Nucleosomal Core Histones Mediate Dynamic Regulation of Poly(ADP-ribose) Polymerase 1 Protein Binding to Chromatin and Induction of Its Enzymatic Activity
2007; Elsevier BV; Volume: 282; Issue: 44 Linguagem: Inglês
10.1074/jbc.m705989200
ISSN1083-351X
AutoresAaron D. Pinnola, Natasha Naumova, Meera Shah, Alexei V. Tulin,
Tópico(s)Genomics and Chromatin Dynamics
ResumoPoly(ADP-ribose) polymerase 1 protein (PARP1) mediates chromatin loosening and activates the transcription of inducible genes, but the mechanism of PARP1 regulation in chromatin is poorly understood. We have found that PARP1 interaction with chromatin is dynamic and that PARP1 is exchanged continuously between chromatin and nucleoplasm, as well as between chromatin domains. Specifically, the PARP1 protein preferentially interacts with nucleosomal particles, and although the nucleosomal linker DNA is not necessary for this interaction, we have shown that the core histones, H3 and H4, are critical for PARP1 binding. We have also demonstrated that the histones H3 and H4 interact preferentially with the C-terminal portion of PARP1 protein and that the N-terminal domain of PARP1 negatively regulates these interactions. Finally, we have found that interaction with the N-terminal tail of the H4 histone triggers PARP1 enzymatic activity. Therefore, our data collectively suggests a model in which both the regulation of PARP1 protein binding to chromatin and the enzymatic activation of PARP1 protein depend on the dynamics of nucleosomal core histone mediation. Poly(ADP-ribose) polymerase 1 protein (PARP1) mediates chromatin loosening and activates the transcription of inducible genes, but the mechanism of PARP1 regulation in chromatin is poorly understood. We have found that PARP1 interaction with chromatin is dynamic and that PARP1 is exchanged continuously between chromatin and nucleoplasm, as well as between chromatin domains. Specifically, the PARP1 protein preferentially interacts with nucleosomal particles, and although the nucleosomal linker DNA is not necessary for this interaction, we have shown that the core histones, H3 and H4, are critical for PARP1 binding. We have also demonstrated that the histones H3 and H4 interact preferentially with the C-terminal portion of PARP1 protein and that the N-terminal domain of PARP1 negatively regulates these interactions. Finally, we have found that interaction with the N-terminal tail of the H4 histone triggers PARP1 enzymatic activity. Therefore, our data collectively suggests a model in which both the regulation of PARP1 protein binding to chromatin and the enzymatic activation of PARP1 protein depend on the dynamics of nucleosomal core histone mediation. Eukaryotic chromatin organization involves the fundamental nucleosomal unit, which consists of four core histones plus a linker histone (1Wolffe A.P. Essays Biochem. 2001; 37: 45-57Crossref PubMed Scopus (59) Google Scholar). Recently, it has been shown that the activity of transcription complexes at nucleosomes is regulated by the PARP1 3The abbreviations used are: PARP1, poly(ADP-ribose) polymerase 1; PARPe, poly(ADP-ribose) polymerase embryonic; PARG, poly(ADP-ribose) glycohydrolase; pADPr, poly(ADP-ribose); FRAP, fluorescence recovery after photobleaching; DTT, dithiothreitol; MNase, micrococcal nuclease; GFP, green fluorescent protein; MS, mass spectrometry; DsRed, red fluorescent protein; EGFP, enhanced green fluorescent protein; EYFP, enhanced yellow fluorescent protein; ECFP, enhanced cyan fluorescent protein; UAS, upstream activating sequence; PCR, polymerase chain reaction. protein (2Tulin A. Kim A. Science. 