the ability of BCL2 to suppress PARP1 function, we hypothesized that ectopic BCL2 expression would kill PARP inhibitor-sensitive cells. Strikingly, BCL2 expression reduced the survival of PARP inhibitor-sensitive breast cancer and lung cancer cells by 90-100%, and these effects were reversed by ABT-737. Taken together, our findings demonstrate that a novel interaction between BCL2 and PARP1 blocks PARP1 enzymatic activity and suppresses PARP1-dependent repair. Targeted disruption of the BCL2-PARP1 interaction therefore may represent a potential therapeutic approach for BCL2-expressing tumors resistant to apoptosis.

Keywords: BCL2, PARP1, ABT-737, necrosis, lymphoma

Introduction

Many tumors harbor genetic, epigenetic or post-transcriptional alterations that result in overexpression of the anti-apoptotic protein BCL2 (1-3). In these tumors, BCL2 inhibits apoptosis from a number of stresses, including DNA damage, microtubule perturbation, and oncogene activation (4), by binding the BH3 (Bcl-2 homology-3) domain of pro-apoptotic factors. Small molecules that occupy the BH3 binding pocket of BCL2, such as ABT-737 (5), are under development for the treatment of malignancies dependent on BCL2 (6, 7). These agents function by disrupting the interaction between BCL2 and pro-apoptotic BH3 domain-containing proteins, such as BIM. Upon displacement, certain pro-apoptotic proteins can initiate oligomerization of the apoptosis effector proteins BAX and BAK.

Cells can overcome dependence on BCL2 in three ways (8). First, they can upregulate other anti-apoptotic factors through cell autonomous or non-cell autonomous processes. For example, long-term selection of BCL2-overexpressing diffuse large B cell lymphoma (DLBCL) cells in the presence of ABT-737 promotes resistance through upregulation of the anti-apoptotic BCL2 family members MCL1 and/or BFL1 (9). Second, cells can downregulate pro-apoptotic signaling through changes in either gene expression or post-translational modifications. Third, cells can lose BAX and/or BAK and thereby become incapable of orchestrating intrinsic apoptosis. Each of these mechanisms can confer broad resistance to apoptosis induced by various stimuli, including DNA damaging agents and microtubule binders (10, 11).

Essentially all cases of follicular lymphoma and many cases of diffuse large B-cell lymphoma (DLBCL), chronic lymphocytic leukemia (CLL) and other subsets of non-Hodgkin lymphoma depend on BCL2 for survival (8). In the upfront setting, these diseases are broadly sensitive to an array of anti-neoplastic agents. However, relapse after treatment with one or more therapeutic regimens is commonly associated with resistance to multiple classes of chemotherapy as a consequence of epigenetic, genetic and/or post-translational modifications in the intrinsic pathway of apoptosis. Even in cells lacking autonomous resistance to apoptosis, bone marrow and lymphoid node stroma provide attachment sites and surface bound growth factors as survival signals that reduce the apoptotic response to a broad range of chemotherapies (12, 13).

Novel approaches are clearly needed for inducing death in tumor cells with resistance to apoptosis. Here we identify a novel mechanism for target non-apoptotic cell death that is dependent on PARP1 and suppressed by BCL2.

Materials and Methods

These studies were approved by the DFCI Institutional Review Board. All p-values were calculated by two-sided T-test using GraphPad software. Additional details are provided in the Supplementary Methods.

Cell culture and reagents

HT, Toledo, and OCI-LY8 were provided by Margaret Shipp. 1OCI-Ly1-7R, OCI-Ly1-10R, 1863 and 3256 were previously described (14, 15). Cell lines were not authenticated prior to these studies. N-Methyl-N′-nitro-N-nitrosoguanidine (MNNG), etoposide, and hydrogen peroxide (H2O2) were purchased from Sigma. BCL2-DKO cells were generated from a pool of DKO cells transfected with pcDNA3.1-BCL2 and selected in 1μg/mL G418 (Gold Biotechnology). ABT-737 and ABT-888 were kindly supplied by Abbott Laboratories.

Cell death assay

Cells were treated with 500μM MNNG for 15 minutes, washed with media, and cultured for 16 hours with DMSO, 100nM ABT-737, 100nM ABT-888 or combination. Cells were stained with propidium iodide (PI, 1μg/ml; Roche) and annexin V-FITC (Upstate) and flow cytometry was performed on a FACSCanto II (BD BioSciences) at the DFCI Division of Hematologic Neoplasia FACS Core Facility.

