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Originally published In Press as doi:10.1074/jbc.M606470200 on September 8, 2006

J. Biol. Chem., Vol. 281, Issue 45, 34394-34405, November 10, 2006
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MNNG-induced Cell Death Is Controlled by Interactions between PARP-1, Poly(ADP-ribose) Glycohydrolase, and XRCC1*

Claudia Keil, Tina Gröbe, and Shiao Li Oei1

From the Institut für Biochemie, Freie Universität Berlin, Thielallee 63, 14195 Berlin, Federal Republic of Germany

Received for publication, July 7, 2006 , and in revised form, August 25, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
PARP-1 (poly(ADP-ribose) polymerases) modifies proteins with poly(ADP-ribose), which is an important signal for genomic stability. ADP-ribose polymers also mediate cell death and are degraded by poly(ADP-ribose) glycohydrolase (PARG). Here we show that the catalytic domain of PARG interacts with the automodification domain of PARP-1. Furthermore, PARG can directly down-regulate PARP-1 activity. PARG also interacts with XRCC1, a DNA repair factor that is recruited by DNA damage-activated PARP-1. We investigated the role of XRCC1 in cell death after treatment with supralethal doses of the alkylating agent MNNG. Only in XRCC1-proficient cells MNNG induced a considerable accumulation of poly(ADP-ribose). Similarly, extracts of XRCC1-deficient cells produced large ADP-ribose polymers if supplemented with XRCC1. Consequently, MNNG triggered in XRCC1-proficient cells the translocation of the apoptosis inducing factor from mitochondria to the nucleus followed by caspase-independent cell death. In XRCC1-deficient cells, the same MNNG treatment caused non-apoptotic cell death without accumulation of poly(ADP-ribose). Thus, XRCC1 seems to be involved in regulating a poly(ADP-ribose)-mediated apoptotic cell death.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Poly(ADP-ribosylation) of proteins is involved in the regulation of basal cellular processes and seems to be crucial for genomic integrity and cell survival. Responsible for the synthesis of poly(ADP-ribose) (PAR)2 are poly(ADP-ribose) polymerases (PARPs) (1). The most abundant and active PARP enzyme is PARP-1, a predominantly nuclear protein of 113 kDa. PARP-1 rapidly binds to DNA breaks, is thereby activated, and covalently automodifies itself under consumption of NAD+. To a lesser extent, some other nuclear proteins are also modified with PAR polymers (2). The primary structure of PARP-1 is well conserved between species. The N-terminal DNA binding domain, which contains two zinc finger motifs, is linked to a nuclear localization signal, the main acceptor sites of automodification are located within the central domain, and the 55-kDa C-terminal domain of the enzyme contains the catalytic site (1). PARP-1 appears to physically interact with multiple proteins involved in DNA metabolism, such as histones, transcription factors, replication factors, and DNA repair enzymes (3). Among DNA repair proteins, PARP-1 interacts with x-ray repair cross-complementing protein 1 (XRCC1) (4). A mutant line of Chinese hamster ovary (CHO) cells, which displays hypersensitivity to a broad range of genotoxins was established and termed EM9 cells (5). It turned out that EM9 cells have a reduced ability to rapidly repair DNA single-strand breaks and are genetically unstable as a consequence of XRCC1 deficiency. Different analyses revealed that XRCC1 physically interacts with several DNA repair enzymes, thereby regulating their corresponding activities (6). XRCC1, a polypeptide of 70 kDa, contains two breast cancer C-terminal domains and a nuclear localization signal but is lacking any known enzymatic activity (6). Notably, recruitment of XRCC1 to single-strand breaks strictly depends on PARP-1 activity (7, 8). PAR polymers are synthesized in response to DNA breaks, which can arise directly or indirectly, for example, after treatment with alkylating agents. Produced PAR modifications are rapidly degraded by poly(ADP-ribose) glycohydrolase (PARG), which cleaves the polymers with high specificity at the glycosidic bonds, generating free ADP-ribose. PARG is the physiological counterpart for all PARP enzymes, encoded by a unique gene (9). Human PARG, encoded by 18 exons is 110 kDa in size (10) and the catalytic domain resides in the C-terminal part of the enzyme (11, 12). Interestingly, recently a 39-kDa protein termed ARH3 has been isolated (13), which possesses a glycohydrolase activity although is structurally unrelated to the conventional 110-kDa PARG. In addition, several PARG isoforms with different sizes, resulting either from alternative initiation events or from post-translational proteolysis, have been described in mammalians (14). Human PARG contains several putative localization signals: nuclear localization signal, nuclear export signals, and a mitochondrial localization signal (15, 16). Whereas PARG activity is detected predominantly in the cytoplasm, full-length PARG is localized to the nucleus (17, 18). Knockout of the full-length isoform of PARG in mice resulted in an increased sensitivity to genotoxic and endotoxic stress (19) and the loss of PARG activity in Drosophila melanogaster caused progressive neurodegeneration (20). Finally, it was demonstrated that after complete abrogation of PARG expression, murine embryonic cells were only viable in the presence of PARP inhibitors. After withdrawal of these inhibitors an accumulation of PAR polymers was observed and cells underwent apoptosis (21). Thus, the metabolism of PAR plays a fundamental role for the decision of the cell to survive or die (22).

Nevertheless, the induction of PARP-1 activity by irreparable amounts of DNA breaks can deplete the cell of NAD and ATP, finally leading to cell death (23). For example, PARP-1-dependent necrosis can be triggered by treatment with 1 mM H2O2 (24). Yu et al. (25) described yet another cell death program depending on PARP-1 activity. In response to an exposure to 0.5 mM N-methyl-N'nitro-N-nitrosoguanidine (MNNG), PAR polymers accumulate, instantly provoking the translocation of apoptosis inducing factor (AIF) from mitochondria to the nucleus. This death stimulus then induces nuclear shrinkage and finally caspase-independent cell death (25, 22). How an accumulation of PAR is accomplished, whether by overactivation of PARP-1 or by repression of PARG activity, is not known. Therefore, the analysis of the relationship between PARP-1 and PARG is necessary to unravel the role of PAR metabolism in cell death.