2003; 299: 560-562Crossref PubMed Scopus (388) Google Scholar, 3Kim M.Y. Kim S. Gevry N. Lis J. T. Kraus W. L. Cell. 2004; 119: 803-814Abstract Full Text Full Text PDF PubMed Scopus (456) Google Scholar). Notwithstanding these findings, major gaps in our present understanding exist that involve the mechanism by which the PARP1 protein binds to specific chromatin domains and the mechanism by which the local PARP1 protein is activated in response to developmental and environmental stimuli. After histones, PARP1 is the most abundant nuclear protein (4Virag L. Kim C. Pharmacol. Rev. 2002; 54: 375-429Crossref PubMed Scopus (1235) Google Scholar). The distribution of PARP1 in chromatin is broad and occurs in regions characterized by distinct cell types (2Tulin A. Kim A. Science. 2003; 299: 560-562Crossref PubMed Scopus (388) Google Scholar, 3Kim M.Y. Kim S. Gevry N. Lis J. T. Kraus W. L. Cell. 2004; 119: 803-814Abstract Full Text Full Text PDF PubMed Scopus (456) Google Scholar, 5Dantzer F. Kim H.P. Vonesch J.L. de Murcia G. Menissier-de Murcia J. Nucleic Acids Res. 1998; 26: 1891-1898Crossref PubMed Scopus (138) Google Scholar). Nevertheless, exactly how the PARP1 enzyme interacts with chromatin in vivo has not been thoroughly investigated, and the molecular basis for PARP1 binding to chromatin remains poorly understood. Although zinc fingers within the PARP1 protein contribute to DNA binding in vitro, they specifically recognize damaged DNA (6Gradwohl G. Kim J.M. Molinete M. Simonin F. Koken M. Hoeijmakers J.H. de Murcia G. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 2990-2994Crossref PubMed Scopus (228) Google Scholar) and therefore do not contribute to the association of PARP1 with intact chromatin. Moreover, a PARP1 paralog, PARP2, that has no zinc fingers and no direct DNA binding capability, nevertheless exhibits a pattern of chromatin association similar to PARP1 and is able to partially complement PARP1 functions in a PARP1 null mutant (7Amé J.C. Kim V. Scureiber V. Niedergang C. Apiou F. Decker P. Muller S. Hoger T. Ménissier-de Murica J. de Murica G. J. Biol. Chem. 1999; 274: 17860-17868Abstract Full Text Full Text PDF PubMed Scopus (616) Google Scholar, 8Babiychuk E. Kim P.B. Storozhenko S. Fuangthong M. Chen Y. O'Farrell M.K. Van Montagu M. Inze D. Kushnir S. Plant J. 1988; 15: 635-645Crossref Google Scholar, 9Meder V.S. Kim M. de Murcia G. Schreiber V. J. Cell Sci. 2005; 118: 211-222Crossref PubMed Scopus (147) Google Scholar). This suggests that PARP1 and PARP2 both bind chromatin indirectly, through an interaction with one or more DNA-binding proteins. A key aim of this study is to determine the specific mechanisms by which PARP1 protein associates with chromatin in vivo. Considerable evidence now suggests that PARP1 interacts with chromatin by binding to histones (10D'Amours D. Kim S. D'Silva I. Poirier G.G. Biochem. J. 1999; 342: 249-268Crossref PubMed Scopus (0) Google Scholar). For example, histones H1, H2A, and H2B are efficient targets for PARP1 binding in vitro (11Buki K.G. Kim P.I. Hakam A. Kun E. J. Biol. Chem. 1995; 270: 3370-3377Abstract Full Text Full Text PDF PubMed Scopus (60) Google Scholar) and are enzymatically modified by PARP1 (12Aubin R.J. Kim A. de Murcia G. Mandel P. Lord A. Grondin G. Poirier G.G. EMBO J. 1983; 2: 1685-1693Crossref PubMed Scopus (57) Google Scholar, 13Krupitza G. Kim P. Biochemistry. 1988; 28: 4054-5060Crossref Scopus (44) Google Scholar, 14Poirier G.G. Kim C. Champagne M. Mazen A. Mandel P. Eur. J. Biochem. 