Cisplatin Resistance Associated with PARP Hyperactivation
Judith Michels1,4,5, Ilio Vitale1,4,5, Lorenzo Galluzzi4,10,12, Julien Adam2,4,5, Ken Andr�e Olaussen2,4,5,Oliver Kepp1,4,5, Laura Senovilla1,4,5, Ibtissam Talhaoui6, Justine Guegan3,7, David Pierre Enot1,9,10,12,Monique Talbot3, Ang�elique Robin2,4,5, Philippe Girard11, C�edric Or�ear8, Delphine Lissa1,4,5,Abdul Qader Sukkurwala1,4,5, Pauline Garcia1,4,5, Parviz Behnam-Motlagh15, Kimitoshi Kohno16,Gen Sheng Wu17, Catherine Brenner14, Philippe Dessen3, Murat Saparbaev6, Jean-Charles Soria2,4,5,Maria Castedo1,4,5, and Guido Kroemer1,2,9,10,12,13
AbstractNon–small cell lung carcinoma patients are frequently treated with cisplatin (CDDP), most often yielding
temporary clinical responses. Here, we show that PARP1 is highly expressed and constitutively hyperactivatedin a majority of human CDDP-resistant cancer cells of distinct histologic origin. Cells manifesting elevatedintracellular levels of poly(ADP-ribosyl)ated proteins (PARhigh) responded to pharmacologic PARP inhibitorsas well as to PARP1-targeting siRNAs by initiating a DNA damage response that translated into cell deathfollowing the activation of the intrinsic pathway of apoptosis. Moreover, PARP1-overexpressing tumor cellsand xenografts displayed elevated levels of PAR, which predicted the response to PARP inhibitors in vitro andin vivo more accurately than PARP1 expression itself. Thus, a majority of CDDP-resistant cancer cells appearto develop a dependency to PARP1, becoming susceptible to PARP inhibitor–induced apoptosis. Cancer Res;73(7); 2271–80. �2013 AACR.
IntroductionSeveral among the 18 PARP proteins described so far
constitute prospective targets for anticancer therapy(1, 2). PARP1 and 2 are the most abundant and best-char-acterized members of this family and are involved in tran-scriptional regulation, DNA repair, as well as in the main-tenance of genomic stability (3–5). PARP1, which is mostlylocalized to the nucleus, accounts for approximately 75%
of overall PARP enzymatic activity, fixing PAR polymers onmultiple nuclear, cytoplasmic, and mitochondrial sub-strates, including proteins that modulate chromatin struc-ture (e.g., histones, topoisomerases I and II), factors thatstimulate DNA synthesis and repair (e.g., XRCC1, DNApolymerases a and b, DNA ligases I and II), and transcrip-tional regulators (e.g., p53; refs. 6–8).
PARP1 has been implicated in DNA repair via the baseexcision repair (BER) pathway (9). Upon DNA damage, PARP1is recruited to single-strand breaks (SSB), where it becomesactivated (10) and catalyzes the poly(ADP-ribosyl)ation of localsubstrates, including histones. As poly(ADP-ribosyl)ationadvances and PAR polymers extend, histones progressivelyacquire negative charges, causing their electrostatic repulsionfrom interacting proteins and DNA (11). As a result, nuclearpoly(ADP-ribosyl)ation facilitates the relaxation of supercoiledDNA structures, improving the accessibility of DNA to repairenzymes (12).
PARP1 is characterized by 3 functionally distinct domains:(i) an N-terminal DNA-binding domain that includes 2 zincfingers for the detection of DNA SSBs (13, 14); (ii) an interme-diate domainwith autoinhibitory poly(ADP-ribosyl)ation sites,and (iii) a C-terminal catalytic domain that fixes NAD andtransfers its ADP-ribose moiety (which bears one negativecharge) to proteins in an ATP-consuming reaction (15). Severalchemically distinct inhibitors of PARP have been synthesizedand explored for their therapeutic profile against cancer, eitheras single agents or within combination regimens (16, 17).Current clinical trials are evaluating the safety and antineo-plastic profile of at least 9 distinct PARP inhibitors (http://www.clinicaltrials.gov/).
Authors' Affiliations: 1U848, 2U981, 3U985, INSERM; 4Institut GustaveRoussy; 5Universit�e Paris Sud; 6UMR8200, CNRS; Plateformes de 7Bioin-formatique, 8G�enomique, and 9Metabolomique, Institut Gustave Roussy;Villejuif; 10Universit�e Paris Descartes; 11Department of Thoracic Oncology,InstitutMutualisteMontsouris; 12Equipe 11 labellis�ee par la LigueNationalecontre le Cancer, Centre de Recherche des Cordeliers; 13Pole de Biologie,Hopital Europ�een Georges Pompidou AP-HP, Paris; 14U769, INSERM-LabEx LERMIT, Universit�e Paris-Sud, Chatenay Malabry, France;15Department of Medical Biosciences, Clinical Chemistry, Umea
�Univer-
sity, Umea�, Sweden; 16Department of Molecular Biology, School of Med-
icine, University of Occupational and Environmental Health, Kitakyushu/Fukuoka, Japan; and 17Molecular Therapeutics Program, KarmanosCancer Institute, Department of Oncology and Pathology, Wayne StateUniversity School of Medicine, Detroit, Michigan
Note: Supplementary data for this article are available at Cancer ResearchOnline (http://cancerres.aacrjournals.org/).