Previously we reported that PARG interacts with human PARP-1 from HeLa cell extracts (26). In line with our findings, an affinity of PARG for PARP-1 was shown in a recent proteomic approach (27). Here we characterize the functional relationship between PARP-1 and PARG. Both enzymes interact directly and PARG has the ability to modulate PARP-1 activity. In addition, PARG also interacts with XRCC1. Above all, we provide evidence that the interplay between PARP-1, PARG, and XRCC1 regulates apoptotic cell death induced by supralethal MNNG doses.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cells and Extracts—HeLa S3 cells (ATCC) were grown in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% fetal calf serum (Biochrom, Berlin, Germany). CHO cell lines AA8, EM9 (28), EM9-V, and EM9-XH (29) were kindly provided by Dr. K. W. Caldecott and cultured in {alpha}-minimal essential medium (Invitrogen) as described by Loizou et al. (30). Nuclear extracts were prepared by the method of Schreiber et al. (31).

Cloning and Purification of Proteins—The cDNA encoding amino acids 378-976 of human PARG (accession number DQ867088 [GenBank] ) was cloned as a SalI fragment into the vector pGEX-5X-3 (Amersham Biosciences). Expression of glutathione S-transferase (GST) or the GST-PARG65 construct was performed in Escherichia coli BL21 Codon Plus-RIL cells (Stratagene), which additionally were transfected with the pREP4 plasmid (Qiagen). Cells were cultured in LB broth overnight at 25 °C and overexpression was induced by adding 0.1 mM isopropyl 1-thio-beta-D-galactopyranoside for 2 h. GST-PARG65 was bound to glutathione-Sepharose (Amersham Biosciences) in phosphate-buffered saline (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4 (7 H2O), 1.4 mM KH2PO4), supplemented with 1% Triton X-100, 5 mM dithiothreitol. After washing, proteins were eluted with phosphate-buffered saline supplemented with 2% N-octyl-beta-D-glucoside, 40 mM glutathione, pH 8.0. Eluted GST-PARG65 was dialyzed in phosphate-buffered saline supplemented with 1 mM dithiothreitol. XRCC1, Lig III, and PARP-1 constructs were expressed and purified as described before (32, 33).

PAR Metabolism—[{alpha}-32P]NAD+ was obtained from Amersham Biosciences and gallotannins were from Sigma. Poly-(ADP-ribosylation) activities were determined as described previously (26) with minor modifications. Purified recombinant proteins as specified in the legends to the figures or 20 µg of nuclear proteins were incubated in 25 µl of phosphate buffer (50 mM potassium phosphate, pH 7.2, 200 µM EDTA, 10 mM beta-mercaptoethanol, 100 µg/ml bovine serum albumin) with 10 µM [{alpha}-32P]NAD+ and 10 µg/ml nicked DNA at 30 °C for 20 min. All reactions were performed in the presence of 200 µM EDTA and absence of Mg2+ ions to reduce endogenous phosphodiesterase and/or ADP-ribose pyrophosphatase activities that convert PAR and ADPR to AMP (34). Reactions were stopped by trichloroacetic acid precipitation and incorporation of [32P]PAR was determined. For determination of relative PARG activity [32P]PAR was synthesized in vitro as described earlier (33) and incubated with purified proteins or nuclear extracts (20 µg of proteins per 30 µl reactions) in phosphate buffer for 30 min at 30 °C. Reactions were stopped by precipitation with acetone. Precipitated nucleotides were dissolved in TE buffer. Samples containing equal amounts of radioactivity were applied to cellulose-coated plates. Thin layer chromatography was performed using the solvent system 0.3 M LiCl, 1 M acetic acid. After separation, dried cellulose plates were subjected to autoradiography and quantified using a phosphorimager or by Cerenkov counting of excised thin layer slices. Protein modifications with [{alpha}-32P]ADP-ribose were analyzed by PAGE and autoradiography. Alternatively, reactions were stopped by precipitation with acetone and nucleotides were separated by thin layer chromatography.

Antibodies and Immunostaining Analyses—Antibodies against PARG were raised in rabbits immunized with the purified His-tagged catalytic domain of PARG (26). These PARG antibodies recognize the catalytic 65-kDa PARG fragment and to a lesser extent also the human full-length PARG. Anti-pentahistidine antibodies were obtained from Qiagen. Antibodies directed against XRCC1 (H-300, from rabbit), AIF (H-300, from rabbit), GST (Z-5, from rabbit), PARP-1 (A-20, from goat), and YY 1 (C-20, from rabbit) were from Santa Cruz. Monoclonal {alpha}-tubulin antibodies (DM1A, from mouse) were from Sigma (Taufkirchen, Germany), polyclonal anti-PAR antibodies were from Alexis (96-10-04 from rabbit). For immunofluorescence analyses, cells were fixed with 3% formaldehyde, 0.25% Triton X-100, and antibodies against PAR or AIF were used and visualized using Alexa Fluor-conjugated secondary antibodies (Invitrogen). Nuclei were counterstained with DAPI (3 µM, Invitrogen). Necrotic cells were stained by incubating unfixed cells with 100 µg/ml propidium iodide (Molecular Probes) in BBS (3.1 mM KCl, 134 mM NaCl, 1.2 mM CaCl2, 1.2 mM MgSO4, 0.25 mM KH2PO4, 15.7 mM NaHCO3, 2 mM glucose, pH 7.2) for 30 min at 37 °C and analyzed immediately using a fluorescence microscope. Photomicrographs were obtained at room temperature with a microscope (Leica, Wetzlar, Germany) equipped with a digital camera. The relative amount of PAR accumulation was quantified using imaging software (Matrix Vision, Oppenweiler, Germany). Localization of AIF in cells and nuclear shrinkage of cells were estimated by visual inspection. At least 300 cells were counted for each sample and all experiments were repeated three times.

