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PARP-1 Deficiency Increases the Severity of Disease in a Mouse Model of Multiple Sclerosis

2009; Elsevier BV; Volume: 284; Issue: 38 Linguagem: Inglês

10.1074/jbc.m109.013474

ISSN

1083-351X

Autores

Vimal Selvaraj, Mangala M. Soundarapandian, Olga Chechneva, Ambrose J. Williams, Maxim Sidorov, Athena M. Soulika, David Pleasure, Wenbin Deng,

Tópico(s)

Cell death mechanisms and regulation

Resumo

Poly(ADP-ribose) polymerase-1 (PARP-1) has been implicated in the pathogenesis of several central nervous system (CNS) disorders. However, the role of PARP-1 in autoimmune CNS injury remains poorly understood. Therefore, we studied experimental autoimmune encephalomyelitis (EAE), a model for multiple sclerosis in mice with a targeted deletion of PARP-1. We identified inherent physiological abnormalities in the circulating and splenic immune composition between PARP-1−/− and wild type (WT) mice. Upon EAE induction, PARP-1−/− mice had an earlier onset and developed a more severe EAE compared with WT cohorts. Splenic response was significantly higher in PARP-1−/− mice largely because of B cell expansion. Although formation of Th1 and Th17 effector T lymphocytes was unaffected, PARP-1−/− mice had significantly earlier CD4+ T lymphocyte and macrophage infiltration into the CNS during EAE. However, we did not detect significant differences in cytokine profiles between PARP-1−/− and WT spinal cords at the peak of EAE. Expression analysis of different PARP isozymes in EAE spinal cords showed that PARP-1 was down-regulated in WT mice and that PARP-3 but not PARP-2 was dramatically up-regulated in both PARP-1−/− and WT mice, suggesting that these PARP isozymes could have distinct roles in different CNS pathologies. Together, our results indicate that PARP-1 plays an important role in regulating the physiological immune composition and in immune modulation during EAE; our finding identifies a new aspect of immune regulation by PARPs in autoimmune CNS pathology. Poly(ADP-ribose) polymerase-1 (PARP-1) has been implicated in the pathogenesis of several central nervous system (CNS) disorders. However, the role of PARP-1 in autoimmune CNS injury remains poorly understood. Therefore, we studied experimental autoimmune encephalomyelitis (EAE), a model for multiple sclerosis in mice with a targeted deletion of PARP-1. We identified inherent physiological abnormalities in the circulating and splenic immune composition between PARP-1−/− and wild type (WT) mice. Upon EAE induction, PARP-1−/− mice had an earlier onset and developed a more severe EAE compared with WT cohorts. Splenic response was significantly higher in PARP-1−/− mice largely because of B cell expansion. Although formation of Th1 and Th17 effector T lymphocytes was unaffected, PARP-1−/− mice had significantly earlier CD4+ T lymphocyte and macrophage infiltration into the CNS during EAE. However, we did not detect significant differences in cytokine profiles between PARP-1−/− and WT spinal cords at the peak of EAE. Expression analysis of different PARP isozymes in EAE spinal cords showed that PARP-1 was down-regulated in WT mice and that PARP-3 but not PARP-2 was dramatically up-regulated in both PARP-1−/− and WT mice, suggesting that these PARP isozymes could have distinct roles in different CNS pathologies. Together, our results indicate that PARP-1 plays an important role in regulating the physiological immune composition and in immune modulation during EAE; our finding identifies a new aspect of immune regulation by PARPs in autoimmune CNS pathology. Poly(ADP-ribose) polymerase-1 (PARP-1) 2The abbreviations used are: PARPpoly(ADP-ribose) polymeraseCNScentral nervous systemEAEexperimental autoimmune encephalomyelitisMOGmyelin oligodendrocyte glycoproteinMHCmajor histocompatibility complexIFNγinterferon γILinterleukinMBPmyelin basic proteinCFAcomplete Freund's adjuvantHBSSHanks' balanced salt solutionPBSphosphate-buffered salineFITCfluorescein isothiocyanateFACSfluorescence-activated cell sorterqPCRquantitative PCRTNFαtumor necrosis factor-αTGFβtransforming growth factor-βiNOSinducible nitric-oxide synthaseICAMintercellular adhesion moleculeVCAMvascular cell adhesion moleculeDNMT1DNA methyltransferase 1CTCFCCCTC binding factorTh1 and Th2T helper type 1 and T helper type 2WTwild type. belongs to a family of enzymes that regulate several cellular processes by adding poly(ADP-ribose) polymers to specific proteins (1Chambon P. Weill J.D. Mandel P. Biochem. Biophys. Res. Commun. 