Determination of NAD and ATP concentrations

MEFs were treated 30 minutes with 500 μM MNNG in the presence of ABT-737, ABT-888, DMSO, or combinations. MTT (Sigma) and CellTiter-Glo Luminescent Cell Viability Assay (Promega) were used to determine cellular NAD and ATP, respectively, using Softmax Pro5.

Enzyme-linked immunosorbent assay

Increasing concentrations of purified GST-BCL2 were added to fractionated lysates from HT cells. Similarly, increasing concentrations of ABT-737 were added to fractionated lysates from OCI-Ly1-10R. The protein concentration of each fraction was measured using the BCA method (Thermo-Fisher). PARP1 enzymatic activity on immobilized histones was measured using an ELISA (Trevigen).

Comet assay

OCI-LY1-10R cells were treated with 50 μM MNNG for 15 minutes. Cells were collected prior to MNNG treatment (pre), immediately after washing out MNNG (post MNNG) or after washing followed by 3 hour treatment with DMSO, ABT-888 (100nM) or ABT-737 (100nM). Cells were subjected to the alkaline comet assay (Trevigen).

Immunofluorescence

Cells were fixed in 4% paraformaldehyde for 30 minutes at room temperature, and permeabilized for 10 min in 5% BSA/PBS containing 0.25% Triton X-100. Cells were washed with PBS containing 0.02% Triton X-100 and incubated with antibodies diluted in 2.5% BSA/PBS containing 0.1% Triton X-100 overnight. Images were captured on a Zeiss Axio Observer.A1 fluorescence microscope. Confocal images were captured on a Zeiss 710 laser scanning confocal microscope and analyzed using ImageJ software.

Primary CLL cell culture

The day before the experiment, 5×104 human stromal cells per well were seeded onto 48 well plates. CLL cells were collected from peripheral blood, isolated by Ficoll separation and viably frozen in liquid nitrogen. On the day of the experiment, the CLL cells were thawed and added to the stromal culture at 20:1 ratio. For comparison, CLL cells were also cultured in suspension at 3×106 cells/ml. For assessment of stroma-derived resistance, 50μM MNNG, 100nM ABT-737 and/or 200nM ABT-888 were added 24 hours later. At the indicated time points, CLL cells were collected by gently pipetting off the CLL cells. After appropriate treatments, cells were washed once with PBS and resuspended in PBS containing 1% FBS and FcR blocking reagent. Cells were stained with pacific blue-CD19 and APC-CXCR4 antibody to differentiate B cells from stromal cells. Cells were resuspended in 50μl of Annexin binding buffer and stained with Annexin V-FITC and propidium iodide. Cell numbers were quantified after the addition of 2500 Fluoresbrite counting beads (Polyscience) per sample. Mitochondrial sensitivity was determined by BH3 profiling, as described (11).

BCL2 transduction colony assay

Cells were plated in 6-well plates (1000 cells per well) the day before the retroviral infection. The cells were transduced and 24 hours later, they were selected in 1 μg/mL G418 (Gold Biotechnology). After 24 hours, cells were washed and incubated in the presence or absence of ABT-737. Cells were fixed in methanol for 1 hour and stained with Giemsa.

Results

Overexpressed BCL2 localizes in the nucleus of lymphoid tumor cells

Multiple previous studies have identified BCL2 within the nucleus of tumor cell lines and fibroblasts with high levels of BCL2 expression (16-19). We first examined BCL2 localization in the DLBCL cell lines OCI-LY1, OCI-LY8 and Toledo. All three lines harbor translocation t(14;18), which results in expression of BCL2 under IGH transcriptional control. Immunoblotting of nuclear and cytoplasmic fractions demonstrated BCL2 within the nucleus of all three cell lines in the presence or absence of MNNG, ionizing radiation (IR) or ABT-737 (Figure 1A). Immunofluorescence of OCI-LY8 cells confirmed the nuclear localization of BCL2 (Figure S1). BCL2 also localized to the nucleus in two murine B-lineage leukemia lines that overexpress BCL2 and MYC (15).

Figure 1. Nuclear BCL2 interacts with PARP1.