Affinity Precipitation and Pulldown Assays—For GST pull-down experiments, GST or GST-PARG65 (7.5 µg) together with recombinant PARP-1 or XRCC1 constructs (20 µg) were incubated with glutathione-Sepharose (50 µl) in 0.5 ml of BP (10 mM Tris/HCl, 7 mM MgCl2, 150 mM NaCl, 50 µM ZnCl2, 0.05% (v/v) Nonidet P-40, 1 mM dithiothreitol, pH 8.0) by head over head rotation at 4 °C for 45 min. After 5 washing steps with BP, bound proteins were extracted with SDS gel loading buffer and subjected to Western blot analysis. Precipitation of XRCC1-bound proteins from EM9-XH cells was performed as described by Caldecott et al. (35). For immunoprecipitation, 40 µl of nuclear extracts from HeLa S3 cells (75 µg of protein) were diluted with 120 µl of immunoprecipitation buffer (20 mM HEPES, 1 mM EGTA, 1 mM EDTA, 1 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, pH 7.5) and pre-cleared by incubation with 50 µl of protein A-Sepharose (Sigma), covalently coupled with control goat antibodies, at 4 °C for 30 min. Thereafter, the unbound fraction was incubated with 10 µg of goat anti-PARP-1 antibodies (PARP-1 (A-20), Santa Cruz) covalently coupled to 35 µl of protein A-Sepharose at 4 °C for 30 min. The Sepharose beads were washed three times with immunoprecipitation buffer including 100 mM NaCl, and bound proteins were extracted subsequently with SDS gel loading buffer and subjected to Western blot analysis.

Yeast Two-hybrid Analysis—Human full-length PARG was cloned into the two-hybrid vector pGADT7 (Clontech). pAS-XRCC1 (36) was kindly provided by K. W. Caldecott. Plasmids were transformed into the yeast strain PJ69-4A (37) and diploids were selected using synthetic medium lacking leucine and tryptophan. For validation of protein-protein interaction, colonies were transferred to histidine- or adenine-free medium.

Treatment of CHO Cells—Stock solutions of MNNG, 3-aminobenzamide (3-ABA), staurosporine, and Z-VAD(OMe)-fmk (Bachem, Weil am Rhein, Germany) were dissolved in Me2SO and serially diluted with BBS or growth medium immediately before use. Nonconfluent CHO cells were treated with 0.5 mM MNNG in BBS for 10 min. Then cells were washed with medium and incubated in fresh medium at 37 °C for time periods as indicated in the figure legends. Perchloric acid extracts were prepared from CHO cells and the cellular ATP level was determined as described earlier (32) using the luciferase assay. The cellular NAD level was determined from perchloric acid extracts as described by Jacobson and Jacobson (38).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
PARG Interacts with PARP-1—Previously, we demonstrated an interaction between full-length PARG immobilized to tannin-Sepharose and endogenous PARP-1 from HeLa cell extracts (26). We also detected interaction of PARP-1 with the immobilized 65-kDa C-terminal fragment of PARG (amino acids 378-976), in the following termed PARG65, which contains the catalytic activity (data not shown). To further characterize the interaction between PARP-1 and PARG we performed GST pulldown experiments with purified recombinant constructs (Fig. 1A). We expressed catalytically active GST-PARG65 in E. coli, purified it, and incubated it with His-tagged full-length PARP-1. Fractions from GST pull-down experiments were separated by SDS-PAGE and subjected to Western blot analysis using anti-polyhistidine and anti-GST antibodies, respectively (Fig. 1A). As evident, after GST pulldown full-length PARP-1 was detected only in fractions eluted from GST-PARG65 beads (Fig. 1A, uppermost panels, fourth lane). If His-tagged constructs of isolated domains of PARP (33) were applied, only the automodification domain showed an affinity to GST-PARG65, whereas no interaction was observed with the other PARP domains (Fig. 1A). These experiments were performed in the absence of NAD+ or PAR, thus demonstrating a direct protein-protein interaction between catalytically active PARG65 and the automodification domain of PARP-1. Next, purified full-length PARP-1 and GST-PARG65 were incubated as described before and the elution fraction from the GST pull-down was analyzed for PARP activity by incubation with [{alpha}-32P]NAD and nicked DNA. Eluted PARP-1 was catalytically active (Fig. 1B, second lane), indicating that native PARP-1 bound to the GST-PARG65 beads. Menard et al. (39) reported that PARG was no acceptor for PAR modifications in vitro, when PARP-1, PARG, histones, DNA, and NAD were incubated together at physiological ratios. Similarly, we observed that the predominant reaction in elution fractions from pull-down with GST-PARG65 was automodification of PARP-1, whereas [32P]ADP-ribosylation of PARG65 was only detectable if GST-PARG65 was added in excess (data not shown).

To study the impact of the interaction between PARP-1 and PARG in vivo, we performed co-immunoprecipitation experiments. First, Western blot analyses showed that PAR modifications were virtually absent in extracts of untreated HeLa cells (Fig. 1C). PARG65 was detected after co-imunoprecipitation of these extracts with anti-PARP-1 antibodies (Fig. 1D). Thus, a preformed complex of PARG and PARP-1 might exist in the nucleus of HeLa cells, independent of PARP-1 activation. To further study the interplay of these enzymes, we supplemented nuclear extracts of HeLa cells with increasing amounts of recombinant GST-PARG65 and incubated the extracts with [{alpha}-32P]NAD and nicked DNA. After reaction for 20 min, modifications with [32P]poly(ADP-ribose) were visualized in SDS-PAGE (Fig. 2B). Amounts of released [32P]ADP-ribose were determined by evaluation of thin layer chromatograms (Fig. 2A), and polymer chain lengths were analyzed by gel electrophoresis (Fig. 2C). As expected, the addition of increasing amounts of catalytically active GST-PARG65 enhanced the levels of released [32P]ADP-ribose (Fig. 2A, bars 1-4), whereas the sizes of the produced [32P]PAR polymers were decreased (Fig. 2C, lanes 1-4). Previously we presented evidence that tannins elevate the level of PAR in HeLa cell extracts by inhibition of PARG (26). In the presence of nuclear proteins 150 µM tannins had no modulating influence on PARP-1 activity (26). If similar experiments as shown in Fig. 2, bars/lanes 1-4, were performed in the presence of 150 µM tannins, the levels of [32P]ADP-ribose were not affected by the addition of PARG65 (Fig. 2A, bars 5-8), but, notably, the sizes of the polymers were significantly reduced (Fig. 2, B and C, lanes 5-8). In conclusion, even independent of its hydrolyzing activity PARG65 may affect the catalytic activity of PARP-1 by direct protein-protein interaction, resulting in the synthesis of only short polymers.


