PARP-2 controls adipocyte differentiation and adipose tissue function through the regulation of the activity of the RXR/PPAR γγγγ heterodimer

polymerase-1; factor-1;

The peroxisome proliferator activated receptor-γ γ γ γ (PPARγ γ γ γ, NR1C3) ) ) ) in complex with the retinoid X receptor (RXR) plays a central role in white adipose tissue (WAT) differentiation and function, regulating the expression of key WAT proteins. In this report we show that poly(ADP-ribose) polymerase-2 (PARP-2), also known as an enzyme participating in the surveillance of the genome integrity, is a member of the PPARγ γ γ γ/RXR transcription machinery. PARP-2 -/mice accumulate less WAT, characterized by smaller adipocytes. In the WAT of PARP-2 -/mice the expression of a number of PPARγ γ γ γ target genes is reduced despite the fact that PPARγ γ γ γ1 and γ γ γ γ2 are expressed at normal levels. Consistent with this, PARP-2 -/mouse embryonic fibroblasts fail to differentiate to adipocytes. In transient transfection assays, PARP-2 siRNA decreases basal activity and liganddependent activation of PPARγ γ γ γ, whereas PARP-2 overexpression enhances the basal activity of PPARγ γ γ γ, although it does not change the maximal ligand-dependent activation. In addition, we show a DNA-dependent interaction of PARP-2 and PPARγ γ γ γ/RXR heterodimer by chromatin immunoprecipitation. In combination, our results suggest that PARP-2 is a novel cofactor of PPARγ γ γ γ activity.
Adipose tissue is composed of adipocytes that store energy in the form of triglycerides. Excessive accumulation of white adipose tissue (WAT) leads to obesity, while its absence leads to lipodystrophic syndromes. The peroxisome proliferator-activated receptorγ (PPARγ, NR1C3) is the main protein orchestrating the differentiation and function of WAT, as evidenced by the combination of in vitro studies, the analysis of mouse models and the characterization of patients with mutations in the human PPARγ gene (1), (2). PPARγ acts as heterodimer with the retinoid X receptor (RXR) (3). The PPARγ/RXR receptor dimer is involved in the transcriptional control of energy, lipid, and glucose homeostasis (4); (5). The actions of PPARγ are mediated by two protein isoforms, the widely expressed PPARγ 1 and adipose tissue-restricted PPARγ 2 , both produced from a single gene by alternative splicing and differing only by an additional 28 amino acids in the NH 2terminus of PPARγ 2 (6); (3).
PPARγ is activated by binding of small lipophilic ligands, mainly fatty acids, derived from nutrition or metabolic pathways, or synthetic agonists, like the anti-diabetic thiazolidinediones (2), (7); (8). Docking of these ligands in the ligand binding pocket alters the conformation of PPARγ, resulting in transcriptional activation subsequent to the release of corepressors and the recruitment of coactivators. Many corepressors and coactivators have been described such as the nuclear receptor corepressor (N-CoR) and the steroid receptor coactivators (SRCs), also known as p160 proteins (9); (10) (11). These corepressors and coactivators determine transcriptional activity by altering chromatin structure via enzyme such as histone deacetylases (HDACs) and histone acetyltransferases (CBP/p300). Other mechanisms include DNA-methylation, ATP-dependent remodeling, protein phosphorylation, SUMOylation and ubiquitinylation and poly(ADP-ribosyl)ation.
Poly(ADP-ribose) polymerase-2 (PARP-2) was described by Ame et al in 1999 as a 66.2 kDa nuclear protein with poly(ADP-ribosyl)ating activity. Through its DNA-binding domain in the Nterminus (amino acid 1-62), PARP-2 can bind to DNase I treated DNA and to aberrant DNA forms and its subsequent activation results in poly(ADP-ribose) polymer (PAR) formation (12). According to the general scheme of PARP activation, the active enzyme catalyses the polymerisation of PAR onto different acceptor proteins and itself using NAD + as a substrate (13). PARP-2 shares a similar catalytic domain (amino acid 202-593) as poly(ADP-ribose) polymerase-1 (PARP-1) (14), the founding member of the PARP family, though PARP-2 has a smaller reaction velocity compared to PARP-1 (12).