Figure 1

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A. Nuclear (N) and cytoplasmic fractions from Eμ-MYC/MMTV-BCL2 B-cell acute lymphoblastic leukemias (1863 and 3256) (15) were immunoblotted for BCL2, tubulin (cytoplasmic marker) and Histone H3 (nuclear marker). DLBCL lines were analyzed by immunoblotting 1 hour after irradiation or treatment with MNNG in the presence or absence of ABT-737. B. HEK293T cells were transfected with an expression plasmid for HA-BCL2 and fractionated lysates were collected after 24 hours. C. OCI-LY8 cells were exposed to ionizing radiation. After 30 minutes, whole cell lysate (WCL) or subcellular fractions were isolated and subjected to immunoprecipitation with anti-BCL2 antibody followed by separation on SDS-PAGE gel and Coomassie staining. Mass spectrometry of the ∼113 kDa band (top arrow) revealed multiple peptides from PARP1. Mass spectrometry of the lower two bands indicated by arrows identified multiple histones. D. OCI-LY8 cells were irradiated and then treated with vehicle (DMSO), ABT-737 (737), an ABT-737 enantiomer (enant), or ABT-888 (888). Subcellular fractions were immunoprecipitated with anti-BCL2 antibody and immunoblotted for BCL2, PARP1 or KU70. Loading and fractionation were confirmed by immunoblotting (Input). WCL indicates whole cell lysate. E. Immunoblotting against PAR in the presence or absence of 100 μM MNNG, 100 nM 100 nM ABT-888, the combination or 100 nM ABT-737.

To determine whether ectopically expressed BCL2 can localize to the nucleus, we transfected hemagglutinin (HA)-tagged BCL2 into 293T cells. Immunoblotting with anti-HA antibody on cellular fractions revealed that BCL2 was present in the nucleoplasm before and after irradiation (Figure 1B). Irradiation also promoted the recruitment of BCL2 to chromatin (Figure 1B). Even with ectopic expression, the nuclear and cytoplasmic levels of BCL2 remained lower than those observed in OCI-LY1 and OCI-LY8 cells (Figure S2).

BCL2 and PARP1 interact in DLBCL cells

Localization of BCL2 to irradiated chromatin suggested that BCL2 interacts with one or more factors involved in the DNA damage response. To identify proteins in the chromatin fraction that interact with BCL2, we isolated the chromatin fractions from OCI-LY8 cells after irradiation and performed immunoprecipitation with an anti-BCL2 antibody (Figure 1C). Mass spectrometry of a 113 kDa band present only in the irradiated chromatin fraction (Figure 1C) identified 18 distinct peptides from PARP1 with greater than 99% confidence (Table S1).

PARP1 plays a role in several nuclear processes, including base excision repair, transcription regulation, DNA methylation and chromatin modeling (20). PARP1 responds to DNA damage by utilizing NAD+ to transfer polymers of ADP-ribose (PAR) to acceptor proteins, including histones and PARP1 itself.

The BCL2-PARP1 interaction is disrupted by ABT-737

To determine whether the PARP1-BCL2 interaction involves the BH3 binding groove of BCL2, we exposed OCI-LY8 cells to irradiation followed by 30 minute treatment with DMSO, 100nM ABT-737 or 100nM of an inactive ABT-737 enantiomer (5). ABT-737 displaced approximately 65% of PARP1 from BCL2, while the enantiomer had no effect (Figure 1D). ABT-737 had little or no effect compared with its inactive enantiomer on the interaction between BCL2 and the nonhomologous end-joining protein KU70 (Figure 1D), which does not bind within the BH3 binding groove (18).

PARP1 undergoes auto-PARylation in response to DNA damage. This creates a negatively-charged scaffold that can mediate nonspecific protein interactions. Thus, the PARP1-BCL2 interaction could involve PAR, rather than PARP1 itself. Treatment with the PARP1 inhibitor ABT-888 (21) completely blocked MNNG-induced PARylation (Figure 1E) but had no effect on the PARP1-BCL2 interaction (Figure 1D), indicating that the interaction with BCL2 is independent of PAR.

BCL2 interacts directly with PARP1 in vitro

To clarify whether PARP1 interacts with BCL2 directly or via intermediary proteins, we performed co-immunoprecipitation on a mixture of purified PARP1 and GST-tagged BCL2. In the absence of DNA and NAD+, GSH beads coated with GST-BCL2 recovered PARP1 (Figure 2A). In contrast, no PARP1 was recovered when GSH beads were coated with either GST-BCL-xL or GST-BCL-w (Figure 2A), two BCL2 family members that are also bound by ABT-737 (5).

Figure 2. ABT-737 disrupts the BCL2-PARP1 interaction.