Figure 1
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  		FIGURE 1.
The catalytic domain of PARG interacts with the automodification domain of PARP-1. A, modular organization of PARP-1 and PARG65 constructs. The localization of zinc fingers (ZF), nuclear localization signal, breast cancer C-terminal region, putative nuclear export signals (put. NES), and mitochondrial targeting element (mito) are indicated. GST or GST-PARG65 were incubated with His-tagged full-length PARP-1 or PARP-1 domains and glutathione-Sepharose. After the washing steps, bound proteins were eluted and subjected to Western blot analysis with anti-polyhistidine or anti-GST antibodies as indicated. 1, input; 2, unbound proteins; 3, wash fractions; 4, eluted proteins. B, after GST pulldown, glutathione-Sepharose bound proteins were incubated with 10 µM [32P]NAD+ and nicked DNA for 20 min. The proteins were separated by SDS-PAGE and radiolabeled PARP-1 was visualized by autoradiography. C, nuclear extracts, prepared from untreated or MNNG-treated (0.5 mM MNNG; 10 min) HeLa cells were subjected to Western blot analysis with anti-PAR and anti-PARP-1 antibodies as indicated. The positions of full-length PARP-1 and proteolysis fragments (*) are indicated. D, immunoprecipitation of HeLa nuclear extracts (NE) with anti-PARP-1 or control antibodies coupled to protein A-Sepharose. Precipitated proteins were subjected to Western blot analysis with anti-PARP-1 and anti-PARG antibodies as indicated. The anti-PARG antibodies recognize the full-length PARG and the 65-kDa form of PARG, which in part is generated by proteolysis during the procedure. B-D, relative molecular weights of marker proteins and the positions of PARP-1 and PARG are indicated. IB, immunoblot.

 
PARG Interacts with XRCC1—PARG65 interacts with the automodification domain of PARP-1 (Fig. 1A), which contains a breast cancer C-terminal domain and is known to be the region important for interactions with other proteins (33) such as for instance XRCC1 (4). To analyze a potential direct connection between PARG and XRCC1, we investigated a direct interaction of these proteins using the following set of experiments. First of all, human full-length PARG was cloned into the yeast two-hybrid reporter plasmid pGADT7 and interaction with an appropriate construct containing XRCC1 (pAS-XRCC1; Ref. 36) was studied in yeast cells (Fig. 3A). Indeed, only in the presence of both PARG and XRCC1 was a supplementation of histidine biosynthesis observed, whereas controls using empty vectors were negative (Fig. 3A). The interaction between PARG65 and XRCC1 was further verified by in vitro GST pulldown experiments. For that purpose, His-tagged XRCC1 was purified from E. coli and incubated with purified GST-PARG65. After GST pulldown, fractions were analyzed by Western blot as described above for PARP-1 (cf. Fig. 1A). Fig. 3B shows that His-tagged XRCC1 bound to GST-PARG65. The interaction between XRCC1 and GST-PARG65 was stable toward salt concentrations of up to 150 mM NaCl (data not shown). To analyze the interaction between PARG and XRCC1 in vivo, we made use of EM9 cells, complemented with either empty vector (EM9-V) or a vector containing His-tagged, human wild-type XRCC1 (EM9-XH) (29). Whole cell extracts of exponentially growing CHO cells were incubated with Ni-NTA-Sepharose. After washing the Ni-NTA beads, elution fractions were subjected to Western blot analysis using specific antibodies against PARG65. As shown in Fig. 3C, PARG clearly bound to His-tagged XRCC1. In conclusion, PARG has the ability to interact with PARP-1 and also with XRCC1, even in the absence of its substrate PAR.


Figure 2
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  		FIGURE 2.
The catalytic activity of PARP-1 is influenced by the interaction with PARG. Purified GST-PARG65 was added to nuclear extracts of HeLa cells in the absence (-) or presence (+) of 150 µM tannins. GST-PARG65 concentrations were without PARG (1 and 5) at 150 (2 and 6), 300 (3 and 7), or 600 nM (4 and 8). 10 µM [32P]NAD and nicked DNA were added and reactions were stopped after 20 min. A, reactions were precipitated with acetone and [32P]ADP-ribose intermediates were separated by thin layer chromatography. The evaluation of [32P]ADP-ribose release is shown. B, in parallel, reactions were subjected to SDS-PAGE (8%) and autoradiography. Relative molecular weights of marker proteins and the position of PARP-1 are indicated. C, the same reactions as in A were precipitated with trichloroacetic acid, polymers were detached by alkaline lysis and subjected to polymer chain length analysis using a 20% polyacrylamide gel. The autoradiogram of the gel is shown and the numbers of ADP-ribose units are indicated.

 
XRCC1 Enhances the Synthesis of Poly(ADP-ribose) in MNNG-treated Cells—XRCC1 interacts with PARP-1 (4) and also with PARG (cf. Fig. 3). Thus, XRCC1 might have the capacity to regulate cellular PAR metabolism. Therefore, in the following experiments we addressed the impact of XRCC1 on PAR synthesis using EM9 cells. First, EM9-V and EM9-XH cells were treated with 0.5 mM MNNG, a supralethal dose to induce poly-(ADP-ribosylation). Interestingly, immunofluorescence analyses clearly showed that higher amounts of PAR polymers were produced in XRCC1-containing EM9-XH cells compared with XRCC1-deficient EM9-V cells (Fig. 4A). Evaluation of quantified PAR signals revealed that in EM9-XH cells, PAR accumulation was increased about 3-4-fold compared with the accumulation observed in EM9-V cells (Fig. 4, B and C). Western blot analyses of cell extracts confirmed this (Fig. 4D). Even 30 min after MNNG treatment, considerably higher amounts of PAR and larger PAR modifications were still detectable in EM9-XH cells, whereas only lower amounts and smaller sizes of PAR modifications remained in EM9-V cells (compare lanes 3 with 6 in Fig. 4D). Hence, after treatment with toxic MNNG doses XRCC1 influences PAR metabolism in living CHO cells.