1963; 11: 39-43Crossref PubMed Scopus (488) Google Scholar, 2Kim M.Y. Zhang T. Kraus W.L. Genes Dev. 2005; 19: 1951-1967Crossref PubMed Scopus (655) Google Scholar). Based on sequence homology, 18 PARP family members have been identified in the human genome (3Amé J.C. Spenlehauer C. de Murcia G. BioEssays. 2004; 26: 882-893Crossref PubMed Scopus (1231) Google Scholar). Of these, PARP-1 is the most abundantly expressed PARP in most cell types (2Kim M.Y. Zhang T. Kraus W.L. Genes Dev. 2005; 19: 1951-1967Crossref PubMed Scopus (655) Google Scholar), accounting for up to 85% of maximally activated cellular PARP activity (4Moroni F. Curr. Opin. Pharmacol. 2008; 8: 96-103Crossref PubMed Scopus (133) Google Scholar). Initially, PARP-1 function was associated only with DNA single-stranded break repair and maintenance of genomic integrity (5Masson M. Niedergang C. Schreiber V. Muller S. Menissier-de Murcia J. de Murcia G. Mol. Cell. 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In addition, excessive activation of PARP-1 has been shown to induce a form of caspase-independent cell death, which involves the nuclear translocation of the flavoprotein apoptosis-inducing factor from the mitochondria (16Yu S.W. Wang H. Poitras M.F. Coombs C. Bowers W.J. Federoff H.J. Poirier G.G. Dawson T.M. Dawson V.L. Science. 2002; 297: 259-263Crossref PubMed Scopus (1569) Google Scholar, 17Koh D.W. Dawson T.M. Dawson V.L. Pharmacol. Res. 2005; 52: 5-14Crossref PubMed Scopus (211) Google Scholar, 18Modjtahedi N. Giordanetto F. Madeo F. Kroemer G. Trends Cell Biol. 2006; 16: 264-272Abstract Full Text Full Text PDF PubMed Scopus (265) Google Scholar). This type of cell death by PARP-1 activation, termed "parthanatos," (19Andrabi S.A. Dawson T.M. Dawson V.L. Ann. N.Y. Acad. Sci. 2008; 1147: 233-241Crossref PubMed Scopus (240) Google Scholar) has been implicated as an important mediator of various central nervous system (CNS) disorders (20Kauppinen T.M. Chan W.Y. Suh S.W. Wiggins A.K. Huang E.J. Swanson R.A. Proc. Natl. Acad. Sci. U.S.A. 2006; 103: 7136-7141Crossref PubMed Scopus (178) Google Scholar). Therefore, several studies have examined PARP-1 as a target for therapeutic intervention (21Decker P. Muller S. Curr. Pharm. Biotechnol. 2002; 3: 275-283Crossref PubMed Scopus (209) Google Scholar, 22Kauppinen T.M. Suh S.W. Berman A.E. Hamby A.M. Swanson R.A. J. Cereb. Blood Flow Metab. 2009; 29: 820-829Crossref PubMed Scopus (77) Google Scholar, 23Graziani G. Szabó C. Pharmacol. Res. 2005; 52: 109-118Crossref PubMed Scopus (127) Google Scholar, 24Virág L. Szabó C. Pharmacol. Rev. 2002; 54: 375-429Crossref PubMed Scopus (1226) Google Scholar), and PARP-1 inhibitors are currently the focus of several clinical trials (25Helleday T. Petermann E. Lundin C. Hodgson B. Sharma R.A. Nat. Rev. Cancer. 2008; 8: 193-204Crossref PubMed Scopus (1244) Google Scholar). poly(ADP-ribose) polymerase central nervous system experimental autoimmune encephalomyelitis myelin oligodendrocyte glycoprotein major histocompatibility complex interferon γ interleukin myelin basic protein complete Freund's adjuvant Hanks' balanced salt solution phosphate-buffered saline fluorescein isothiocyanate fluorescence-activated cell sorter quantitative PCR tumor necrosis factor-α transforming growth factor-β inducible nitric-oxide synthase intercellular adhesion molecule vascular cell adhesion molecule DNA methyltransferase 1 CCCTC binding factor T helper type 1 and T helper type 2 wild type. Mice lacking PARP-1 (PARP-1−/−) have also shown significant protection against various forms of CNS injury (26Kauppinen T.M. Neurochem. Int. 2007; 50: 954-958Crossref PubMed Scopus (37) Google Scholar). These mice are resistant to neuronal cell death after ischemia-reperfusion injury and neurodegeneration (27Goto S. Xue R. Sugo N. Sawada M. Blizzard K.K. Poitras M.F. Johns D.C. Dawson T.M. Dawson V.L. Crain B.J. Traystman R.J. Mori S. Hurn P.D. Stroke. 