Figure 2

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A. Purified PARP1 was incubated with GST-tagged BCL2, GST-BCL-xL or GST–BCL-w. Mixtures were subjected to immunoprecipitation with anti-GST antibody followed by immunoblotting for PARP1. Ponceau staining was performed on immunoprecipitates to confirm equal recovery. B. Fluorescence polarization assay of GST-BCL2 mixed with increasing concentrations of ABT-737, NOXA or PARP1, as previously described (22). Error bars indicate standard error of measurement. C. GST-BCL2 and PARP1 were incubated along with increasing concentrations of ABT-737. Immunoprecipitation and immunoblotting were performed, as in (A). D. Schematic representation of full-length PARP1, deletion constructs and brct domain constructs. F1 and F2 are zinc finger 1 and zinc finger 2 domains, respectively. N indicates the nuclear localization signal. E. Bacterially expressed PARP1 proteins were separated on SDS-PAGE gel and immunoblotted with anti-PARP1 antibody. Arrows indicate the expected sizes. F. His-PARP1 contructs were incubated with purified GST-BCL2 followed by immunoprecipitation with anti-His antibody and immunoblotting against BCL2. Purified GST-BCL2 was used as a control for BCL2 immunoblotting. Incubation of the His-ΔBRCT and GST-BCL2 was performed in the presence or absence of 100 nM ABT-737. G. GST-BCL2 was incubated with brct-only or brctΔBH3 constructs followed by immunoprecipitation and immunoblotting with an antibody specific for the PARP1 BRCT domain. Ponceau staining was performed to confirm equal loading.

We performed a fluorescence polarization assay, as previously described (22), to confirm the interaction between full-length PARP1 and BCL2. As expected, full-length ABT-737 avidly interacted with BCL2 (Ki, 24.9 nM) while the negative control NOXA failed to interact with BCL2 (Figure 2B). Consistent with the findings by co-immunoprecipitation, full-length PARP1 interacted with BCL2 (Ki, 77.0 nM) (Figure 2B). The addition of ABT-737 displaced purified PARP1 from GST-BCL2 in a dose-dependent fashion (Figure 2C), confirming the ability of ABT-737 to directly disrupt the BCL2-PARP1 interaction.

The PARP1 BRCT domain is required for interaction with BCL2

Full length PARP1 includes an N-terminal DNA-binding domain, a C-terminal catalytic domain, and central PADR, BRCT and WGR domains (Figure 2D). To map the region that interacts with BCL2, we generated His-tagged C-terminal deletion mutants of PARP1 (Figure 2D). Immunoblot analysis with antibody to PARP1 confirmed all protein products (Figure 2E). We mixed equimolar amounts of each His-PARP1 protein with BCL2 and performed co-immunoprecipitation. Western blot analysis of BCL2 in eluted His-tagged protein complex revealed that a C-terminal deletion mutant lacking the BRCT domain did not interact with BCL2 (Figure 2F).

To confirm the involvement of the BRCT domain, we generated an internal BRCT domain deletion mutant (ΔBRCT), which also had markedly reduced binding of BCL2 (Figure 2F). To determine whether the PARP1 BRCT domain alone can bind BCL2, we generated a 15 kDa fragment that contains the PARP1 BRCT domain (Figure 2D). After mixing with GSH beads coated with GST-BCL2, immunoblotting with an antibody specific for the PARP1 BRCT domain demonstrated that the BRCT domain alone is sufficient to bind BCL2 (Figure 2G). The PARP1 BRCT domain contains a single BH3 motif (LXXXXD) beginning at codon 401. Mutation of both L410 and D406 to alanines markedly reduced binding of BCL2 (Figure 2G).

BCL2 blocks PARP1 enzymatic activity in DLBCL cells

To determine whether the PARP1-BCL2 interaction affects PARP1 function, we measured in vitro PARylation of immobilized histones within cell lysates from the DLBCL cell line HT, which lacks significant BCL2 expression (11). Addition of purified BCL2 reduced PARP1 activity in the nucleoplasm and chromatin fractions of HT cells (Figure 3A). In contrast, addition of MCL1, BCL-xL or BCL-w had no effect on PARP1 activity (Figure 3B). These results are similar to a previous report in HL60 and U937 cells, in which ectopic overexpression of BCL2 suppressed both basal PARP1 activity and the PARP1 response to a topoisomerase II inhibitor (23).

Figure 3. BCL2 suppresses PARP1 enzymatic function.