To further elucidate the regulation of PAR metabolism by XRCC1, PARP and PARG activities were analyzed and compared in different CHO lines deficient or proficient in XRCC1. Western blot analyses revealed no significant differences in PARP-1, or PARG110/PARG65 levels, whereas levels of DNA ligase III (Lig III) were considerably reduced in XRCC1-deficient cells (data not shown). Consequently, whereas Lig III is stabilized by XRCC1 (35), cellular XRCC1 does not appear to be essential for the stability of PARP-1 or PARG. Previously, Masson et al. (4) proposed that XRCC1 down-regulates PARP-1 activity. Accordingly, we observed an inhibition of poly(ADP-ribosylation) in a reconstituted system with recombinant proteins, when XRCC1 was incubated with PARP-1 in a molar ratio of 8:1 (Fig. 5A). Furthermore, we found that PARG activity was unaltered, even if an excess of XRCC1/Lig III was added to GST-PARG65 (Fig. 5B). Analyzing relative PARP and PARG activities in nuclear extracts of different CHO lines deficient or proficient in XRCC1 resulted in comparable PARP and PARG activities in all cell lines (Fig. 5, C and D). Similarly, it has been reported that cellular NAD content and relative PARP activity appeared normal in EM9 cells (40). These seemingly contradictory observations are explainable because in living cells the level of XRCC1 is not higher than the level of PARP-1. A direct decreasing effect on PARP-1 activity was only observed if an excess of XRCC1 was added to isolated PARP-1 (Fig. 5A).

In response to treatment with genotoxic agents, XRCC1 is recruited to DNA single-strand breaks in a PARP-1-dependent fashion (7). In comparison to PARP-1, the cellular concentration of XRCC1 is lower (41). Therefore, recruitment might represent an increase of the local concentration of a limited factor at sites of recruitment. Because neither PARP-1 nor PARG activities are directly regulated by XRCC1 (Fig. 5, C and D), XRCC1 recruitment might be required for the observed PAR accumulation in MNNG-treated XRCC1-proficient cells (Fig. 4). Therefore, in the next experiment, we reconstituted XRCC1 recruitment in vitro by supplementation of XRCC1-deficient nuclear extracts with an excess of recombinant XRCC1. For that purpose, nuclear extracts from XRCC1-deficient EM9 cells were complemented with recombinant XRCC1 during monitoring of PARP activity for a period of 20 min using nicked DNA and [{alpha}-32P]NAD+. [32P]Poly(ADP-ribosylation) was visualized by SDS-PAGE (Fig. 6A) and polymer chain lengths were analyzed in parallel (Fig. 6B). Indeed, we observed that PAR synthesis was dramatically enhanced after supplementation with XRCC1 (Fig. 6). Both, the amounts and the sizes of formed polymers increased significantly after supplementing XRCC1 to EM9 extracts (Fig. 6, A and B, right lanes). Large and branched PAR polymers, which were unable to enter the gel, were only produced when XRCC1 was present (Fig. 6, A and B, top of right lanes). The fractions of [32P]poly(ADP-ribosylated) proteins with large modifications were quantified as insoluble fractions after acetone precipitation (PAR* in Fig. 6C). Soluble [32P]ADP-ribose metabolites PAR, ADP-ribose, AMP, and NAD+ were separated by thin layer chromatography and relative ratios of all [32P]ADP-ribose metabolites were determined (Fig. 6C). The evaluation of the overall [32P]ADP-ribose metabolism revealed that in the presence of XRCC1 a significant and reproducible accumulation of large [32P]PAR modifications (PAR*) was achieved (Fig. 6C). Moreover, the levels of [32P]ADP-ribose decreased when XRCC1 was added (Fig. 6C). Thus, the observed enhanced formation of [32P]PAR polymers was partly caused by a suppression of PARG activity. Nevertheless, XRCC1 stimulated PARP activity because the consumption of [32P]NAD+ was significantly increased in reactions supplemented with XRCC1 (Fig. 6C). Similarly, supplementation of nuclear extracts from HeLa cells with recombinant XRCC1 resulted in increased poly(ADP-ribosylation) (data not shown). A recruitment of or supplementation with XRCC1 appears to be necessary for efficient PAR accumulation. One possible explanation for this is that XRCC1 displaces PARG from binding PAR or from associations with other proteins. However, because PARG and PARP-1 interact (cf. Fig. 1) and PARG thereby down-regulates PAR syntheses (cf. Fig. 2), it is conceivable that the observed accumulation of PAR polymers is a result of alternating interactions between PARG, PARP-1, and XRCC1.


Figure 3
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  		FIGURE 3.
PARG interacts with XRCC1. A, pGADT7-PARG110, pAS-XRCC1, and control plasmids as indicated were transformed into yeast cells and diploids selected using medium lacking leucine and tryptophan (+His). For validation of protein-protein interaction, colonies were transferred to histidine-free plates (-His). B, His-tagged XRCC1 was incubated with GST or GST-PARG65 and glutathione-Sepharose. After the washing steps, bound proteins were eluted and subjected to Western blot analysis with anti-polyhistidine or anti-GST antibodies as indicated. 1, input; 2, unbound proteins; 3, wash fractions; 4, eluted proteins. C, whole cell extracts of EM9-V or EM9-XH cells were incubated with Ni-NTA-Sepharose. Proteins bound to the Sepharose beads were eluted with 80 mM imidazole and subjected to Western blot analysis with anti-XRCC1 or anti-PARG antibodies as indicated. 1, input; 2, unbound proteins; 3, eluted proteins.