2002; 33: 1101-1106Crossref PubMed Scopus (116) Google Scholar, 28Chiarugi A. Pharmacol. Res. 2005; 52: 15-24Crossref PubMed Scopus (76) Google Scholar). Furthermore, PARP-1 deficiency is suggested to alter immune responses, as PARP-1−/− mice showed resistance to lipopolysaccharide-induced endotoxic shock (29Oliver F.J. Ménissier-de Murcia J. Nacci C. Decker P. Andriantsitohaina R. Muller S. de la Rubia G. Stoclet J.C. de Murcia G. EMBO J. 1999; 18: 4446-4454Crossref PubMed Scopus (540) Google Scholar). This immunomodulation has been suggested to be a result of PARP-1 regulation of transcriptional networks involved in immune cell activation and inflammatory cytokine production (30Saenz L. Lozano J.J. Valdor R. Baroja-Mazo A. Ramirez P. Parrilla P. Aparicio P. Sumoy L. Yélamos J. BMC Genomics. 2008; 9: 171Crossref PubMed Scopus (42) Google Scholar). Modulation of the activity of transcription factors, such as nuclear factor-κB (NFκB) and nuclear factor of activated T cells (NFAT) by PARP-1 has been reported in T lymphocytes (29Oliver F.J. Ménissier-de Murcia J. Nacci C. Decker P. Andriantsitohaina R. Muller S. de la Rubia G. Stoclet J.C. de Murcia G. EMBO J. 1999; 18: 4446-4454Crossref PubMed Scopus (540) Google Scholar, 31Kameoka M. Ota K. Tetsuka T. Tanaka Y. Itaya A. Okamoto T. Yoshihara K. Biochem. J. 2000; 346 (Pt 3): 641-649Crossref PubMed Scopus (106) Google Scholar, 32Valdor R. Schreiber V. Saenz L. Martinez T. Muñoz-Suano A. Dominguez-Villar M. Ramirez P. Parrilla P. Aguado E. García-Cózar F. Yélamos J. Mol. Immunol. 2008; 45: 1863-1871Crossref PubMed Scopus (64) Google Scholar). Moreover, T cell-dependent antibody responses are also found to be markedly reduced in PARP-1−/− mice (33Ambrose H.E. Willimott S. Beswick R.W. Dantzer F. de Murcia J.M. Yelamos J. Wagner S.D. Immunology. 2009; 127: 178-186Crossref PubMed Scopus (35) Google Scholar). In addition to immune activation, there is also evidence indicating that PARP-1 could be required in earlier stages of immune system maturation. For example, in vitro experiments have suggested that PARP-1 is critical for the functional maturation of dendritic cells (34Aldinucci A. Gerlini G. Fossati S. Cipriani G. Ballerini C. Biagioli T. Pimpinelli N. Borgognoni L. Massacesi L. Moroni F. Chiarugi A. J. Immunol. 2007; 179: 305-312Crossref PubMed Scopus (56) Google Scholar). Furthermore, activation and proinflammatory gene expression in resident immune cells in the brain (microglia and astrocytes) have also been found altered in vitro by the deficiency of PARP-1 (11Ha H.C. Proc. Natl. Acad. Sci. U.S.A. 2004; 101: 5087-5092Crossref PubMed Scopus (57) Google Scholar, 35Kauppinen T.M. Swanson R.A. J. Immunol. 2005; 174: 2288-2296Crossref PubMed Scopus (157) Google Scholar) or in the presence of PARP inhibitors (35Kauppinen T.M. Swanson R.A. J. Immunol. 2005; 174: 2288-2296Crossref PubMed Scopus (157) Google Scholar, 36Phulwani N.K. Kielian T. J. Neurochem. 2008; 106: 578-590Crossref PubMed Scopus (40) Google Scholar). In animal models, administration of putative PARP-1 inhibitors, such as 6(5H)-phenanthridinone, benzamide, PJ-34, and 5-aminoisoquinolinone, has been shown to ameliorate clinical signs of experimental autoimmune encephalomyelitis (EAE) (37Chiarugi A. Br. J. Pharmacol. 2002; 137: 761-770Crossref PubMed Scopus (82) Google Scholar, 38Scott G.S. Kean R.B. Mikheeva T. Fabis M.J. Mabley J.G. Szabó C. Hooper D.C. J. Pharmacol. Exp. Ther. 2004; 310: 1053-1061Crossref PubMed Scopus (68) Google Scholar), experimental autoimmune arthritis (39Gonzalez-Rey E. Martínez-Romero R. O'Valle F. Aguilar-Quesada R. Conde C. Delgado M. Oliver F.J. PLoS ONE. 2007; 2: e1071Crossref PubMed Scopus (39) Google Scholar) and experimental autoimmune diabetes (40Suarez-Pinzon W.L. Mabley J.G. Power R. Szabó C. Rabinovitch A. Diabetes. 