Figure 3

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A. Purified GST-BCL2 was added to fractionated lysates from HT cells or ABT-737 was added to fractionated lysates from OCI-LY8 cells. PARP1 activity on immobilized histones was measured by enzyme linked immunosorbent assay. Error bars represent standard error of measurement. B. GST-MCL1, GST-BCL-xL and GST-BCL-w were added to nucleoplasm extracts from HT cells and PARP1 activity was assayed, as above. C. OCI-LY1-10R cells treated with MNNG for 15 minutes were washed and exposed to vehicle (DMSO), ABT-888 or ABT-737. Cells were collected prior to MNNG treatment (pre), immediately after washing MNNG (post MNNG) or 3 hours after wash. Olive moments were quantified for 500 cells in each category. Error bars indicate standard error of measurement. Pictures are 100X magnification.

To determine whether ABT-737 can increase PARP1 activity by displacing BCL2 from PARP1, we measured the effect of ABT-737 on PARP1 activity in fractionated lysates from OCI-LY8 cells. ABT-737 potentiated PARP1 enzymatic activity in a dose-dependent manner in both nucleoplasm and chromatin fractions (Figure 3A). Together, these findings indicate that the PARP1-BCL2 interaction blocks PARP1 activity while ABT-737 can displace PARP1 from BCL2 and restore that activity.

Next, we asked whether inhibition of PARP1 function by BCL2 suppresses PARP1-dependent DNA repair. Assaying the effects of BCL2 inhibition on DNA repair is complicated by the dependence of most BCL2-overexpressing cells on BCL2 for survival. We utilized OCI-LY1 and SU-DHL4 clones previously selected for resistance to ABT-737 by long-term culture in the presence of the drug (24). These clones maintain BCL2 expression but have also upregulated the anti-apoptotic BCL2 family members MCL1 and/or BFL1(24).

OCI-LY1-10R cells were treated for 15 minutes with MNNG, which is known to activate PARP1-dependent DNA repair (25). After washing away MNNG, the cells were allowed to repair the damage for 3 hours in the presence of vehicle, ABT-888 or ABT-737. We performed the alkaline comet assay, which quantifies both single- and double-stranded DNA breaks. Cells treated with ABT-737 had significantly smaller comet tails, indicating more extensive resolution of MNNG-induced DNA damage compared to cells treated with MNNG and vehicle (Figure 3C). The addition of ABT-888 did not further suppress repair, suggesting that most of the PARP1-mediated repair in these cells is inactive (Figure 3B). Of note, neither ABT-737 nor MNNG affect the nuclear localization of BCL2 (Figure 1A).

BCL2 blocks PARP1-dependent, non-apoptotic death in DLBCL cells

PARP1 enzymatic activity is required for a form of non-apoptotic cell death previously termed ‘parthanatos’ (26-28). This pathway can be induced by ischemia or treatment with the alkylating agent MNNG, which both result in DNA damage that requires PARP1-dependent repair. This form of cell death does not involve the intrinsic apoptosis pathway, as wild-type and Bax-/-Bak-/- mouse embryonic fibroblasts have similar sensitivity to MNNG-induced non-apoptotic death (25, 29).

We hypothesized that inhibition of PARP1 enzymatic activity by BCL2 would suppress this form of non-apoptotic cell death. Supporting this hypothesis, previous studies have reported that BCL2 can inhibit nitric oxide-induced, non-apoptotic cell death of PC12 and HeLa cells (30) as well as non-apoptotic death of neural cells induced by glutathione depletion (31). We assayed the survival of ABT-737-resistant clones treated with MNNG, ABT-737, ABT-888 or combinations (Figure 4A). As expected, very little cell death was observed with single-agent ABT-737 (Figure 4A). However, ABT-737 significantly increased death induced by MNNG in ABT-737-resistant OCI-LY1 and SU-DHL4 clones as well as Toledo cells (Figure 4A, 4B; p<0.05 compared to MNNG plus vehicle in all lines). Importantly, MNNG+ABT-737-induced death was partially reversed by ABT-888, supporting a specific role for PARP1 (Figure 4A, 4B).

Figure 4. BCL2 suppresses MNNG-induced death in DLBCL cells.