1982; 127: 437-442Crossref PubMed Scopus (38) Google Scholar). This idea is, however, complicated by the fact that Drosophila histone H1 was recently reported as an antagonist of PARP1 binding to chromatin (3Kim M.Y. Kim S. Gevry N. Lis J. T. Kraus W. L. Cell. 2004; 119: 803-814Abstract Full Text Full Text PDF PubMed Scopus (456) Google Scholar). In addition, accumulation of PARP1 interactors, which have to date been identified through in vitro experiments, has resulted in findings suggesting that almost none significantly co-localizes with PARP1 in chromatin. To clarify the many issues involved with PARP1 protein binding activity to chromatin and the activation of its enzymatic activity, we sought an appropriate experimental model. As both an organismal and genetic model system, Drosophila was selected as the best tool for studying the function and dynamics of PARP1 protein interaction with chromatin in vivo. Unlike mammals, which have multiple PARP-related proteins, only a single nuclear PARP1 gene (15Adams M.D. Kim S.E. Holt R.A. Evans C.A. Gocayne J.D. Amanatides P.G. Scherer S.E. Li P.W. Hoskins R.A. Galle R.F. George R.A. Lewis S.E. Richards S. Ashburner M. Henderson S.N. et al.Science. 2000; 287: 2185-2195Crossref PubMed Scopus (4854) Google Scholar, 16Hanai M. Kim M. Kobayashi S. Miwa M. Uchida K. J. Biol. Chem. 1998; 273: 11881-11886Abstract Full Text Full Text PDF PubMed Scopus (24) Google Scholar, 17Tulin A. Kim D. Spradling A.C. Genes Dev. 2002; 16: 2108-2119Crossref PubMed Scopus (178) Google Scholar) is present in the Drosophila genome. Therefore, in this paper we study Drosophila PARP1 to first identify the components of nucleosomal core particles, which are responsible for PARP1 protein binding and activation, and then, based on these findings, deconstruct and analyze the machinery responsible for PARP1-chromatin interaction. Drosophila Strains and Genetics—Genetic markers are described in FlyBase 1999, and stocks were obtained from the Bloomington Stock Center, except as indicated. pP{w1, UAST::PARP1-DsRed}, called UAS::Parp1-DsRed, was described in Ref. 17Tulin A. Kim D. Spradling A.C. Genes Dev. 2002; 16: 2108-2119Crossref PubMed Scopus (178) Google Scholar. The transgenic stock with pP{w1, UAST:: PARG-EGFP}, called UAS::Parg-EGFP, was described in Ref. 18Tulin A. Kim N.M. Menon A.K. Spradling A.C. Genetics. 2006; 172: 363-371Crossref PubMed Scopus (48) Google Scholar. The following GAL4 driver strains were used: 69B-GAL4 (19Manseau L. Kim A. Brower D. Budhu A. Elefant F. Phan H. Philp A.V. Yang M. Glover D. Kaiser K. Palter K. Selleck S. Dev. Dyn. 1997; 209: 310-322Crossref PubMed Scopus (198) Google Scholar) and arm::GAL4 (Bloomington stock no. 1560). Balancer chromosome carrying Kr::GFP, i.e. TM3, P{w1, Kr-GFP} (20Casso D. Kim F. Kornberg T.B. Mech. Dev. 2000; 91: 451-454Crossref PubMed Scopus (109) Google Scholar), was used to identify heterozygous and homozygous ParpCH1 (17Tulin A. Kim D. Spradling A.C. Genes Dev. 2002; 16: 2108-2119Crossref PubMed Scopus (178) Google Scholar). Construction of Transgenic Drosophila—To construct UAS::H2A-ECFP and UAS::H1-EYFP, we generated full-length histone H2A and histone H1 open reading frame using PCR. Primers used were as follows: for H1 cloning, h1d, CACCatgtctgattctgcagttg, and h1r, ctttttggcagccgtag; and for H2A cloning, h2ad, CACCatgtctggacgtggaaaagg, and h2ar, ggccttcttctcggtcttcttg. We used wild-type Drosophila genomic DNA as a template for PCR. The resulting PCR products were cloned through The Drosophila Gateway™ vector cloning system (Carnegie Institution of Washington) into the corresponding vector for Drosophila transformation. Transformation was performed as described in Ref. 21Spradling A.C. Kim G.M. Science. 1982; 218: 341-347Crossref PubMed Scopus (1174) Google Scholar, with modifications (22Prokhorova A.V. Kim M.A. Shostak N.G. Barskii V.E. Golubovskii M.D. Genetika (Moscow). 1994; 30: 874-878PubMed Google Scholar). Fluorescence Recovery After Photobleaching (FRAP) Assay—FRAP experiments on live Drosophila tissues were performed as described in Ref. 23Phair R.D. Kim T. Nature. 