2003; 52: 1683-1688Crossref PubMed Scopus (38) Google Scholar). These effects have been suggested to be caused mainly by attenuation of inflammatory responses involving multiple immune cell types (41Masutani M. Nakagama H. Sugimura T. Cell. Mol. Life Sci. 2005; 62: 769-783Crossref PubMed Scopus (74) Google Scholar). Although PARP-1 is believed to be the primary target in these studies, the specificity of these inhibitors on the different PARP isozymes has not been completely examined (42Nottbohm A.C. Hergenrother P.J. Huang Z. Drug Discovery Research: New Frontiers in the Post-Genomic Era. John Wiley and Sons, Hoboken, NJ2007: 163-185Crossref Scopus (6) Google Scholar). Moreover, recent evidence suggests that there may be distinct but cooperative roles for PARP-1 and other PARP family members in their immunomodulatory effects (36Phulwani N.K. Kielian T. J. Neurochem. 2008; 106: 578-590Crossref PubMed Scopus (40) Google Scholar). Despite all of these previous studies, the specific role of PARP-1 in CNS immune injury seen in multiple sclerosis remains largely unclear. In this study, we examined the autoimmune response and progression of EAE in PARP-1−/− mice. Our results unexpectedly demonstrated that the deficiency of PARP-1 markedly alters the immune phenotype, leading to an exacerbated EAE in PARP-1−/− mice. All chemicals were purchased from Sigma-Aldrich unless otherwise specified. New England Peptide (Gardner, MA) synthesized the peptide containing amino acids 35–55 of myelin oligodendrocyte glycoprotein (MOG) used in this study. Antibodies against CD4, CD8, B220, CD25, CD11c, CD11b, CD45, MHC Class II, IFNγ, and IL-17 were purchased from BD Biosciences, CD14 and F4/80 were from eBioscience (San Diego), CD68 was from AbD Serotec (Raleigh, NC), myelin basic protein (MBP) was from Novus Biologicals (Littleton, CO), Iba1 was from Wako Pure Chemical Industries (Osaka, Japan), and AlexaFluor 488- and 555-conjugated secondary antibodies were from Invitrogen. Liberase RI and DNase I were purchased from Roche Applied Science. Percoll was from GE Healthcare. MACS® reagents for CD11c+ cell purification were from Miltenyi Biotec (Bergisch Gladbach, Germany). Quantitative PCR reagents were purchased from Applied Biosystems (Foster City, CA). Breeding pairs with a homozygous deletion of the PARP-1 gene (129S-Parp1tm1Zqw/J) (43Wang Z.Q. Auer B. Stingl L. Berghammer H. Haidacher D. Schweiger M. Wagner E.F. Genes Dev. 1995; 9: 509-520Crossref PubMed Scopus (714) Google Scholar) and wild type (WT) controls in the same 129S genetic background (129S1/SvImJ) were purchased from The Jackson Laboratories (Bar Harbor, ME) and were bred and genotyped for the experiments. Animals were maintained in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals, and all experiments were approved by the Institutional Animal Care and Use Committee of the University of California, Davis. In 10-week-old mice, EAE was induced by injecting an emulsion of 300 μg of MOG peptide in complete Freund's adjuvant (CFA) subcutaneously on either hind flank as two injections. In addition, 250 ng of pertussis toxin was injected intraperitoneally on the same day as MOG-CFA, and another dose was administered after 48 h. Body weights of mice were recorded before MOG-CFA injection and then continuously at 2-day intervals. Disease development was monitored daily, and the severity of clinical signs was scored based on a standard neurological scoring system for EAE as follows: 1, limp tail or waddling gait; 2, limp tail and ataxia; 2.5, single limb paresis and ataxia; 3, double limb paresis; 3.5, single limb paralysis and paresis of second limb; 4, full paralysis of two limbs; 4.5, moribund; 5, dead. Scoring was performed in a blind fashion, and all animals were genotyped again for confirmation at the end of each experiment. Blood was collected by cardiac puncture under anesthesia (100 mg/kg ketamine and 10 mg/kg xylazine, intraperitoneally) and placed in EDTA-coated vacutainers (BD Biosciences Pharmingen) and stored at 4 °C. Samples were