Figure 4

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A. Cells were treated with indicated agents for 20 hours and analyzed by flow cytometry after staining with propidium iodide (PI) and FITC-conjugated annexin V. Percent alive is the fraction of Annexin V/PI-negative cells relative to the same cell line treated with DMSO and no MNNG. Error bars indicate standard deviation. * indicates p<0.05. B. Example of flow cytometric plots of OCI-LY1-10R cells treated with agents for 20 hours and stained with PI and FITC-annexin V. C. Transmission electron micrographs of OCI-LY1-10R cells fixed 9 hours after treatment with DMSO, 500 μM MNNG, 100 nM ABT-737, 50 μM etoposide or combinations.

Transmission electron microscopy of OCI-LY1-10R and SU-DHL4-2R cells treated with MNNG+ABT-737 demonstrated ultrastructural features of non-apoptotic cell death, including loss of membrane integrity with preservation of nuclear architecture (Figure 4C). These findings are most consistent with necrotic cell death. Double-membrane structures characteristic of autophagy were not present. In contrast, treatment with ABT-737 plus the topoisomerase II poison etoposide resulted in features characteristic of apoptosis, including chromatin condensation and preservation of membrane integrity (Figure 4C). Thus, ABT-737 can promote PARP1-dependent non-apoptotic death in DLBCL cells.

We treated Bax-/-Bak-/- MEFs (referred to as ‘DKO cells’) with MNNG and confirmed that non-apoptotic cell death is independent of BAX and BAK (Figure 5A, 5B). DKO cells that stably overexpress BCL2 (‘BCL2-DKO cells’) were less sensitive to MNNG (p<0.05 compared to DKO cells) (Figure 5A), consistent with the inhibition of PARP1-dependent cell death by BCL2. In both lines, MNNG-induced death was completely blocked by ABT-888 (Figure 5A). Thus, BCL2 inhibition can promote at least two forms of cell death. The first is BAX/BAK-dependent intrinsic apoptosis, while the second involves PARP1-dependent non-apoptotic death that is independent of BAX and BAK.

Figure 5. ABT-737 promotes MNNG-induced cell death in BCL2-overexpressing cells that is independent of BAX/BAK.

Figure 5

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A. Bax-/-Bak-/- MEFs (DKO) and BCL2-transgenic Bax-/-Bak-/- MEFs (BCL2-DKO) were treated with indicated agents for 20 hours and analyzed by flow cytometry after staining with propidium iodide (PI) and FITC-conjugated annexin V. Percent alive is the fraction of Annexin V/PI-negative cells relative to the same cell line treated with DMSO and no MNNG. Error bars indicate standard deviation. * indicates p<0.05. B. Transmission electron micrographs at 890X and 2900X magnifications fixed 9 hours after treatment with the indicated agents. C. Concentrations of NAD+ and ATP were determined in lysates isolated after 9 hour treatment with MNNG and the indicated agents. All values are relative to the same cell line treated with vehicle and no MNNG.

PARP1-mediated cell death induced by MNNG is characterized by reductions in cellular NAD+ and ATP (25, 29). We confirmed that both NAD+ and ATP concentrations are significantly reduced in DKO cells after 30 minute treatment with MNNG (Figure 5C). In DKO cells, reductions in NAD+ and ATP after MNNG treatment were attenuated by the PARP1 inhibitor ABT-888 but were unaffected by ABT-737 (Figure 5C). MNNG treatment led to a lesser reduction of ATP levels in BCL2-DKO cells, consistent with the attenuation of PARP1 activity by BCL2 (Figure 5C). Unlike DKO cells, ABT-737 reduced the levels of NAD+ and ATP in MNNG-treated BCL2-DKO cells (p<0.05 for both NAD+ and ATP), and these effects were significantly reversed by ABT-888 (p<0.05 for both NAD+ and ATP; Figure 5C).

ABT-737 promotes non-apoptotic death in primary CLL cells independent of stromal protection

We hypothesized that non-apoptotic cell death induced by MNNG+ABT-737 could be utilized to overcome stroma-induced resistance to apoptosis. CLL cells are known to be dependent on BCL2 and sensitive to treatment with ABT-737 (32). Co-culture of primary CLL cells with murine or human marrow stromal cells (HMSCs) reduces apoptosis of CLL cells upon treatment with antineoplastic agents or ABT-737 through upregulation of anti-apoptotic BCL2 family members (12, 13).