2000; 404: 604-609Crossref PubMed Scopus (972) Google Scholar. To conduct these experiments, we used a Leica TCS SP2 confocal microscope with capacity for FRAP. To avoid the oxidative stress and other damage that lasers can cause, we used only the minimal level of laser power. This step extended the “bleaching” phase but did not affect our results. To collect FRAP data, we employed the “FlyMode” program, which allows data collection even during the bleaching phase. The recordings were performed via a 63× 1.4 NA oil immersion objective. We found that all the fluorescent epitopes we tested (ECFP, EYFP (Venus), EGFP, and DsRed) were appropriate for FRAP assays, as well as for regular confocal analysis. We did not detect epitope-specific biases in the function, expression dynamics, or localization of any fused moiety. We used transgenic fly stocks that express appropriate fluorescent epitope-tagged protein. Tissues were dissected in Grace’s medium, and dynamic movement of fluorescent proteins was analyzed for 20-30 min following dissection. Nuclei Isolation and Micrococcal Nuclease Digestion—0.5 g of fresh pupae were homogenized in 10 ml of buffer A1 (15 mm Tris-HCl, pH 7.5, 60 mm KCl, 15 mm NaCl, 5 mm MgClB2B, 0.5% Triton X-100, 0.1 mm EGTA, 0.5 mm DTT, and CompletePTMP protease inhibitors (Roche Applied Science)), using a Potter homogenizer (Pyrex). The homogenate was filtered through two layers of Miracloth (Calbiochem), homogenized using a Dounce homogenizer (Pestle B) (Kontes Glass Co.) with 10-15 strokes, and centrifuged for 4 min at 4000 × g at 4 °C. The pellet was washed once with 10 ml of the A1 buffer, then resuspended in 6 ml of A1, loaded onto 3 ml of buffer A1/0.3 m sucrose, and centrifuged for 6 min at 1500 × g at 4 °C. The nuclei were washed once with 3 ml of micrococcal nuclease (MNase) digestion buffer (15 mm Tris-HCl, pH 7.5, 60 mm KCl, 15 mm NaCl, 1 mm CaClB2B, 0.3 m sucrose, 0.5 mm DTT, and EDTA-free CompletePTMP protease inhibitors (Roche Applied Science)), diluted by MNase digestion buffer to 1 ml, and incubated with ∼200 units of MNase (Worthington) at 37 °C for 3 min, 650 rpm in Thermomixer (Eppendorf). An amount of MNase sufficient for complete chromatin digestion to mononucleosomes was chosen in preliminary experiments for each aliquot of the enzyme. The reaction was stopped by 25 μl of 0.5 m EDTA. After the addition of 200 μl of M buffer (190 mm Tris-HCl, pH 7.5, 25% glycerol, 440 mm NaCl, 5 mm MgClB2B, 125 mm NaF, 5 mm NaB3BVOB4B, 5 mm EDTA, 1% Nonidet P-40, 5 mm DTT, and 2× CompletePTMP protease inhibitors (Roche Applied Science)), the nuclei were lysed on a rotating platform at 4 °C for 20 min. The nuclei extract was clarified by centrifugation for 20 min at 17,000 × g at 4 °C. Sucrose Gradient—300 μl of nuclear extract were loaded onto 12 ml of 10-30% linear sucrose gradient in buffer B (30 mm Tris-HCl, pH 7.6, 100 mm NaCl, 0.7 mm EDTA, 0.1 mm phenylmethylsulfonyl fluoride, and CompletePTMP protease inhibitors (Roche Applied Science)) and poured into UltraClear ultracentrifuge tubes (Beckman, no. 344059), using Hoefer SG15 gradient maker (Hoefer Scientific Instruments) and Pharmacia Biotech Pump P1. The probes were centrifuged using Sw41Ti rotor (Beckman) (35,000 rpm, 20 h, 4 °C). 1-ml fractions were collected manually through the hole made in the bottom of a tube. Analysis of Gradient Fractions—Proteins were trichloroacetic acid-precipitated from 700 μl of 1-ml gradient fraction, dissolved in 200 μl of 2× Laemmli, and analyzed by Western blot (30 μl for one assay) on 4-12% Bis-Tris NuPAGE Gel (Invitrogen). The primary antibodies used were as follows: mouse monoclonal antibody H1 (Santa Cruz Biotechnology, sc-8030) (1:500), monoclonal antibody H3 (Upstate Biotechnology, Inc., no. 05-499) (1:1000), rabbit polyclonal antibody H2A#618 (1:3000) from Dr. R. Glaser (Division of Genetic Disorders, Wadsworth