 


Figure 4
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  		FIGURE 4.
MNNG treatment triggers increased PAR synthesis in EM9-XH cells. A, nonconfluent growing EM9-V and EM9-XH cells were treated with 0.5 mM MNNG in the absence or presence of 1 mM 3-ABA as indicated. 10 min thereafter, cells were fixed and immunostained with anti-PAR antibodies (left panels) and counterstained with DAPI (right panels). Representative images are shown. B, diagrams of the fluorescence signals from the single cells marked with an asterisk in A. C, quantification of PAR accumulation in MNNG-treated cells in the absence or presence of 3-ABA as indicated. D, EM9-V or EM9-XH cells were not treated or treated with 0.5 mM MNNG and whole cell extracts were prepared and subjected to Western blot analysis with anti-PAR or anti-{alpha}-tubulin antibodies, respectively. For lanes 1 and 4, extracts were prepared from untreated cells, whereas the cells represented by lanes 2 and 5 were MNNG treated for 12 min. For lanes 3 and 6, cells were treated with MNNG for 10 min and thereafter incubated in drug-free medium for a further 20 min. Relative molecular sizes of marker proteins are indicated.

 


Figure 5
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  		FIGURE 5.
Influence of XRCC1 and Lig III on PARP-1 and PARG activity. A, 50 nM recombinant PARP1 was combined with 400 nM recombinant XRCC1 and 100 nM Lig III as indicated. 10 µM [{alpha}-32P]NAD and 10 µg/ml nicked DNA were added and stopped 20 min thereafter. Relative [32P]ADP-ribose incorporation was estimated by trichloroacetic acid precipitation. B, 50 nM recombinant GST-PARG65 was mixed with 400 nM recombinant XRCC1 and 100 nM Lig III as indicated. 3 µM [32P]PAR was added and reactions were precipitated with acetone after 20 min. [32P]PAR metabolites were dissolved in TE and separated by thin layer chromatography. Positions of [32P]PAR and [32P]ADP-ribose are indicated. C and D, nuclear extracts from CHO cell lines AA8, EM9, EM9-V, and EM-XH were prepared. Relative activities are given in percentages of the values obtained with extracts of AA8 cells. C, relative [32P]ADP-ribose incorporation by trichloroacetic acid precipitation was estimated for all four CHO cell lines. D, relative PARG activity was determined for all four cell lines.

 
XRCC1-induced Accumulation of PAR Leads to Apoptotic Cell Death—It is known that an accumulation of PAR in response to high doses of MNNG triggers several further apoptotic events, including translocation of AIF from mitochondria to the nucleus and nuclear shrinkage within a few hours (25). Because we observed increased PAR accumulation in EM9-XH cells compared with EM9-V cells after treatment with supralethal MNNG doses (cf. Fig. 4), in the next experiments we monitored translocation of AIF and nuclear shrinkage in CHO cells, 3 or 6 h after MNNG treatment (Fig. 7). The cellular content of AIF was comparable in EM9-V and EM9-XH cells (Fig. 7C). Translocation of AIF after MNNG treatment could be detected in only a few EM9-V cells and in most EM9-XH cells, but not in the presence of the PARP inhibitor 3-ABA (Fig. 7, A and B). Because XRCC1 increases the amount of PAR formation (Figs. 4 and 6), it appears that XRCC1 might as well regulate PARP-1-dependent translocation of AIF from mitochondria to the nucleus. Notably, 6 h after MNNG treatment, shrunken nuclei were detected in more than 80% of EM9-XH cells but only in 10% of EM9-V cells (Fig. 7D). When the XRCC1 proficient parental cell line AA8 was treated with MNNG, similar apoptotic features as obtained with the EM9-XH cells were observed (data not shown). The caspase inhibitor Z-VA-D(OMe)-fmk (100 µM) failed to block MNNG-induced nuclear shrinkage in EM9-XH cells (Fig. 7D). In a control experiment, the impact of XRCC1 on induction of apoptosis was analyzed, using staurosporine, another trigger of cell death (Fig. 8). Staurosporine has two effects on nuclear structure, either causing caspase-independent partial nuclear condensation (stage I) or caspase-dependent advanced nuclear condensation and fragmentation (stage II) (42). We observed that AIF translocation and nuclear condensation induced by staurosporine were comparable in EM9-V and EM9-XH cells (Fig. 8). Furthermore, Z-VAD(OMe)-fmk did not prevent AIF translocation but effectively blocked the formation of shrunken nuclei (stage II) in both cell lines (Fig. 8). Thus, the impact of XRCC1 on apoptotic events appears to be restricted to MNNG-induced caspase-independent cell death.


Figure 6
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  		FIGURE 6.
Supplementation of EM9 nuclear extracts with recombinant XRCC1 increases the PAR level. Nuclear extracts (15 µg of protein/reaction) from EM9 cells were supplemented with 1 µg of bovine serum albumin (BSA) or recombinant XRCC1 as indicated. 10 µM [32P]NAD and 10 µg/ml nicked DNA were added and reactions were stopped after 20 min. A, proteins were subjected to SDS-PAGE and autoradiography. Relative molecular weights of marker proteins and the positions of PARP-1 and XRCC1 are indicated. B, reactions were precipitated with trichloroacetic acid, polymers were detached and subjected to polymer chain length analysis using an 8% polyacrylamide gel. The autoradiogram of the gel is shown and the numbers of ADP-ribose units are indicated. C, reactions were precipitated with acetone. Amounts of insoluble precipitates, which contained proteins modified with large PAR polymers, were termed PAR* fractions and quantified. Soluble 32P metabolites were separated by thin layer chromatography and signals of [32P]PAR, -ADP-ribose, -AMP, and -NAD were quantified. The evaluation of relative ratios of all 32P metabolites is shown. In the control reaction (diagram bottom right) no proteins were present.

 


Figure 7
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  		FIGURE 7.
MNNG treatment triggers translocation of AIF and nuclear shrinkage in EM9-XH cells. A, nonconfluent growing EM9-V and EM9-XH cells were treated with 0.5 mM MNNG in the absence or presence of 1 mM 3-ABA as indicated. 10 min thereafter cells were washed with fresh medium and incubated for a further 3 h at 37 °C. Then cells were fixed and immunostained with anti-AIF antibodies (left panels) and counterstained with DAPI (middle panels). Representative images are shown. B, quantification of cells with AIF translocation, 3 h after MNNG treatment in the absence or presence of 1 mM 3-ABA as indicated. C, Western blot analysis of whole cell extracts of EM9-V and EM9-XH cells with antibodies against XRCC1, AIF, and YY1. D, quantification of cells with nuclear shrinkage, 6 h after MNNG treatment in the absence or presence of 100 µM Z-VAD(OMe)-fmk as indicated. Representative images of control and shrunken nuclei are given above.