We cultured primary CLL cells from 14 patients (Table S2) with or without HMSCs for 48 hours (Figure 6). As expected, cells cultured in the presence of human HMSCs downregulated CXCR4 (Figure 6A), the CLL cell surface homing receptor for SDF-1/CXCR12 (12, 33). ABT-737 potently induced mitochondrial depolarization in primary CLL cells cultured in the absence of HMSCs (Figure 6B) while cells co-cultured with HMSCs had significant resistance to 100 nM ABT-737 (p<0.01; Figure 6C-D). In contrast, coculture with HMSCs did not block the additive killing from the combination of 50 μM MNNG plus 100 nM ABT-737 but killing was reduced by ABT-888 (p<0.05; Figure 6C-D), demonstrating partial dependence on PARP1. Of note, 100 nM ABT-737 plus 50 μM MNNG resulted in only 20% loss of viability among the HMSC cells (data not shown).

Figure 6. ABT-737 promotes non-apoptotic death in primary CLL cells independent of stroma.

Figure 6

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A. CLL cells were cultured with or without primary marrow stromal cells (HMSCs) and surface CXCR4 expression was quantified by flow cytometry. B. Mitochondria from six CLL samples were exposed to ABT-737 and the relative depolarization was quantified based on cytochrome C release, as previously described (32). FCCP was used as a positive control. C. CLL cells from patient D1001 were cultured in the presence or absence of HMSCs for 24 hrs and then treated with DMSO, 100nM ABT-737, 50μM MNNG, 200nM ABT-888 or combinations, as shown. The cells were stained with FITC-conjugated Annexin V, propidium iodide and pacific blue-conjugated CD19 antibodies. CD19-positive cells were gated and cells negative for both PI and Annexin V were considered alive. D. Percent of cells alive under various treatment conditions for all 16 CLL samples. Bars indicate mean and 95% confidence interval. * indicates p<0.05.

BCL2 expression induces death in PARP inhibitor-sensitive cells

A logical extension of our findings that BCL2 overexpression phenocopies PARP1 inhibition is that cells sensitive to therapeutic PARP inhibitors could be killed by overexpression of BCL2. As a proof-of-principle, we retrovirally transduced BCL2 into the BRCA1-mutated breast cancer cell line MDA-MB-436 (34). Strikingly, ectopic expression of BCL2 completely blocked colony formation, which was partially reversed by co-administration of ABT-737 (Figure 7A).

Figure 7. Model for PARP1 involvement in DNA repair and cell death.

Figure 7

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A. MDA-MB-436 breast cancer cells were transduced with BCL2 or control virus and selected in neomycin in the presence or absence of increasing concentrations of ABT-737. The experiment was performed in triplicate. B. Sensitivity of lung cancer cell lines to the PARP1 inhibitor AG014699. A replicate experiment is shown in Figure S3. C. Lung cancer cells were transduced and selected, as in (A). Representative data of three independent experiments is shown. D. Basal DNA damage from endogenous processes like replication requires PARP1 for repair. In the absence of functional PARP1, single-strand lesions can become double-strand breaks that require homology-directed repair (HDR) for resolution. In cells with impaired HDR, the accumulation of DNA double-strand breaks induces apoptosis. Similarly, extensive DNA damage from irradiation or treatment with alkylating agents can induce apoptosis, which is suppressed by BCL2. The same DNA damage can initiate repair and non-apoptotic cell death dependent on PARP1. The latter pathway is also suppressed by BCL2.

To expand this finding, we screened approximately 30 lung cancer cell lines for sensitivity to the PARP1 inhibitor AG014699. The lung cancer cell lines H520 (GI50, 30 nM) and HCC827 (GI50, 65 nM) were highly sensitive to AG014699, compared with insensitive lines like H1299 (GI50, >1 μM) (Figures 7B, S3). Similar to MDA-MB-436 cells, ectopic expression of BCL2 in the PARP inhibitor-sensitive cell lines reduced colony formation by 90-100% but did not affect colony formation in H1290 cells (Figure 7C). The effect from BCL2 was reversed by ABT-737 at therapeutic concentrations (Figure 7C).

Discussion

BCL2 suppresses the intrinsic pathway of apoptosis by binding pro-apoptotic BCL2 family members (8). We have identified two additional pathways that can modulate survival through BCL2. First, inhibition of PARP1 activity by BCL2 delays the repair of DNA damage as well as non-apoptotic cell death. The pharmacologic BH3 mimetic ABT-737, which was designed to induce BAX/BAK-dependent apoptosis in cells dependent on BCL2 (5), can also promote PARP1-dependent non-apoptotic death by dissociating PARP1 from BCL2 and thereby restoring PARP1 enzymatic activity.