J.-C. Soria, M. Castedo, and G. Kroemer share senior coauthorship.
I. Vitale, L. Galluzzi, and J. Adam contributed equally to this work.
Corresponding Authors:Guido Kroemer, INSERMU848, Institut GustaveRoussy, 114 rue Edouard Vaillant, F-4805 Villejuif, France. Phone: 33-1-4211-6046; Fax 33-1-4211-6047; E-mail: [email protected]; and MariaCastedo, E-mail: [email protected]
PARP inhibitors are currently being evaluated for thetreatment of tumors bearing loss-of-function mutations ofBRCA1 and BRCA2 (18, 19), in particular breast, ovarian, andprostate cancers. BRCA1 mutations compromise DNA repairvia homologous recombination (HR), obliging cells to correctdouble-strand breaks via the less accurate non-homologousend-joining (20). By interfering with BER, PARP inhibitorsindirectly promote the accumulation of DNA double-strandbreaks (20). In line with this notion, PARP inhibition and loss-of-function BRCA1mutations display a robust synthetic lethal-ity (21). Although initial phase I clinical trials tended to supportthis concept (22, 23), less promising results have been obtainedin phase II studies (18, 19), raising an urgent need for predictivebiomarkers. In addition, PARP inhibitors have been used tosensitize cancer cells to DNA-damaging agents including (butnot limited to) ionizing irradiation, the alkylating agent temo-zolomide, the topoisomerase inhibitor camptothecin as well asplatinum compounds such as cis-diammineplatinum(II)dichloride, also known as cisplatin (CDDP), and carboplatin(24–26). Some of these combinatorial chemotherapeutic regi-mens are currently under clinical evaluation (27, 28).
Driven by the facts that non–small cell lung carcinoma(NSCLC) is the leading cause of cancer-related morbidityand mortality worldwide (29) and that NSCLC patientsoften develop resistance against CDDP-based therapies, weaddressed the question as towhetherNSCLCcellsmay respondto PARP inhibitors. Here, we report the unexpected findingsthat NSCLC cells that have become resistant to CDDP oftenoverexpress PARP1, constitutively exhibit high levels of PARPenzymatic activity and apparently rely on it for their survival, asPARP inhibition kills NSCLC cells that harbor hyperactivatedPARP1.
Materials and MethodsCell lines, culture conditions, and chemicals
The following culture media were used for both wild-type(WT) cells and their CDDP-resistant counterparts (generat-ed as depicted in Supplementary Fig. S1A): Glutamax-con-taining Dulbecco's Modified Eagle's Medium/F12 mediumfor human NSCLC A549 cells; RPMI-1640 medium for humanNSCLC H460 and H1650 cells; MCDB105/M199 medium forhuman ovarian cancer TOV-112D cells, and Eagle's MinimalEssential culture medium with Earl's salts for human meso-thelioma P31, NSCLC H1299, and cervical carcinoma HeLacells. Media for cell culture were invariably supplementedwith 10% fetal bovine serum, 10 mmol/L HEPES buffer, 100U/mL penicillin G sodium salt, and 100 mg/mL streptomycinsulfate. Cell lines were routinely maintained at 37�C under5% CO2 in T175 flasks and seeded in appropriate supports(6, 12, or 96 wells plates) 24 hours before experimentaldeterminations. Authenticated WT cells were obtained fromAmerican Type Culture Collection, immediately amplified toconstitute liquid nitrogen stocks and (upon thawing) neverpassaged for more than 1 month before use in experimentaldeterminations. Once generated, CDDP-resistant cells weretreated similarly. CEP 8983-07 (CEP) and Z-Val-Ala-Asp(OMe)-fluoromethylketone (Z-VAD-fmk) were purchasedfrom Cephalon and Bachem, respectively.
RNA interferencesiRNA heteroduplexes specific for PARP1 (PARP1.a, sense
50-CAAACUGGAACAGAUGCCGdTdT-30; PARP1.b, sense 50-GCCUCCGCUCCUGAACAA UdTdT-30; PARP1.c, sense 50-GA-UAGAGCGUGAAGGCGAAdTdT-30; ref. 30), as well as 2 non-targeting siRNAs (UNR, sense 50-GCCGGUAUGCCGGUUAA-GUdTdT-30; EMR, sense 50-CCGUGCUCCUGGGGCUGGGdT-dT-30; ref. 31), were purchased from Sigma-Aldrich. A549 cellspreseeded in 12-well plates were transfected with siRNAs at30% to 40% confluence by means of the HiPerFect transfectionreagent (Qiagen), as previously described (32, 33). Cells wereused for experiments no earlier than 24 hours aftertransfection.