 
Poly(ADP-ribosylation) is an energy-consuming process. Consequently, cellular levels of ATP and NAD were considerably reduced within 60 min after MNNG treatment in all tested cell lines (Fig. 9, A and B). 13 h after MNNG treatment, EM9-V cells were permeable for staining with propidium iodide, whereas most of EM9-XH cells had already died (Fig. 9D). Finally, 24 h after MNNG treatment nearly all EM9-V cells had died, whereas about 20% of EM9-XH survived (Fig. 9C). Thus, in EM9-V cells cell death was presumably caused by energy depletion.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The investigation presented here suggests a new level of coordination of MNNG-induced cell death regulated by interactions between PARP-1, PARG, and XRCC1. Several conclusions can be drawn from our study. First of all, PARG interacts with PARP-1 and XRCC1. Second, PARG can regulate PARP-1 activity. Third, XRCC1 regulates PAR-mediated apoptotic cell death induced by supralethal doses of MNNG.

The creation of different PARG knockout models revealed the importance of PAR polymers for cellular survival. For example, genetic PARG inactivation in D. melanogaster resulted in a severe PAR accumulation in neuronal cells and lethality at the larval stage (20) and mouse cells lacking PARG showed an accumulation of PAR leading to cell death by apoptosis (21). Thus, PAR can induce apoptosis and PAR signaling appears to play an important role in embryonic development. In a recent proteomic approach, localization of a fraction of PARG to messenger ribonucleoparticles and an interaction of PARG with Fragile-X-related protein was discovered (27). Given the fact that PARG activity is involved in many different cellular events these observed interactions could be important also for the regulation of the activities of PARP enzymes other than PARP-1. All current models regarding the functions of poly-(ADP-ribosylation) are based on the interplay between PARP-1 and PARG, but a direct interaction between both enzymes has not been described before. In the study presented here we show for the first time that PARP-1 interacts with PARG, even independently of the substrates NAD or PAR. In addition, we characterize XRCC1 as another interaction partner of PARG. Remarkably, phosphorylation of XRCC1 by protein kinase CK2 influences its interaction with the DNA repair protein polynucleotide kinase, as recently shown (30). Furthermore, in a large scale characterization of nuclear phosphoproteins from HeLa cells, phosphorylated PARG was detected (43). Thus, it is tempting to speculate that PARG activity or its interaction with other proteins might be regulated by phosphorylation. Nevertheless, functional analyses of how PARG activity is modulated in living cells are lacking yet.


Figure 8
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  		FIGURE 8.
EM9 cells are susceptible to AIF-mediated apoptosis. A, nonconfluent growing EM9-V and EM9-XH cells were treated with 800 nM staurosporine in the absence or presence of 100 µM Z-VAD(OMe)-fmk as indicated. 6 h thereafter cells were fixed and immunostained with anti-AIF antibodies (left panels) and counterstained with DAPI (right panels). Representative images are shown and stage I or stage II shrunken nuclei are indicated. B, the evaluations of counted cells with AIF translocation and nuclear shrinkage stage I or stage II are given below.

 


Figure 9
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  		FIGURE 9.
In EM9-V cells MNNG-induced cell death is caused by energy depletion. Nonconfluent growing EM9-V and EM9-XH cells were treated with 0.5 mM MNNG for 10 min, thereafter cells were washed with fresh medium and incubated at 37 °C for further time periods as indicated. A, the cellular levels of ATP compared with untreated cells are shown. B, the cellular levels of NAD compared with untreated cells are shown. C, 24 h after MNNG treatment cells were trypsinized, mixed with trypan blue, and cell survival was estimated by counting vital cells. The surviving fraction was calculated by dividing the average number of surviving cells on treated plates by the average number on untreated plates. A-C, all data are the mean (±S.D.) of at least three independent sets of experiments. D, 13 h after MNNG treatment, unfixed cells were stained with propidium iodide (PI) (left panels) and DAPI (right panels). Representative images are shown.

 
The biological significance of PARP-1 activity has been the subject of numerous publications but it is poorly understood how this catalytic activity is modulated (1-3). As it is known, PARP-1 activity is induced by DNA breaks, introduced either directly or indirectly, and depends on the availability of NAD. Among the proteins interacting with PARP-1, XRCC1 and DNA-dependent protein kinase have been reported to negatively regulate PARP-1 activity (4, 44). On the other hand, we characterized transcription factors that have the ability to directly stimulate PARP-1 activity (45-47). Furthermore, an allosteric activation of PARP-1 automodification by Mg2+, Ca2+, histones H1 and H2, and polyamines has been demonstrated (48). Here, with PARG we present another PARP-1-interacting protein that may function as a negative regulator. In line with these findings, Cortes et al. (19) reported that when the full-length isoform of PARG was knocked-out in mice, residual PARG60 severely reduced the automodification activity of PARP-1 in vivo. Thus, it is feasible that PARG, when interacting with PARP-1, inhibits the automodification reaction and thereby guarantees that large PAR modifications are not generated. This inhibition of PARP-1 activity might be achieved directly or indirectly by displacement of PARP-1 activators as histones or transcription factors. Thus, PARG appears to have a dual function in PAR metabolism, degradation of PAR and down-regulation of PAR synthesis. In contrast to PARG, PARP-1 is highly abundant in the nucleus (14). In situations of low levels of DNA damage, PAR synthesis and degradation are balanced and no accumulation of PAR occurs. It has therefore been assumed that the high specific activity of PARG compensates for its low cellular concentration (39). Based on our findings it can be speculated that at low levels of DNA damage, only a minor fraction of nuclear PARP-1 is catalytically activated and at the same time down-regulated by PARG. Hence, the mechanism of regulation proposed here ensures that in the absence of high amounts of DNA lesions, PAR polymers do not accumulate. Accordingly, it has been reported that in response to moderate doses of alkylating agents, such as 15 µM MNNG, the production of PAR is limited, whereas higher MNNG doses trigger the formation of large PAR polymers in cells peaking within 10-15 min (49).