Center, Albany, NY), polyclonal antibody PAR (Calbiochem) (1:4000), and polyclonal antibody GFP (1:1000-1:1500) (TP401, Torrey Pines Biolabs). The remaining 300 μl of each fraction was digested with 100 μg/ml proteinase K in 1% SDS at 50 °C for 2 h, 650 rpm in Thermomixer (Eppendorff). DNA was then recovered by phenol chloroform extraction, followed by ethanol precipitation with glycogen as a carrier. Pellet was dissolved in 40 μl of HB2BO, incubated with 2 μg of RNase A for 30 min at 37 °C, and analyzed on 1.2% agarose gel. Immunoprecipitation—For one immunoprecipitation reaction, 300 μl of nuclear extract was incubated with 60 μl of protein G-Sepharose 4B (Sigma P3296-5ML) on a rotating platform for 1 h at 4 °C.The beads were removed by spinning for 5 min at 15,000 × g.25 μg of anti-GFP polyclonal antibody (Torrey Pines, TP401) were added to the extract and incubated for 2 h or overnight on a rotating platform at 4 °C. Then 50 μl of protein G-Sepharose 4B were added to the extract and incubated for 2 h at 4 °C with rotation. The beads were washed five times for 3 min in 1.2 ml of the buffer (50 mm Tris-HCl, pH 7.5, 125 mm NaCl, 5% glycerol, 0.2% Nonidet P-40, 1.5 mm MgClB2B,BB25 mm NaF, 1 mm NaB3BVOB4B, 1 mm EDTA, and CompletePTMP protease inhibitors (Roche Applied Science)). Bound proteins were eluted by 100 μl of 2× Laemmli with heating at 90 °C for 5 min. Mass Spectrometry Analysis—Mass spectrometric identification of proteins was carried out as described in Ref. 24Bouwmeester T. Kim A. Ruffner H. Angrand P.O. Bergamini G. Croughton K. Cruciat C. Eberhard D. Gagneur J. Ghidelli S. Hopf C. Huhse B. Mangano R. Michon A.M. Schirle M. Schlegl J. Schwab M. Stein M.A. Bauer A. Casari G. Drewes G. Gavin A.C. Jackson D.B. Joberty G. Neubauer G. Rick J. Kuster B. Superti-Furga G. Nat. Cell Biol. 2004; 6: 97-105Crossref PubMed Scopus (880) Google Scholar. Complete lanes from protein gels were cut into slices (narrow for specific bands) and analyzed by liquid chromatography-tandem MS. The tandem MS data were analyzed using the SEQUESTP Pprogram. Protein complexes from four purifications were analyzed by liquid chromatography-tandem MS, and a total of 22 proteins were identified. It should be emphasized that it is essential to perform several purifications for a given bait protein to obtain a reliable view of its interaction network, because significant interactors are expected to be reproducibly identified in more than one experiment. In Vitro Interaction Assays—Histones and histone octamers were isolated or assembled according to Ref. 25Luger K. Kim T.J. Richmond T.J. Methods Enzymol. 1999; 304: 3-19Crossref PubMed Scopus (574) Google Scholar. Protein coupling to CnBr-activated Sepharose beads (GE Healthcare) and in vitro binding assays were adapted from (26Cirillo L.A. Kim F.R. Cuesta I. Friedman D. Jarnik M. Zaret K.S. Mol. Cell. 2002; 9: 279-289Abstract Full Text Full Text PDF PubMed Scopus (879) Google Scholar). Briefly, beads coupled to histone octamer (20 pmol), PARP1 (30 pmol, Trevigen), or individual histones (400 pmol) were washed once for 10 min in binding/washing buffer (10 mm Tris-HCl, pH 8, 140 mm NaCl, 3 mm DTT, and 0.1% Triton X-100). Washed beads were incubated with octamer (22.12 pmol), PARP1 (8 pmol), or rabbit IgG (8 pmol, Sigma) in binding/washing buffer for 20 min. The beads were then washed five times for 10 min in binding/washing buffer. All of the binding/washing was done at room temperature with gentle rotation. Full-length PARP1 and rabbit IgG were visualized on Western blots with anti-PARP1 (mouse monoclonal, 1:500, Serotec) and anti-rabbit horseradish peroxidase (1:3000, Jackson ImmunoResearch Labs), respectively. Anti-PARP1 C terminus (rabbit polyclonal, 1:1000) and anti-PARP1 N terminus (rabbit