Cytofluorometric studiesFor the simultaneous quantification of plasma membrane
integrity and mitochondrial transmembrane potential (Dym),both adherent and nonadherent cells were collected, washed,and costained with 1 mg/mL propidium iodide (PI, which onlyincorporates into dead cells) and 40 nmol/L 3,30-dihexylox-acarbocyanine iodide DiOC6(3), a mitochondrial transmem-brane potential (Dym)-sensitive dye, (Molecular Probes–Invi-trogen), following standard protocols (34, 35). Cytofluoro-metric acquisitions were conducted on a FACSCalibur (BDBiosciences), FACScan (BD Biosciences), or Gallios cytometer(Beckman Coulter). First-line statistical analyses were per-formed by means of the CellQuest (BD Biosciences) or Kaluzasoftware (Beckman Coulter), upon gating on events exhibitingnormal forward scatter and side scatter parameters.
ImmunoblottingCells were collected, washed with cold PBS, and lysed as
previously described (32, 36). Thereafter, protein extracts (30mg/lane) were separated on precast 4% to 12% SDS-PAGE gels(Invitrogen), followed by electrotransfer to Immobilon mem-branes (Sigma-Aldrich) and immunoblotting with antibodiesspecific for PAR (10H; Calbiochem, Merck KGaA), PARP1 (CellSignaling Technology Inc.), POLb (Abcam), and XRCC1(Abcam). An antibody that recognizes glyceraldehyde-3-phos-phate dehydrogenase (GAPDH; Millipore-Chemicon Interna-tional) was used to monitor equal lane loading. Finally,membranes were incubated with appropriate horseradishperoxidase-conjugated secondary antibodies (Southern Bio-tech), followed by chemiluminescence detection with theSuperSignal West Pico reagent and either CL-XPosure X-rayfilms (both from Thermo Scientific-Pierce) or the ImageQuantLAS 4000 Biomolecular Imager (GE Healthcare Life Sciences).
Clonogenic survival assaysTo evaluate clonogenic survival, cells were seeded at differ-
ent concentrations (from 0.2 to 4 � 103 per well) into 6-wellplates, let adhere overnight, treated for 48 hours with CDDPand PARP inhibitors, and then cultured in drug-free mediumfor up to 10 days. Eventually, colonies were stained with0.25% (w/v) crystal violet, 70% (v/v) methanol, and 3%(v/v) formaldehyde in water and quantified as previouslydescribed (37). Only colonies made of 30 cells and more wereincluded in the quantification. The surviving fraction (SF) was
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calculated according to the formula: SF¼ (number of coloniesformed/number of cells seeded).
In vivo experimentsMice were maintained in specific pathogen-free conditions
and all animal experiments were approved by the local EthicsCommittee (CEEA IRCIV/IGR n�26, registered with the FrenchMinistry of Research), were in compliance with Directive EU63/2010 and followed the Federation for Laboratory AnimalScience Associations (FELASA) guidelines. Athymic nu/nu 8weeks old female mice (Charles River Laboratories) weresubcutaneously xenografted with 1 � 106 WT or CDDP-resis-tant R6 A549 cells (suspended in 200 mL PBS). Mice weretreated starting from day 8 postinoculation with 5 mg/Kg PJ34or an equivalent volume of vehicle (PBS) intraperitoneally 3times a week for 3 consecutive weeks. Tumor growth wasroutinely monitored with a common caliper, and tumor vol-ume was calculated according to the formula V ¼ maximaldiameter � perpendicular diameter2.
Statistical proceduresUnless otherwise specified, all experiments were conducted
in triplicates and independently repeated at least 2 times,yielding comparable results. Data were analyzed with Micro-soft Excel (Microsoft Co.) and statistical significance wasassessed by means of one-tailed Student t tests. P-values wereconsidered significant when lower than 0.05. Minimum effec-tive doses (MED)were defined as the smallest dose to achieve astatistically significant effect over control conditions. MEDswere computed on the basis of a 4-parameter logistic or 3-parameter Emax dose-response models (38). When MEDscould not be determined within the dose range, maximumdoses are reported. Paired Wilcoxon tests were used to assessthe one-sided hypothesis that MEDs would not differ betweenWT cells and their CDDP-resistant counterparts. See alsoSupplementary Information.
ResultsPARP inhibitors interactwithCDDP to induceNSCLC celldeathTo identify pharmacologic agents that may influence the
activity of the commercial PARP inhibitor PJ34 hydrochloridehydrate (PJ), we assessed nuclear shrinkage (pyknosis), whichis indicative for the induction of apoptosis (35, 39), in NSCLCA549 cells exposed to 1,040 U.S. Food and Drug Administration(FDA)-approved drugs alone or combined with PJ for 48 hours.This screen led to the identification of 10 compounds thatinduce significant apoptosis in the presence, but not in theabsence, of PJ, including alkylating agents such as CDDP,thiotepa, dacarbazine, and mitomycin C (Fig. 1A and B). Theapparent interaction between PJ and CDDPwas confirmed in aset of independent experiments (Fig. 1C), and this observationextended to another PARP inhibitor, CEP, whose precursor(CEP-9722) is currently being tested in phase I clinical trials(http://www.clinicaltrials.gov; Fig. 1D). On the basis of theseresults, we decided to investigate the possibility that PARPinhibitors might be useful for the treatment of tumors thatbecome insensitive to CDDP-based chemotherapy.