Figure 10
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  		FIGURE 10.
XRCC1 regulates MNNG-induced cell death by coordinating the activities of PARP-1 and PARG. Toxic doses of MNNG introduce irreparable DNA lesions. PARP-1 interacts with PARG and both enzymes are catalytically activated, and PAR synthesis and degradation are induced. In EM9-XH cells (right) XRCC1 is recruited by automodified PARP-1, resulting in a displacement of PARG. As a consequence, the extent and length of PAR polymers increase. Accumulated PAR polymers then trigger AIF release, nuclear shrinkage, and apoptotic cell death. In XRCC1-deficient EM9-V cells (left) activation of PARP-1 and PARG cause depletion of NAD and ATP leading to cell death.

 
XRCC1, for which no enzymatic activity has been described, plays a central role in DNA single-strand break repair (6). Here we suggest an unexpected role for XRCC1, influencing apoptotic cell death in response to treatment with supralethal doses of MNNG. XRCC1 binds preferentially to automodified PARP-1, most likely via a putative PAR recognition and binding motif within its internal breast cancer C-terminal I motif (4, 50). Furthermore, it is established that XRCC1 is immediately recruited to DNA lesions by automodified PARP-1 (7, 8). At physiological ratios a direct modulation of PARP-1 or PARG activity by XRCC1 was not detected (Fig. 5, C and D). Notably, the effect of XRCC1 on PAR accumulation was only observed in response to treatment of cells with toxic doses of MNNG (Fig. 4). Because XRCC1 preferentially interacts with automodified PARP-1 (4, 51, 52) it may be suspected that PARG could be displaced from its interaction with the PARP-1 automodification domain. Presumably, the repression of PARP-1 activity by PARG is abrogated by XRCC1, resulting in increased PAR synthesis. Based on the experiments presented here we propose the following hypothesis (Fig. 10): PARP-1 interacts with PARG, even in the absence of significant automodification. In response to treatment with supralethal MNNG doses massive DNA lesions are introduced, PAR synthesis and degradation proceed, and automodified PARP-1 instantly recruits XRCC1. It can be speculated that if amounts of DNA lesions are beyond the cellular repair capacity, PARP-1 still remains catalytically active, whereas PARG might be displaced by XRCC1. As a result, the number and length of PAR polymers would increase further. Accumulated PAR polymers then trigger the translocation of AIF from mitochondria and nuclear shrinkage, ultimately leading to caspase-independent apoptotic cell death (25). In XRCC1-deficient cells, the combined activities of PARP-1 and PARG cause NAD and ATP depletion resulting in non-apoptotic cell death (Fig. 9). In addition, it has been reported that after methyl methanesulfonate treatment NAD depletion was augmented in EM9-V cells compared with EM9-XH cells (53). Cell killing effects of alkylating agents depend on the doses of agents and the growth state of CHO cells (54). Here we demonstrate that in response to treatment with a cytotoxic dose of MNNG, rapid accumulation of PAR and induction of apoptotic cell death occurred only in XRCC1 proficient cells. However, treatment with H2O2 introduces oxidative DNA damage including direct single-strand breaks, so that in difference to treatment with MNNG, substantial amounts of PAR are still detectable 30 min after the treatment (55). Equal amounts of PAR foci triggered by 20 mM H2O2 were detected in EM9-V and EM9-XH cells (55). Consequently, these PAR foci seem to arise independent of the presence of XRCC1. Presumably, H2O2-induced PAR foci persist because of the limiting amount of PARG molecules compared with the amount of damage-activated PARP-1 molecules. In line with this, we observed nuclear condensation also of most of EM9-V cells in response to a treatment with 1 mM H2O2 (data not shown). Thus, the impact of XRCC1 on PAR accumulation and cell death depends on the type and amount of DNA lesions introduced and obviously seems to be specific for treatment with toxic MNNG doses.

When a cell is damaged, the cell has two options: repair or die. If the damages are too extensive the cell must also decide which cell death pathway to follow. Several ATP-dependent steps are required for apoptotic signal transduction and an excessive NAD/ATP depletion below 50% is believed to induce cell death by necrosis (56). Apparently, PARP-1 plays a dual role in triggering cell death. On the one hand PARP-1 consumes NAD, thus PARP-1 activation may cause energy deprivation. Accordingly, PARP-1 overactivation leads to necrotic cell death (24). Moreover, PARP-1-mediated necrotic cell death after treatment with 0.5 mM MNNG was detected in mice fibroblasts by HMGB1 exclusion (57), whereas AIF localization was not monitored in that study. On the other hand, accumulating PAR polymers appear to be an effective apoptotic stimulus signaling AIF release (25, 22). Additionally, studies with cortical neurons revealed that an accumulation of PAR and not excessive NAD consumption was responsible for initiation of apoptosis (22, 58).

Efficient progression of both DNA repair and apoptosis are essential for genome integrity. Poly(ADP-ribosylation) plays relevant roles in DNA damage sensing/repair and apoptosis. From our study we suggest that XRCC1 might be another important determinant regulating both processes.


   FOOTNOTES
 
* This work was supported by Deutsche Forschungsgemeinschaft Grant ZI 541/3. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed. Tel.: 49-3083852194; Fax: 49-3083857194; E-mail: lity{at}chemie.fu-berlin.de.

2 The abbreviations used are: PAR, poly(ADP-ribose); AIF, apoptosis inducing factor; GST, glutathione S-transferase; CHO, Chinese hamster ovary; Lig III, DNA ligase III; MNNG, methyl-N'-nitro-N'-nitrosoguanidine; PARG, poly-(ADP-ribose) glycohydrolase; PARP, poly(ADP-ribose) polymerase; XRCC1, x-ray repair cross-complementing 1; 3-ABA, 3-aminobenzamide; DAPI, 4',6-diamidino-2-phenylindole; Ni-NTA, nickel-nitrilotriacetic acid; Z, benzyloxycarbonyl; fmk, fluoromethyl ketone. Back


   ACKNOWLEDGMENTS
 
We thank Dr. K. W. Caldecott for gifts of CHO cells and plasmids encoding human XRCC1 and Dr. E. Petermann for critical reading the manuscript. We further thank G. Buchlow for DNA sequencing.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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