PARP-2 has multiple in vivo functions comprising DNA surveillance and DNA repair processes (reviewed in (15)), spermatogenesis (16), (17), inflammation and oxidative injury (18), (19), (20). Most of these functions are accomplished through protein-protein interactions. In PARP-2, the interaction platforms can be mapped to the DNA-binding domain and to the domain E (amino acid 63-202) (21); (22); (23); (24); (25). A role for PARP-2 in the regulation of transcription has already been described. In lung epithelial cells PARP-2 interacts with thyroid transcription factor-1 (TTF1). TTF1 is a homeodomaincontaining transcription factor of the Nkx-2 family. In these cells, PARP-2 regulates the expression of the surfactant protein-B by affecting TTF1 activity (25). In this study we show that PARP-2 affects the transcriptional activity of PPARγ both in vitro and in vivo.

EXPERIMENTAL PROCEDURES
Materials -All chemicals were from Sigma-Aldrich (St. Louis, MO, USA) unless stated otherwise. Animals -PARP-2 -/mice and their wildtype (WT) littermates (26) coming from heterozygous crossings were used. Mice were housed separately, had ad libitum access to water and chow, and were kept under a 12 h dark-light cycle. The animals were killed at the age of 7 months by cervical dislocation after 4 hours of fasting and tissues were collected. Cell culture -3T3-L1 cells were maintained in DMEM (Invitrogen, Carlsbad, CA), 10% NCS (Invitrogen), Gentamicine (Invitrogen) and HEK and mouse embryonic fibroblasts (MEFs) in DMEM, 10% FCS (Adgenix, Voisins le Bretonneux, France), Gentamicine (Invitrogen). The 3T3-L1 cells were maintained subconfluent. MEF preparation and differentiation -MEFs were prepared from embryos as described elsewhere (26;26). For the differentiation studies 4x10 5 MEFs were seeded in 12 well plates and were maintained in DMEM, 10% FCS. The medium was changed every 2 days until confluence. The cells were maintained at confluency for 2 days. Cells were then differentiated in DMEM, 10% NCS, 5 µM troglitazone (TZD), 5 µM dexamethasone (Dex), 500 µM IBMX and 10 µg/ml insulin (later defined as differentiation mix), while the control cells received DMEM, 10% FCS and DMSO as vehicle. The medium with the differentiation mix was replaced every 2 days and the cells were differentiated for 8 days. Control cells after confluency were cultured in DMEM+10% FCS containing only vehicle (DMSO 0.21%). DNA constructs -To create an siRNA expressing construct, double stranded oligonucleotides were cloned into the pSuper vector (for sequences see Table 1.) (27). The oligonucleotides siPARP-2sense and siPARP-2antisense (containing the siRNA sequence), as well as the control scrPARP-2sense and scrPARP-2antisense (scrambled version of the siRNA sequence) respectively were annealed in annealing buffer (150mM NaCl, 1mM EDTA, 50 mM Hepes pH. 8.0). The resulting duplexes carried BglII and HindIII sites and were cloned into pSuper using these sites resulting in pSuper-siPARP-2 (oligos siPARP-2sense + siPARP-2antisense) and pSuper-scrPARP-2 (oligos scrPARP-2sense + scrPARP-2antisense). An EcoRV/SmaI fragment encoding mouse PARP-2 was isolated from pBC-mPARP-2 ((23)) and inserted into the SnaBI site of pBABEpuro (Addgene, Cambridge, MA), giving the pBABE-mPARP-2 vector. All other constructs pGL3-(J wt ) 3 TKluc reporter construct (28), pSG-PPARγ 2 , (3), pSG5-PPARα (29), pSG5-PPARβ (30), pCMX-ERβ, and vitellogeninA2-ERE-TKLuc (ER-luc) (31) were described before. The pCMV-βGal was used to control the transfection efficiency. Transfections -Transfections were preformed either by the BES buffered saline (BBS) method (26) or by JetPei (Polyplus Transfections, Illkirch, France). Luciferase activity measurement -3x10 5 HEK cells were seeded in 6 well plates and were transfected with pSuper-siPARP-2, pSuper-scrPARP-2, pBabe or pBabe-PARP-2 using the BBS method. Two days later the cells were once more transfected with the constructs mentioned above. Cells were transfected 24 hours later with 0.6 µg pSuper-siPARP-2/pSuper-scrPARP-2/pBabe/pBabe-PARP-2; 0.4 µg β−galactosidase expression plasmid; 1 µg pSG-PPARα/pSG-PPARβ/ pSG-PPARγ 2 /pCMX-ERβ expression vector; 1 µg PPAR-/ER-responsive construct. Six hours after transfection, cells were scraped and luciferase activity was determined. For the determination of PPAR activity, just before transfection, cells were washed in serum-free DMEM medium, and the transfection was carried out in DMEM + 10% fat-free serum. As ligand we used, fenofibrate (FF) (50 