Selective killing of CDDP-resistant cell clones by PARPinhibitors
CDDP-resistant NSCLC cell clones were isolated from A549cells that had been cultured in the continuous presence ofCDDP (2 mmol/L for 6months followed by 5 mmol/L for 1 year).This procedure generated nine A549 cell clones (R1–R9) that—1 month after the withdrawal of CDDP—exhibited variableextents of CDDP resistance. Thus, as compared with parental,WT A549 cells, all resistant clones exhibited a reducedfrequency of dying (DiOC6(3)
lowPI�) and dead (PIþ) cellsupon exposure to CDDP concentrations ranging from 10 to50 mmol/L (Fig. 2A and B and Supplementary Fig. S1). To getfurther insights into the cytotoxic activity of PARP inhibitors in
Figure 1. Interactions between PARP inhibitors and CDDP. A and B,humanNSCLCA549 cells were cultured in the presence of 1,040 differentcompounds from theUSDrugCollection (final concentration¼ 10mmol/L)alone or in combination with 20 mmol/L PJ for 48 hours, and nuclearshrinkage was measured by robotized fluorescence microscopy andautomatic image analysis upon Hoechst 33342 staining. Representativemicrophotographs and Z-scores are shown in A (scale bar, 10 mm)andB, respectively. Chemicals that inducednuclear shrinkage exclusivelywhen combined with PJ and in a statistically significant fashion arelisted. C and D, A549 cells were kept in control condition or exposed for48hours to the indicatedconcentrationsofPJandCEPaloneorcombinedwith 10 mmol/L CDDP, followed by the image-based quantification ofnuclear shrinkage. Data are reported asmeans� SEM (n¼ 3). �, P < 0.05;���, P < 0.001 (Student t test), compared with cells treated with the samePARP inhibitor at the same concentration.
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Figure 2. CDDP-resistant human NSCLC cell clones are susceptible to cell death induced by PARP inhibition. A–D, WT human NSCLC A549 cells and3 CDDP-resistant derivatives (R) were maintained in control conditions or treated with increasing concentrations of CDDP (10, 15, 20, 30, and50 mmol/L; A and B), CEP (1.25, 2.5, 5, and 10 mmol/L; C), or PJ (10, 20, 30, and 40 mmol/L; D) for 48 hours. Eventually, cells were subjected to thecytofluorometric assessment of apoptosis-related parameters. Representative dot plots and quantitative data are shown in A and B–D, respectively. In A,numbers refer to thepercentageof cells in the correspondingquadrant. InB–D,white andblackcolumns illustrate thepercentage of dying [DiOC6(3)
lowPI�] anddead (PIþ) cells, respectively (means � SEM, n ¼ 3). E and F, WT and CDDP-resistant R1 A549 cells were exposed to 1 mmol/L CEP, 5 mmol/L PJ, 1 mmol/LCDDP (WT cells), or 5 mmol/L CDDP (R1 cells) for 48 hours and then allowed to generate colonies in drug-free medium for 10 days. In E, representativepictures of colonies as observed upon crystal violet staining are shown (scale bar, 1 cm). In F, columns depict the normalized surviving fraction (means�SEM,n¼ 3 parallel wells). G and H,WT and CDDP-resistant R1 A549 cells were left untransfected (–) or transfected with two distinct control siRNAs (UNR, EMR) orwith PARP-specific siRNAs for 24 hours, then subjected to the cytofluorometric assessment of apoptosis-related variables. In G, white and black columnsdepict the percentage of DiOC6(3)
lowPI� and PIþ cells, respectively (means � SEM, n ¼ 3). In H, representative immunoblots confirming PARP1downregulation are reported. GAPDH levels weremonitored to ensure equal loading of lanes. �,P < 0.05; ��,P < 0.01; ���,P < 0.001 (Student t test), comparedwith equally treated WT cells (B–D) or UNR-transfected cells of the same type (G). See also Supplementary Fig. S1.
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the context of CDDP resistance, WT A549 cells and all theirCDDP-resistant derivatives were exposed to increasing dosesof CEP or PJ for 48 hours, followed by the cytofluorometricevaluation of cell death. Unexpectedly, we found that someCDDP-resistant clones (R1, R2, R3, R6, R7) were more sensitivethan parental A549 cells to both PARP inhibitors used in thisstudy (CEP and PJ), although a few others were not (R4, R5) orexhibited intermediate responses (R8, R9). Thus, although 10mmol/L CEP and 30 mmol/L PJ exerted negligible cytotoxiceffects against WT A549 cells, they did induce significantextents of cell death in several of their CDDP-resistant clonalcounterparts (Fig. 2C and D and Supplementary Fig. S1).Similar results were obtained in clonogenic assays. Thus,
although the concentration of CDDP that halved the clonoge-nic potential of parental A549 cells (IC50) was approximately1 mmol/L, the IC50 of the resistant clone R1 was approximately5mmol/L. Conversely, the clonogenic potential ofWTA549 cellswas barely affectedby exposure to 1mmol/LCEPor 5mmol/L PJ,although that of CDDP-resistant R1 cells was significantlycompromised (Fig. 2E and F). In line with these observations,CDDP-resistant A549 cell clones that were sensitive to thecytotoxic activity of PARP inhibitors (such as R1) also died inresponse to PARP1-targeting siRNAs (Fig. 2G and H).Altogether, these results indicate that CDDP resistance
often, but not always, entails a hypersensitivity to PARPinhibition.