polyclonal, 1:1000) were gifts from Dr. Lee Kraus (Department of Molecular Biology and Genetics, Cornell University, Ithaca, NY). PARP1 Activity Assay—0.2 nmol of histones and/or 2.5 μg of endonuclease-digested plasmid DNA were combined with 5× PARP1 reaction buffer (0.05 unit/μl PARP1 enzyme (Trevigen), 500 μm NAD (Sigma), 500 mm Tris, pH 8, 50 mm MgCl2, and 5 mm DTT) in a final volume of 25 μl. PARP1 inhibition was achieved by the addition of 3-aminobenzamine (Sigma) to a final concentration of 12 mm. All of the reactions were carried out for 10 min at room temperature. PARP1 Protein Association with Chromatin Is Dynamic—To study PARP1 interaction with chromatin, we first analyzed the dynamic localization of this protein in vivo by using a FRAP assay. To visualize the PARP1 protein in Drosophila, we used the UAS/Gal4 system (27Brand A.H. Kim N. Development. 1993; 118: 401-415Crossref PubMed Google Scholar) for transgenic expression of PARP1-DsRed (encoding full-length, catalytically active PARP1) and for a contrasting control, PARPe-EGFP (encoding a naturally occurring, catalytically inactive form of PARP1) (Fig. 1A). Previously, we biologically validated those constructs by testing their ability to rescue a ParpCH1 mutation phenotype and by using immunofluorescence to assess recombinant protein localization to chromatin (2Tulin A. Kim A. Science. 2003; 299: 560-562Crossref PubMed Scopus (388) Google Scholar, 17Tulin A. Kim D. Spradling A.C. Genes Dev. 2002; 16: 2108-2119Crossref PubMed Scopus (178) Google Scholar). As references, we also made transgenic flies with core histone H2A-ECFP and linker histone H1-EYFP (Fig. 1A), which served as comparative controls defining the protein mobility of chromatin-associated proteins. As an additional reference for the mobility of a nucleoplasmic soluble protein, we made transgenic flies expressing PARG-EGFP (18Tulin A. Kim N.M. Menon A.K. Spradling A.C. Genetics. 2006; 172: 363-371Crossref PubMed Scopus (48) Google Scholar) (Fig. 1A). To express our transgenic constructs in Drosophila, we used the 69B-GAL4 driver (19Manseau L. Kim A. Brower D. Budhu A. Elefant F. Phan H. Philp A.V. Yang M. Glover D. Kaiser K. Palter K. Selleck S. Dev. Dyn. 1997; 209: 310-322Crossref PubMed Scopus (198) Google Scholar), which allows expression of recombinant protein ubiquitously without excess overproduction (supplemental Fig. S1). All of the recombinant proteins demonstrated exclusive nuclear localization in all tissues of the fruit fly (supplemental Fig. S1). All of the recombinant proteins except PARG-EGFP were also associated with chromatin. Previously, we reported that PARG is a soluble nucleoplasmic protein (18Tulin A. Kim N.M. Menon A.K. Spradling A.C. Genetics. 2006; 172: 363-371Crossref PubMed Scopus (48) Google Scholar). H1-YFP and H2A-CFP histones remain bound to chromatin during all stages of cell cycle, whereas PARPe-EGFP and PARP1-DsRed are partially excluded from mitotic chromosomes (supplemental Fig. S2). To explore whether the catalytic activity of PARP1 influences the dynamics of PARP1 protein interaction with chromatin, we compared the FRAP dynamics of PARPe-EGFP protein with those of full-length, enzymatically active PARP1-DsRed in Drosophila interphase nuclei. We co-expressed both recombinant PARPs in the ParpCH1 mutant animals (17Tulin A. Kim D. Spradling A.C. Genes Dev. 2002; 16: 2108-2119Crossref PubMed Scopus (178) Google Scholar) using Arm::Gal4 driver. The catalytically active PARP1-DsRed and inactive PARPe-EGFP demonstrated exactly the same localization profiles (Fig. 1B) and the same replacement rate (Fig. 1C and Table 1). This suggested that the catalytic domain of PARP1 is not involved in PARP1 protein interaction with chromatin. Based on this last result, we then used the