analyzed within 2 to 3 h after collection using the Hemavet multispecies hematological analyzer 850FS (Drew Scientific, Waterbury, CT) calibrated for use in the mouse. Measurements for erythrocyte, total leukocytes, neutrophils, lymphocytes, monocytes, eosinophils, basophils, platelets, hemoglobin, mean corpuscular volume, mean corpuscular hemoglobin, and mean corpuscular hemoglobin concentration were carried out. A manual differential count on Giemsa-stained blood smears was also performed simultaneously to validate automated results. Sample names were coded, and manual counts were performed in a blind fashion. Spleens were removed from either anesthetized mice before perfusion or immediately after euthanasia in a CO2 chamber and placed in Hanks' balanced salt solution (HBSS) on ice. They were then sliced into small pieces (∼1 mm3) in a Petri dish using surgical blades, and a single cell suspension was prepared by gently forcing the pieces through a 100-μm cell strainer. The resulting cell suspension was then centrifuged and resuspended in red blood corpuscle lysis buffer (Roche Applied Science), allowed to shake at room temperature for 10 min, and washed with RPMI lymphocyte culture medium (RPMI 1640 containing 10% fetal bovine serum and 1 mm sodium pyruvate) by centrifugation and resuspension at 4 °C. Isolated splenocytes were subsequently counted to estimate total cell numbers and prepared for either flow cytometry or cell culture as described below. To examine CD4+ T cell subsets expressing IFNγ (Th1 cells) and IL-17 (Th17 cells), we performed an in vitro treatment of lymphocyte preparations with MOG. For this specific assay, mice were sampled at days 7 and 10 of EAE when there was a high peripheral lymphocyte response. Spleens were processed as described above; but in addition to spleens, lymph nodes (superficial cervicals, axillary, brachial, mesenteric, and inguinal) were also included in the cell isolation. For culture, 3 × 105 cells were plated on Costar® ultra-low-binding 96-well plates (Corning Life Sciences, Lowell, MA). MOG was added at a final concentration of 50 μg/ml, and cells were incubated in a 37 °C incubator with 5% CO2 for 20 h. During the last 5 h of incubation, cells were exposed to brefeldin A (Golgi Stop®, BD Biosciences) to facilitate/enhance intracellular staining by retaining cytokines within the cells. Controls without MOG treatment, to visualize background, and positive controls using phorbol myristyl acetate (50 ng/ml) and ionomycin (750 ng/ml) during the last 2 h of incubation, to evaluate the viability and positive response of the CD4+ T lymphocytes in culture, were performed. At the end of incubation, cells were collected and processed for flow cytometry. Mouse spleens (three pooled for each sample) were collected in RPMI medium containing 0.5 Wünsch units/ml Liberase RI (Roche Applied Science) and 14 μg/ml DNase I (Roche Applied Science). Spleens were then sliced into small pieces using surgical blades and incubated with continuous shaking for 30 min at room temperature. During incubation, spleen pieces were triturated every 10 min by using fire-polished Pasteur pipettes. At the end of incubation, EDTA (10 mm final concentration) was added to stop the enzymatic reaction. Red blood corpuscle lysis was performed as described above, and cells were subsequently separated in a step Percoll gradient. In brief, cells were resuspended in 40% Percoll in RPMI medium with 2 mm EDTA, overlaid on 70% Percoll in the same medium, and centrifuged at 2,800 rpm at 4 °C for 25 min. Cells at the interface were collected with a Pasteur pipette into a separate tube, washed twice, and resuspended in RPMI medium with 2 mm EDTA. Cells were then counted and incubated with rat anti-mouse CD3 and CD19 antibodies for 30 min. Cells were subsequently washed and resuspended in phosphate-buffered saline (PBS) containing 0.1% bovine serum albumin and 2 mm EDTA, added to a tube of equilibrated sheep anti-rat Biomag beads (Qiagen, Valencia, CA), and incubated for 30 min at 4 °C with continuous rotary mixing. After