PARP hyperactivation predicts the sensitivity of cancercells to PARP inhibitorsTo characterize the molecular cascades through which
PARP inhibitors kill CDDP-resistant NSCLC cells, we firstdetermined the expression level and activation status ofPARP1. This was achieved by the immunoblotting-assisteddetection of PARP1 and of the products of its enzymaticactivity, that is, poly(ADP-ribose) (PAR)-containing proteins.As compared with their CDDP-resistant counterparts, WTA549 cells in steady-state conditions exhibited low levels ofboth PARP1 andPAR-containing proteins. The amount of PAR-containing proteins increased (at least to some extent) in WTcells upon addition of CDDP, a phenomenon that could beentirely suppressed by the administration of CEP or PJ. Con-versely, the constitutively high levels of PAR-containing pro-teins observed in CDDP-resistant clones could not be furtherelevated by CDDP, yet were fully suppressed by CEP or PJ(Supplementary Fig. S2). Of note, CDDP-resistant clones thatwere particularly sensitive to the cytotoxic effects of PARPinhibitors were characterized by higher levels of PAR-contain-ing proteins than clones that responded less efficiently to CEPand PJ (Fig. 3A). These results indicate a correlation betweenPARP1 enzymatic activity and the lethal effects of PARPinhibitors in CDDP-insensitive NSCLC clones.To assess whether the intracellular levels of PAR-containing
proteins might predict the susceptibility of cancer cells ofdifferent histologic origin to PARP inhibitors, we explored thisparameter in 3 different NSCLC (H1650, H460, H1299), 1mesothelioma (P31), 1 ovarian (TOV-112D), and 1 cervicalcancer (HeLa) cell lines, invariably comparing parental cellswith their CDDP-resistant derivatives. In 5 out of 6 such
pairwise comparisons, CDDP-resistant cells exhibitedincreased PAR levels. Only in H1299 cells the amount ofPAR-containing proteins was reduced along with the develop-ment of CDDP resistance (Fig. 3B). Importantly, all cell linescharacterized by elevated levels of PAR proteins (PARhigh),were more sensitive to CEP and PJ than their parental PARlow
counterparts (Fig. 3C and Supplementary Table S1). Theseresults underscore a tight link between high intracellular levelsof PAR-containing proteins (and hence an elevated enzymaticactivity of PARPs) and the susceptibility of cancer cells tosuccumb to PARP inhibitors. Of note, although PARhigh cellstend to express high levels of PARP1 as well, we observed noabsolute correlation between these 2 variables (for instance,see P31 and TOV-112D cells, Fig. 3B), indicating that PAR levelspredict the sensitivity of cells to PARP inhibition more accu-rately than the expression levels of PARP1.
Mechanism of cell death induced by PARP inhibitorsCDDP-resistant A549 cells succumbed to PARP inhibitors via
the intrinsic pathway of apoptosis, as indicated by the mito-chondrial release of cytochrome c (assessed by immunofluo-rescence as the redistribution of cytochrome c from a com-partmentalized to a diffuse cytoplasmic localization) andcaspase-3 activation (detected by immunostaining with anantibody specific for the cleaved, proteolytically active form ofcaspase-3; ref. 40; Fig. 4A). In line with this notion, theco-administration of the broad-spectrum caspase inhibitorZ-VAD-fmk blunted the cytotoxic effects of CEP or PJ (Fig.4B). Moreover, CDDP-resistant, but notWT, A549 cells respond-ing to CEP or PJ manifested (at least to some extent) theapoptosis-associated cleavage of PARP1 (41; SupplementaryFig. S2). In response to PARP inhibitors, CDDP-resistant cellsalso mounted a DNA damage response characterized by anaccumulation of nuclear foci containing histone 2AX phospho-rylated on serine 139 (gH2AX) that was far more pronouncedthan that of CDDP-sensitive cells (Fig. 4C). In addition, thepercentage of cells exhibiting gH2AXþ foci in steady-stateconditions was significantly higher among CDDP-resistant cellsthan among their parental counterparts (Fig. 4C). The DNAdamage response elicited by PARP inhibitors was detectable asearly as 6 hours after exposure and could not be suppressed bythe co-administration of Z-VAD-fmk (Supplementary Fig. S3).Of note, DNA repair via HR appeared to be functional in bothparental cells and their CDDP-resistant counterparts, asreflected by the formation of BRCA1þ and RAD51þ nuclearfoci formation in response to g irradiation (Fig. 4D) or treatmentwith PARP inhibitors (Supplementary Fig. S4).