µM), mono-ethyl-hexyl-phthalate (MEHP) (100 µM), TZD (5µM), and β-estradiol (10 µM). After 6 hours of transfection cells were washed with PBS, scraped and stored at -80 o C. Luciferase assay was carried out by standard procedures. Luciferase activity was expressed as luciferase activity/βgalactosidase activity. Nile-Red flow cytometry -To assess the extent of MEF differentiation, cytosolic triglyceride content was assessed by determining Nile-Red uptake (modified from (32)) followed by flow cytometry using a FACSCalibur machine (BD, San Diego, CA, USA). Cells were harvested by adding Trypsin-EDTA and the detached cells were stained with Nile-Red (20 µg/ml, 5 min.). Cells were subjected to flow cytometric analysis with 10000 events collected for each sample, each measurement point was repeated in 4 parallel replicates. Samples for each cell line were normalized against the nondifferentiated cells of the same line. The rate of differentiation was expressed as the percentage of the differentiated cells vs. total number of cells. SDS-PAGE -Western blot -Cells were lysed in lysis buffer (50 mM Tris, 500 mM NaCl, 1 mM EDTA, 1% NP40, 1 mM PMSF, protease inhibitor cocktail, pH 8.0). Proteins were separated by SDS-PAGE and transferred onto nitrocellulose membranes. For the detection of PARP-2, a polyclonal rabbit antibody was used 1:2000, Alexis, Lausen, Switzerland) and actin was as detected using a rabbit polyclonal antibody (Sigma, 1:200 to the manufacturer's instructions. RNA was treated with DNase and 2 µg RNA was used for reverse transcription (RT). cDNA was purified on Qiaquick PCR cleanup columns (Qiagen, Valencia, CA, USA). 50X diluted cDNA was used for quantitative PCR (qPCR) reactions. The qPCR reactions were preformed using the LightCycler system (Roche, Basel, Switzerland) and a qPCR supermix (Qiagen) with the primers summarized in Table 2. Chromatin immunoprecipitation -Chromatin immunoprecipitation was performed according to (33) on 3T3-L1 cells using α-PARP-2, α-PPARγ 2 (Alexis, Lausen, Switzerland), and α-MMP9 (Santa Cruz, Santa Cruz, CA) antibodies. We used also a no antibody control. The chromatin fragments collected upon precipitation with the above antibodies were amplified using promoter-specific primers by qPCR. For the analysis of the coding sequence the same qPCR primer set was used as the one for the quantitation of the given gene. The respective primers are listed in Table 2 and Table 3. The results were normalized for the signal of the input and were expressed as a percentage of the aP2 signal with the PARP-2 antibody.
For the testing of the K19 primer set we used non-confluent 3T3-L1 cells transfected with pCMX-ERβ . Chromatin immunoprecipitation was performed using the α-ERβ, (Santa Cruz), as controls we used an α-MRE11 (Santa Cruz) and a no antibody control. The chromatin fragments collected upon precipitation with the above antibodies were amplified using K19 promoter-specific primers by qPCR.

Microscopy
-Formaldehyde-fixed, paraffin-embedded sections (7 µm) were made from WAT samples and were stained with haematoxylin-eosine (HE). The same sections were stained with a biotin-conjugated F4/80 antibody (Serotec, Raleigh, NC, USA, 1:100 dilution) and the bound primary antibodies were detected using streptavidin-perxidase (Vector ABC kit) and diamino-benzidine as chromogenic substrate. Terminally differentiated MEFs were stained by Oil-Red O as described elsewhere. Triglyceride measurement -Triglyceride content of the MEFs were determined using commercially available Sigma kit according to the manufacturer's instructions. Statistical analysis -Significance was analyzed by Student`s t-test. Error bars represent +/-SEM, unless noted otherwise.

RESULTS
In vivo dysfunction of the PPARγ/RXR heterodimer in the white adipose tissue of PARP-2 -/mice. The different fat-depots (epididymal-, mesenteric-and inguinaland the interscapular brown adipose tissue associated WAT were measured in 7 month old PARP-2 -/mice and their wild type littermates. A proportional loss of the weight of all adipose tissue depots was observed in the PARP-2 -/mice (Fig. 1A).
Histological examination of the PARP-2 -/epididymal WAT showed adipocytes with reduced and irregular size. This tissue contained diluted capillaries, indicative of inflammation, which was confirmed by a faint staining with the macrophage-specific F4/80 antibody in the PARP-2 -/- (Fig. 1B, 1C) and the macroscopic appearance of the WAT (Fig.1A). The F4/80 positive cells were present in the vicinity of the blood vessels.