The second new pathway of survival modulation involving BCL2 results from the ability of BCL2 overexpression to phenocopy PARP deficiency. More specifically, BCL2 overexpression in PARP inhibitor-sensitive cells can promote cell death and this death can be reversed by interrupting the interaction between BCL2 and PARP1. The “pro-death” effect from BCL2 and the “pro-survival” effect from ABT-737 is specific to cells that lack the ability to repair damage that results from PARP deficiency. In this model (Figure 7D), basal levels of damage are repaired through PARP1 without drastically affecting cellular energy stores. PARP1 deficiency, either through interaction with BCL2 or pharmacologic inhibition, results in the conversion of single-strand lesions to double-strand breaks, which can be repaired by homology-directed repair (HDR) (35, 36). Extensive DNA damage resulting from exogenous clastogens can drive both apoptosis and PARP1-mediated non-apoptotic cell death, which are both suppressed by BCL2.

The extent of damage necessary to induce PARP1-mediated non-apoptotic death within any particular cell is likely to depend on several factors, including energy metabolism, repair efficiency, threshold for undergoing apoptosis (37), and the downstream effectors of PARP1-dependent death. One testable prediction based on our findings is that tumors with BCL2 overexpression must maintain some extent of HDR, as concurrent deficiencies in HDR and PARP1 results in synthetic lethality (35, 36). Challenging our model, overexpression of BCL2 or BCL-XL can suppress HDR in rodent (38) and human (39, 40) cells. In addition, BCL2 inhibits the formation of radiation-induced BRCA1 foci in human lymphoma cells (40), further suggesting that BCL2 may negatively modulate HDR. However, another study in human lymphoblastoid cells noted the opposite effect, with a 3-fold increase in HDR upon BCL-XL overexpression (41).

A series of studies have reported additional effects of BCL2 in various cell types on other DNA pathways, including nucleotide excision repair, base excision repair, mismatch repair and homologous recombination (16-19, 38, 39, 42-44). The described effects from BCL2 include both direct binding and decreased transcription of DNA repair factors. The extent to which these effects are present within lymphoid tumor cells and their contribution to sustaining DNA damage after treatment with clastogens has not been addressed.

Several aspects of the BCL2-mediated suppression of PARP1 activity remain unclear. First, the precise function of the PARP1 BRCT domain is poorly understood. In chicken DT40 cells, the PARP1 BRCT domain appears to be involved in mutagenic repair at immunoglobulin loci, possibly through an interaction with nonhomologous end-joining factors (45, 46). Aberrant somatic hypermutation at oncogene loci is a common feature of lymphomas with BCL2 overexpression (47). Thus, it is intriguing to speculate that BCL2-mediated suppression of PARP1 function could promote these events by inhibiting more fidelitous repair.

It also remains unclear whether targeting non-apoptotic death would offer an adequate therapeutic index, as some populations of non-neoplastic cells may be similarly sensitive to PARP1-mediated death. Zong et al. identified greater sensitivity to MNNG-induced death in proliferating versus vegetative cells, despite equivalent amounts of DNA damage and PARP activity (48). The greater sensitivity among proliferating cells resulted from dependence on glycolysis for the generation of ATP (i.e. the Warburg effect).

In conclusion, we have identified a novel interaction between BCL2 and PARP1 that blocks PARP1 enzymatic activity and suppresses PARP1-dependent repair. Targeting of the PARP1-BCL2 interaction with BH3 mimetics is a potential approach for killing BCL2-expressing tumors that are resistant to apoptosis, either through upregulation of other anti-apoptotic BCL2 family members or from defects in the intrinsic pathway (11, 13).

Supplementary Material

Supplementary Tables, Figures and Methods

Acknowledgments

The authors would like to thank Margaret Shipp and Bjoern Chapuy for providing reagents and helpful discussion. We thank Maria Ericsson, Simon Dillon and John Daley for assistance with electron microscopy, mass spectrometry and flow cytometry, respectively.

Grant support: This work was supported by a Burroughs-Wellcome Fund Career Award in the Biomedical Sciences (6150801), the Claudia Adams Barr Program in Cancer Research and the John Stellato Fund. TD is supported by an American Cancer Society Fellowship. NK and GIS are supported by the DF/HCC SPORE in Lung Cancer P50 CA90578.

Footnotes

The authors report no conflicts of interest

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Supplementary Materials

Supplementary Tables, Figures and Methods
NIHMS581586-supplement-Supplementary_Tables__Figures_and_Methods.pdf (14.9MB, pdf)

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