As PARP1 is critically involved in the BER pathway, wecomparatively examined the proficiency of this DNA repairsystem in parental and CDDP-resistant A549 cells. To this aim,we tested the capacity of cell-free extracts to catalyze thecleavage of synthetic DNA substrates in assays that reflect thefirst steps in the BER cascade, as catalyzed by apurinic/apyrimidinic endonuclease 1 (APE1), alkyl-N-purine DNA gly-cosylase, human uracil-DNA glycosylase, human endonucleaseIII, and human 8-oxoguanine glycosylase 1 (42). We found thatWT and CDDP-resistant A549 cells possess similar repairactivities relative to specific DNA substrates (Supplementary
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Figs. S5, S6, and S7), suggesting that the upstream steps of BERare not impaired in these cellular models. SSBs as generated bycombined action of DNA glycosylases and APE1 are normallyrepaired by proteins that operate in downstream steps of BERsuch as X-ray repair cross-complementing protein 1 (XRCC1),PARP1, and polymerase b (POLb; ref. 42). Remarkably, various
(but not all) CDDP-resistant cells characterized by PARP1overexpression and high levels of PAR-containing proteinsexhibited reduced expression levels of XRCC1 and POLb(i.e., A549 R6 to R9, H1650, H460, P31, HeLa). Accordingly,several among these clones were more sensitive to temozolo-mide (a DNA damaging agent reputed to selectively kill
Figure 3. PARP activity predicts response to PARP inhibitors. A, WT and CDDP-resistant (R1–R9) A549 cells were processed for the immunoblotting-assisted detection of PAR-containing proteins and PARP1 expression levels. Representative immunoblots are reported and the abundance of PAR-containing proteins is correlated with responsiveness to PARP inhibitors. Bars illustrate the levels or PAR-containing proteins as quantified by densitometryupon immunoblotting (means�SEM, n¼ 3). Sensitivity to PARP inhibitors (indicated byþ and� symbols) was determined based on the experiments shownin Fig. 2 and Supplementary Fig. S1. Results are representative of 3 independent determinations. B, immunoblotting-based assessment of PARP1 andPAR-containing proteins in WT and CDDP-resistant (R) H1650, H460, P31, TOV-112D, HeLa, and H1299 cells. In A and B, GAPDH levels were monitoredto ensure equal loading of lanes. C, parental and CDDP-resistant H1650, H460, P31, TOV-112D, HeLa, and H1299 cells were kept in control conditions orincubated with the indicated concentration of CDDP (5, 10, 20, 50, and 80 mmol/L), CEP (5, 10, 15, 20, and 40 mmol/L), or PJ (10, 20, 30, 40, and80 mmol/L) for 48 hours, followed by the cytofluorometric assessment of apoptosis-related parameters. �, P < 0.05 (Student t test), compared with equallytreated WT cells of the same type. See also Supplementary Table S1.
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BER-deficient cells; ref. 43) than their CDDP-sensitive counter-parts (Supplementary Fig. S8).Taken together, these results indicate that CDDP-resis-
tant cells that are characterized by high PARP1 expressionlevels and high amounts of PAR-containing proteins executethe early steps of BER as proficiently as their WT counter-parts, but exhibit some degree of deficiency in downstreamreactions such as those catalyzed by XRCC1 and POLb. Inaddition, our results indicate that PARP activity mayincrease in CDDP-resistant cells in the absence of obviousHR defects.
Effects of PARP inhibition on CDDP-resistant tumorxenograftsTo validate the hypothesis that PARP-hyperactivating,
CDDP-resistant NSCLCs might respond to PARP inhibitors invivo, athymic nu/nu mice carrying parental (WT) or CDDP-resistant cell-derived xenografts were treated with PJ. Incontrast to tumors developing from WT A549 cells, whichfailed to respond to PJ monotherapy, the growth of cancersgenerated by CDDP-resistant cells, which exhibit high levels of
PAR-containing proteins (Fig. 3A), was significantly delayed bythe administration of PJ as a standalone intervention (Fig. 5A).Thus, the levels of PAR-containing proteins may be used as abiomarker to estimate the effects of PARP inhibitors in vivo. Toget further insights into this issue, we developed an immuno-histochemical-staining method that specifically detects PAR-containing proteins on paraffin-embedded cell pellets andtissue sections (as shown by the fact that the siRNA-mediateddepletion of PARP1 as well as the treatment with PARPinhibitors result in a complete loss of the signal, Fig. 5B). Inthe absence of chemotherapy, tumors derived from CDDP-resistant R6 cells were characterized by higher levels of PAR-containing proteins thanWTA549 cell-derived xenografts (Fig.5C), indicating that the levels of PAR-containing proteins arepreserved during tumor formation in vivo. Moreover, xeno-grafts featuring high amounts of PAR-containing proteins andexposed to PJ in vivo underwent a consistent reduction of poly(ADP-ribosyl)ation-dependent immunoreactivity (Fig. 5C).Hence, PARP inhibitors appear to exert antineoplastic effectsin vivo along with a significant decrease in PARP enzymaticactivity.