incubation, the Biomag bead-bound cells were removed by magnetic separation, and the non-bound cells were processed for magnetic sorting (MACS®, Miltenyi Biotec) using CD11c+ selection. In brief, cells were resuspended in PBS containing 0.5% bovine serum albumin and 2 mm EDTA and incubated with FcR block (eBioscience) for 10 min at 4 °C. Cells were then incubated with CD11c-specific MACS® beads for 15 min in a rotary mixer at 4 °C. The incubation suspension was then loaded on an equilibrated MACS LS column; bound cells were washed three times and eluted. Cells were then counted, and the purity of each preparation was tested using flow cytometry for CD11c+ cells as described below; only samples of >90% purity were used for further experiments. To examine the antigen uptake potential of PARP-1−/− dendritic cells, we performed specific assays for phagocytosis and macropinocytosis. In this experiment, purified dendritic cells were plated at a density of 3 × 105 cells on Costar® ultra-low-binding 96-well plates. For the phagocytosis assay, FITC-conjugated albumin was added at a final concentration of 20 μg/ml. For the macropinocytosis assay, FITC-conjugated dextran was added at a final concentration of 1 mg/ml. Cells under both conditions were incubated at 37 °C with 5% CO2 for 1 h. Negative controls without albumin or dextran and a more stringent temperature-based control in which cells exposed to either FITC-albumin or FITC-dextran were incubated at 4 °C for 1 h were also set up concurrently. At the end of incubation, the cells were processed for flow cytometry to estimate uptake by quantifying FITC fluorescence along with the detection of CD11c+ cells as described below. Transcardial perfusion was carried out under anesthesia with 20 ml of sterile PBS to clear all vascular blood cells in the CNS. Immediately after perfusion, brain and spinal cord were removed by dissection in a laminar flow unit and placed in HBSS (with 2 mm EDTA) on ice. Collected tissues were chopped into small pieces (∼1 mm3) in a Petri dish with digestion buffer (HBSS with 0.5 Wünsch Units/ml Liberase RI and 14 μg/ml DNase I) using a sterile scalpel blade. The tissue pieces were then incubated in a 37 °C incubator with 5% CO2 and shaking at 100 rpm/min for 25 min. After incubation, the resulting isolated cells were passed through a 100-μm cell strainer and rinsed twice with HBSS. Cells were then pelleted by centrifugation at 1500 rpm for 10 min at 4 °C and subsequently separated in a step Percoll gradient as described for dendritic cells. Cells at the interface were collected with a Pasteur pipette into a separate tube and washed twice by centrifugation and resuspension in HBSS. These cells were counted and prepared for either flow cytometry or cell culture. Before antibody labeling, cells were spun down and resuspended in FACS buffer (PBS containing 1% bovine serum albumin with 0.1% azide). To reduce nonspecific antibody binding, surface Fc receptors were blocked using FcR block for 30 min. Surface labeling using fluorophore-conjugated antibodies (CD4, CD8, B220, CD14, F4/80, MHC Class II, CD11c, CD25, CD11b, CD45, and isotype controls) was carried out directly in FACS buffer with a 2-h incubation at 4 °C. After incubation, cells were washed by centrifugation and resuspension in FACS buffer to remove unbound antibodies. Cells were subsequently either fixed with 1% formaldehyde solution for analysis or permeabilized for intracellular labeling (IFNγ, IL-17, CD68, and isotype controls). For intracellular labeling for IFNγ and IL-17, cells were fixed and permeabilized using the BD Cytofix and Cytoperm buffers (BD Biosciences). Fluorophore-conjugated IFNγ and IL-17 were then added into BD Cytoperm buffer and allowed to label overnight at 4 °C. The next day, cells were washed with BD Cytoperm buffer, fixed using 1% formaldehyde solution, and analyzed. For intracellular labeling of CD68, cells were fixed and permeabilized using Leucoperm (AbD Serotec) and processed as described above for IFNγ and IL-17. Data acquisition from labeled cells was performed in a CyAN flow