PARPe-EGFP isoform to remove the potential for artifacts arising from the expression of catalytically active PARP1-DsRed, e.g. hyper-activation of the pADPr reaction targeting nonphysiological substrates. In the early stages of Drosophila development, catalytically inactive PARPe protein is expressed endogenously, and overexpression of it does not affect Drosophila development (17Tulin A. Kim D. Spradling A.C. Genes Dev. 2002; 16: 2108-2119Crossref PubMed Scopus (178) Google Scholar).TABLE 1Quantification of the fluorescence recovery after photobleachingRecombinant proteinT½PARP1-DsRed104 ± 6 sPARPe-EGFP101 ± 4 sPARG-EGFP3 ± 2 sH1-EYFP108 ± 4 sH2A-ECFP>20 min Open table in a new tab We then compared the PARPe-EGFP protein dynamics after photobleaching to those of histone H2A-ECFP, linker histone H1-EYFP, and the soluble nucleoplasmic protein PARG-EGFP (Fig. 1D and Table 1). As anticipated, the soluble PARG-EGFP demonstrated the highest recovery rate: 78% within 5 s. In contrast, the replacement rate for the control core histone H2A in chromatin was close to zero (Fig. 1D), whereas linker histone H1 showed a ∼49% replacement rate after 100 s of recovery (Fig. 1D), which is similar to previously reported values (28Misteli T. Kim A. Hock R. Bustin M. Brown D.T. Nature. 2000; 408: 877-881Crossref PubMed Scopus (520) Google Scholar). These data indicated that the PARP1 protein recovery kinetics was similar to that of H1 histone. A small, but reproducible difference is only observed during the first “fast” phase of recovery (Fig. 1D). During this phase the PARP1 protein recovery is more rapid, which suggested that the pool of soluble nucleoplasmic PARP1 is higher than the pool of soluble H1 protein. The deviation in the binding kinetics may also reflect differences in mechanisms of PARP1 protein and histone H1 interaction with nucleosomal arrays. PARP Proteins Are Continuously Exchanged between Chromatin Domains—We characterized the dynamics of PARPe-EGFP in different nuclear subcompartments: euchromatic, heterochromatic, and nucleolar. We defined heterochromatin on the basis of morphological criteria as a condensed block of chromatin attached to the nuclear lamina and associated with the nucleolus. In contrast to euchromatin, heterochromatin demonstrated a very low level of PARPe-EGFP fluorescence recovery (Fig. 1E), which might be attributed to low accessibility of compacted heterochromatin. Although the nucleolus is decondensed and accumulates PARPe-EGFP protein, PARPe-EGFP protein recovery to photobleached nucleoli is also minimal. This implies that the mechanism of PARPe-EGFP protein association with nucleolar chromatin may be different from that in other nuclear compartments. Next, we analyzed PARPe-EGFP protein dynamics in respect to chromatin subdomains, as noted above. We photobleached regions of euchromatin in a giant polyploid cell of Drosophila salivary gland expressing PARPe-EGFP and then recorded the recovery of fluorescence signal in the bleached area by time lapse imaging (Fig. 2A). PARPe-EGFP protein recovery had two distinct phases: 1) a fast phase, in which ∼50% of the fluorescent signal was recovered within 100 s after bleaching (Fig. 2A, graph) and 2) a “slow” phase, in which the signal was recovered up to ∼97% of starting levels during 15-20 min (not shown). These results suggested that, in the nucleus, most of the PARPe-EGFP molecules are bound to chromatin at any given time. Following this hypothesis, the pool of free soluble PARPe-EGFP is rapidly depleted for fast recovery, whereas the slow phase recruits PARPe-EGFP, which has dissociated from other chromatin domains. This hypothesis suggests that there is equilibrium of PARPe-EGFP protein association with different domains of chromatin and depletion of