Figure 4. Mechanisms of cell death induction by PARP inhibitors in HR-proficient CDDP-resistant cancer cells. A and B, WT human NSCLC A549 cells andtheir CDDP-resistant derivatives (R1 and R6) were cultured in control conditions or exposed to 10 mmol/L CEP or 30 mmol/L PJ, alone or in combination with 50mmol/L Z-VAD-fmk (Z-VAD), then processed either for the simultaneous immunofluorescence detection of cytochrome c (Cyt c) release and the active form ofcaspase-3 (Casp-3a), 24 hours later, or for the cytofluorometric determination of apoptosis-related parameters, 48 hours later. In A, representativemicrophotographs are depicted (scale bar, 10 mm) and the frequency of cells showing diffuse cytochrome c staining (indicative of mitochondrial outermembrane permeabilization) and caspase-3 activation is reported (means � SEM, n ¼ 3). �, P < 0.05; ��, P < 0.01 (Student t test), compared with untreatedcells of the same type. In B, white and black columns illustrate the percentage of dying [DiOC6(3)
lowPI�] and dead (PIþ) cells, respectively (means � SEM,n ¼ 3). �, P < 0.05 (Student t test), compared with cells of the same type not receiving Z-VAD. C, WT and CDDP-resistant (R1, R6) A549 cells were cultured onglass coverslips in control conditions or in the presence of 10 mmol/L CEP or 30 mmol/L PJ for 6 hours, then processed for the immunofluorescence detectionof phosphorylated histone 2AX (gH2AX, green fluorescence). Representative immunofluorescence microscopy images (scale bar, 10 mm) and quantitative dataare depicted. Columns illustrate the percentage of cells whose nucleus contained more than 5 gH2AXþ foci (means � SEM, n ¼ 300 cells in triplicateexperiments). D, WT and CDDP-resistant (R) NSCLC A549, H460, and H1650 cells were maintained in control conditions or exposed to g rays (6 Gy), andthen were processed for the immunofluorescence-assisted detection of BRCA1 and RAD51. Representative immunofluorescence microscopy images ofirradiated (WT and R) H460 cells (scale bar, 5 mm) and quantitative data are reported. Columns illustrate the percentage of cells whose nucleus contained morethan 5 BRCA1þ or RAD51þfoci (means � SEM, n ¼ 300 cells in triplicate experiments). �, P < 0.05; ��, P < 0.01 (Student t test), compared with equally treatedWT cells (C) or untreated (D) cells. In C and D, Hoechst 33342 (H33342, blue fluorescence) was used for nuclear counterstaining. See also SupplementaryFigs. S3–S8.
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DiscussionHere, we show that NSCLC cell clones that have been
selected for CDDP resistance by prolonged exposure to CDDPcan upregulate PARP1 and hence accumulate elevated levels ofPAR-containing proteins. Both high levels of PAR-containingproteins and CDDP resistance were maintained upon thewithdrawal of CDDP, suggesting the existence of yet elusive,genetic or epigenetic mechanisms that stabilize the upregula-tion of PARP1 and its hyperactivation. Irrespective of theseunresolved issues, the hyperactivation of PARP appears as arelatively frequent event in the context of acquired CDDPresistance, as it could be observed in more than half ofCDDP-resistant cells used in this study. As a caveat, thehyperactivation of PARP is not a universal characteristic ofCDDP resistance but rather constitutes an optional mecha-nism. Indeed, one of the CDDP-resistant cell lines (H1299)characterized in this study exhibited a combined reduction inPARP1 expression levels and in the abundance of intracellularPAR-containing proteins.
The most striking observation reported in the present studyconcerns the ability of PARP inhibitors to kill CDDP-resistant
cells that exhibit constitutive PARP hyperactivation. It is there-fore tempting to speculate that this trait—which can be mon-itored by the abundance of PAR-containing proteins—mayreflect a situation of PARP-dependent survival. Inhibition ofPARP in PARP-hyperactivating cells led indeed to the activationof a DNA damage response (as indicated by the appearance ofgH2AXþ foci) and ignited the intrinsic pathway of apoptosis.Thus, not only oncogenesis but also the development of che-moresistance appears to be coupled to the constitutive activa-tion of stress response mechanisms that may constitute pref-erential targets for chemotherapy or chemosensitization (44).