cytometer (Dako Cytomation, Carpinteria, CA). Isotype and autofluorescence controls were examined for each assay. Acquired data were subsequently analyzed by calculating proportions/percentages and average intensities using FlowJo software (Treestar, Inc., San Carlos, CA). Absolute numbers of cellular subtypes were derived from percentages using the total cell numbers counted after isolation. Spinal cords were dissected from transcardial perfusion-fixed animals and placed in 4% neutral buffered formalin for 24 h. Samples were then embedded in OCT compound (Sakura Finetek, Torrance, CA) and frozen in liquid nitrogen-cooled isopentane. Sections (10 μm thick) were prepared in a cryotome and stored at −20 °C until use. For routine histological examination, sections were stained with hematoxylin and eosin. For localization of MBP and Iba1, slides were post-fixed in 4% neutral buffered formalin containing 0.3% Triton X-100 for 30 min. Nonspecific binding of antibodies was blocked using 10% normal goat serum, and then samples were incubated with primary antibodies against either MBP or Iba1 overnight at 4 °C. Slides were then washed in PBS and incubated with fluorophore-conjugated secondary antibodies (1:500). Fluorescent nuclear counterstaining was carried out using 4,6-diamidino-2-phenylindole (1 μm final concentration; Invitrogen) for 10 min. Slides were then washed in PBS and mounted using ProLong Gold mounting medium (Invitrogen). Images were acquired on an Olympus BX61-DSU microscope (Olympus Corp., Melville, NY) using a Hamamatsu ORCA-ER camera (Hamamatsu Photonics, Shizuoka, Japan). To examine the relative expression of immune cytokines, adhesion molecules, PARP isozymes and epigenetic regulators during EAE in PARP-1−/− and WT mice, we used a real-time quantitative reverse transcriptase-polymerase chain reaction (qPCR) approach. Total RNA was extracted from lumbar spinal cords isolated (after clearing vascular blood cells as described earlier) from non-immunized mice (Day 0) and from mice at their peak of clinical EAE scores (Day 14) using a Qiagen RNeasy lipid mini kit following the standard protocol. For quality control, RNA purity was verified using the OD260/280 ratio to be between 1.8 and 2.0. Total RNA (1.0 μg) was reverse-transcribed to cDNA using MultiscribeTM reverse transcriptase (Applied Biosystems). Subsequent qPCR reactions for TNFα, IFNγ, IL-1β, IL-2, IL-4, IL-5, IL-10, iNOS, and TGFβ were performed in triplicate on a Roche Lightcycler 480 using SYBR Green Master Mix (Roche Applied Science) with specific primers (supplemental Table 1) either designed or selected from published literature (44Overbergh L. Valckx D. Waer M. Mathieu C. Cytokine. 1999; 11: 305-312Crossref PubMed Scopus (522) Google Scholar, 45Cho I.H. Hong J. Suh E.C. Kim J.H. Lee H. Lee J.E. Lee S. Kim C.H. Kim D.W. Jo E.K. Lee K.E. Karin M. Lee S.J. Brain. 2008; 131: 3019-3033Crossref PubMed Scopus (134) Google Scholar). The specificity of products from each primer set was validated by analyzing melting curves (Tm). To examine the induction of PARP-1 and relative changes in PARP-2, PARP-3, intercellular adhesion molecule-1 (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1), DNA methyltransferase 1 (DNMT1), and CCCTC binding factor (CTCF) expression during EAE in WT and PARP-1−/− mice, qPCR reactions were performed in duplicate using validated TaqMan gene expression assays (Applied Biosystems), PARP-1 (Mm00500171_g1), PARP-2 (Mm01319555_m1), PARP-3 (Mm00467486_m1), ICAM-1 (Mm 01175876_g1), VCAM-1 (Mm01320970_m1), DNMT1 (Mm01151065_g1), CTCF (Mm004840 27_m1), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH; Mm99999915_g1). For both SYBR Green and TaqMan primers, validation was performed by calculating qPCR efficiencies by amplification of a standardized dilution series and constructing a relative efficiency plot comparing target and reference ΔCp values to ensure that the absolute slope of fit line was less than 0.1 (46Livak K.J. Schmittgen T.D. Methods. 2001; 25: 402-408Crossref

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