PARPe-EGFP protein from one locus leads to redistribution of PARPe-EGFP in the whole nucleus. To test this idea and better evaluate the kinetics of PARPe-EGFP protein exchange between chromatin subdomains, we bleached an extended rectangular area occupying approximately one-third of the total area of the nucleus (Fig. 2B). We compared the fluorescent signal within four distinct euchromatin subdomains, two (RO1 and RO4) localized outside of the bleached area and two bleached subdomains (RO2 and RO3). The recovery kinetics for the two bleached subdomains was similar to that observed in previous experiments. However, unbleached chromatin subdomains lost PARPe-EGFP fluorescence, whereas fluorescent intensity came to equilibrium in all four areas after ∼150 s (Fig. 2B). This observation directly demonstrated that PARPe-EGFP is continuously exchanged between chromatin regions in the nucleus. Based on the rapid exchange rate, our findings further indicate that the PARPe-EGFP protein is dynamic in its association with chromatin. However, the profile of PARPe-EGFP protein distribution among chromatin subdomains was very stable and reconstituted after recovery from bleaching (Fig. 2B). Thus, there must be high affinity landmarks for PARPe-EGFP binding on chromatin that maintains the stability of local PARPe-EGFP concentration in any given domain of chromatin. To identify these landmarks, we performed purification of PARPe-EGFP-containing protein complexes and identification of PARPe-EGFP protein partners using MS analysis. PARPe-EGFP Protein Co-purifies with Nucleosomal Core Histones—To identify PARPe-EGFP-chromatin targeting proteins, we performed co-immunoprecipitation experiments from a Drosophila stock with ubiquitous expression of the PARPe-EGFP transgenic construct. We purified protein complexes from nuclear extracts prepared from Drosophila pupae. Pupal extracts treated with micrococcal nuclease to produce mononucleosomes were immunoprecipitated with anti-GFP antibodies to collect PARPe-EGFP-associated complexes. As a control, extracts from wild-type flies were immunoprecipitated in parallel reactions to allow identification of proteins specifically interacting with PARPe-EGFP. Immunoprecipitates were analyzed by MS analysis. Based on this analysis, we identified 22 nuclear proteins that specifically interacted with the PARPe-EGFP protein (Table 2). Among these, only the nucleosomal core histones H4, H3, H2A, and H2B (Fig. 3A) were ubiquitous chromatin components. We did not identify H1 histone among the PARPe-EGFP-interacting proteins in multiple experiments, even though the interaction of PARP1 protein with linker histone H1 has been shown in vitro (29Kun E. Kim E. Ordahl C.P. J. Biol. Chem. 2002; 277: 39066-39069Abstract Full Text Full Text PDF PubMed Scopus (64) Google Scholar). The last result correlates with the observation that H1 and PARP1 are antagonists in the chromatin in vivo (3Kim M.Y. Kim S. Gevry N. Lis J. T. Kraus W. L. Cell. 2004; 119: 803-814Abstract Full Text Full Text PDF PubMed Scopus (456) Google Scholar). The interactions of the PARP1 protein with core histones were confirmed in experiments with immunoprecipitation of protein complexes with PARP1-DsRed protein as bait (not shown). Based on these results, together with our earlier data demonstrating the broad distribution of the PARP1 protein in chromatin, we hypothesized that the PARP1 protein interacts either directly or indirectly with nucleosomal particles.TABLE 2Proteins associated with the PARPe-EGFP protein in DrosophilaProtein nameClass of proteinNominal molecular weightMASCOT scoreMASCOT expectSequence coverage%P17763Retroviral genome polyprotein138,031700.004618Fbp1Fat body protein 1119,591730.001423PA
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