The mechanisms through which CDDP-resistant cells upre-gulate PARP1 expression and its enzymatic activity remainlargely obscure. Actually, we found that CDDP-resistant cellscanbe categorized intodifferent classes, those that overexpressPARP1 and contain high levels of PAR-containing proteins andthose that do not. This classification can be correlated withother parameters, including (but perhaps not limited to)XRCC1 and POLb expression levels as well as the cytotoxicresponse to theDNA-damaging agent temozolomide. Althoughthese results are insufficient to establish direct cause–effect
Figure 5. Therapeutic effects ofPARP inhibition against CDDP-resistant, PARP-hyperactivatingxenografts in vivo. A, WT humanNSCLC A549 cells and theirCDDP-resistant derivatives (R6)were subcutaneously xenograftedin athymic nu/nu mice (15 pergroup) and treatedwith 5mg/KgPJ(intraperitoneally 3 times aweek) oran equivalent volume of vehicle(PBS), starting from day 8postinoculation. Tumor growthwas routinely monitored with astandard caliper and is reported asmeans� SEM. �, P < 0.05 (Studentt test), as compared with vehicle-treated mice. B, CDDP-resistantR6 cells were cultured in theabsence or in the presence of10 mmol/L CEP or—alternatively—transfected with a PARP-specificsiRNA, then processed for theimmunocytochemical detection ofPAR-containing proteins.Representative microphotographs(scale bar, 30 mm) and quantitativedata (means � SEM, n ¼ 3) areshown. C, tumors generated byWTand R6 A549 cells and treated invivo as in A were recovered 2 daysafter the last treatment, and thelevels of PAR-containingproteins were assessed byimmunohistochemistry.Representative microphotographs(scale bar, 10 mm) and quantitativedata are reported (means � SEM,n ¼ 3).
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relationships, they suggest a functional connection betweendefects in the downstream steps of the DNA repair via the BERpathway and upregulation/hyperactivation of PARP1.CDDP remains one of the first-line agents for the therapy
for NSCLC, in particular when such tumors lack activatingmutations of the EGF receptor (45). However, NSCLCpatients treated with CDDP often develop chemoresistance,near-to-invariably leading to relapse and therapeutic failure(46). Here we show that CDDP-resistant cells respond moreefficiently than their WT counterparts to PARP inhibitors. Ifthe results of in vitro studies could be extrapolated to theclinics, PARP inhibitors would be most beneficial for (at leasta fraction of) NSCLC patients relapsing after CDDP-basedchemotherapy. In this context, it will be crucial to under-stand not only to which extent and under which specificcircumstances PARP inhibitors may be effective as stand-alone interventions, but also whether the levels of PARproteins may be used in clinical settings to predict theresponsiveness of cancer patients to PARP inhibitor-basedchemotherapeutic regimens.
Disclosure of Potential Conflicts of InterestNo potential conflicts of interest were disclosed.
Authors' ContributionsConception and design: J. Michels, L. Galluzzi, K.A. Olaussen, J.-C. Soria, M.Castedo, G. KroemerDevelopment of methodology: J. Michels, J. Adam, K.A. Olaussen, O. Kepp, M.Talbot, A. Robin
Acquisition of data (provided animals, acquired and managed patients,provided facilities, etc.): J. Michels, I. Vitale, J. Adam, O. Kepp, L. Senovilla, I.Talhaoui, J. Guegan, P. Girard, C. Or�ear, D. Lissa, A.Q. Sukkurwala, P. Behnam-Motlagh, G.S. Wu, C. Brenner, M. Saparbaev, J.-C. SoriaAnalysis and interpretation of data (e.g., statistical analysis, biostatistics,computational analysis): J. Michels, I. Vitale, I. Talhaoui, J. Guegan, D.P. Enot,P. Garcia, P. Dessen, M. Saparbaev, J.-C. Soria, M. CastedoWriting, review, and/or revision of the manuscript: J. Michels, I. Vitale, L.Galluzzi, O. Kepp, D.P. Enot, P. Girard, P. Behnam-Motlagh, C. Brenner, J.-C. Soria,M. Castedo, G. KroemerAdministrative, technical, or material support (i.e., reporting or orga-nizing data, constructing databases): J. Michels, J. Adam, K. KohnoStudy supervision: J. Michels, L. Galluzzi, M. Castedo, G. Kroemer
AcknowledgmentsThe authorswould like to thank Louis Kayitalire fromCephalon (Frazer, USA)
and Genomic Core Facilities-TA2012.
Grant SupportG. Kroemer is supported by the European Commission (ArtForce); Agence
National de la Recherche (ANR); Ligue contre le Cancer (Equipe labellis�ee);Fondation pour la Recherche M�edicale (FRM); Institut National du Cancer(INCa); LabEx Immuno-Oncologie; Fondation de France; Fondation Betten-court-Schueller; AXA Chair for Longevity Research; Canc�eropole Ile-de-Franceand Paris Alliance of Cancer Research Institutes (PACRI). J. Michels is supportedby the French Ministry of Science, FRM, Groupe Pasteur Mutualit�e, and ActionLions "Vaincre le Cancer" (Luxembourg). L. Senovilla, I. Vitale, and L. Galluzzi aresupported byFRM, the LigueNationale contre le Cancer, and the LabEx Immuno-Oncologie, respectively. I. Talhaoui is financed by postdoctoral fellowships fromFondation ARC pour la Recherche sur le Cancer.
The costs of publication of this article were defrayed in part by the payment ofpage charges. This article must therefore be hereby marked advertisement inaccordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received July 30, 2012; revised January 5, 2013; accepted January 7, 2013;published online April 3, 2013.
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