To identify the molecular changes that contribute to the decreased fat accumulation and abnormal adipocyte morphology we determined the expression of the PPARγ target genes, TNFα and hormone sensitive lipase (HSL) by RT-qPCR in the epididymal WAT.
TNFα expression was undetectable in 8 of the 22 mice used for this study (4 out of 14 PARP-2 +/+ and 4 out of 8 PARP-2 -/-). In the TNFα positive mice, expression levels were not different, ruling out a major role for inflammation in the adipose tissue dysfunction in PARP-2 -/mice. The expression level of HSL, which is responsible for lipolysis, was also not different between the 2 genotypes. The expression of several PPARγ target genes, however, was markedly decreased. These include genes involved in chylomicron and VLDL triglyceride hydrolysis (lipoprotein lipase; LPL), FFA uptake (CD36), de novo fatty acid synthesis (fatty acid synthase; FAS) and endocrine signaling (leptin, adiponectin) (Fig. 1D). Interestingly, no difference was detected in PPARγ 1 and PPARγ 2 mRNA levels between the different genotypes.
MEF differentiation is affected by PARP-2 ablation. We next aimed to determine whether MEFs differentiation towards adipocytes was affected by the PARP-2 deletion. Differentiation of PAPR-2 -/-MEFs into adipocytes was decreased as judged by Oil-Red O staining, determination of lipid content and Nile-red staining followed by FACS analysis (Fig. 2A).
The expression of genes involved in adipocyte differentiation and function such as PPARγ 1 and PPARγ 2 were decreased in the PARP-2 -/-MEFs (34). Since the PPARγ transcripts are primarily present in the differentiated cells, these data confirm that PARP-2 -/cells differentiate less into adipocytes. The expression of PPARγ target genes, such as LPL, FAS, leptin, adiponectin and adipocyte fatty acidbinding protein 2 (aP2), were decreased in parallel (Fig. 2B). PARP-2 expression modulates transactivation of PPARs . To measure whether changes in PARP-2 expression affect PPAR transactivation, we used HEK 293 cells transfected with a PPARγ 2 expression vector and a PPARγ responsive luciferase construct. In these experiments we modulated the expression of PARP-2 expression by overexpression and siRNA depletion. For the siRNA depletion of PARP-2 we used the pSuper-siPARP-2 construct, whereas for PARP-2 overexpression we used the pBabe-PARP-2. The pSuper-scrPARP-2 and the empty pBabe vector served as the respective controls. PARP-2 levels were assessed by Western blotting using a PARP-2 specific antibody. For both constructs, the cells were transfected twice, on day 0 and on day 2. On day 3, the specific siRNA decreased PARP-2 protein levels significantly, whereas the scrambled PARP-2 siRNA did not alter the PARP-2 levels. A strong increase in PARP-2 protein was observed on day 3 of the overexpression experiment (Fig 3).
PARP-2 depletion diminished the basal PPARγ activity and abrogated receptor activation by its synthetic ligand, troglitazone.
Conversely, PARP-2 overexpression induced by 3-fold the basal PPARγ activity, although it does not significantly change the ligand-dependent activation by TZD (Fig. 4A). To verify whether this effect of PARP-2 was specific for PPARγ, we performed similar experiments for the related nuclear receptors PPARα (NR1C1) and PPARβ (NR1C2), and the unrelated estrogen receptor β (ERβ, NR3A2). Interestingly, siRNA depletion of PARP-2 increased the basal activity of both PPARα and β ( Figure 4B, C). PARP-2 overexpression did not affect PPARβ, but increased PPARα activity. The activation of PPARα and β with FF and MEHP, respectively, was not modified by the modulation of PARP-2 expression. In addition, neither PARP-2 depletion, nor PARP-2 overexpression had an effect on the basal or ligand-induced activity of ERβ (Fig.  4D). Combined these results indicate specificity of the PARP-2-dependent effect on PPARγ. PARP-2 is the member of the RXR-PPARγ transcription complex. To demonstrate an interaction between PPARγ and PARP-2 we used ChIP assays. To precipitate chromatin from undifferentiated 3T3-L1 cells we used antibodies against PARP-2 and PPARγ 2 . An anti-matrix metalloproteinase-9 (MMP-9) antibody and a no antibody sample served as negative controls. We used qPCR to amplify the promoters of the aP2 (6)  ERβ-regulated promoter (36). PARP-2 and PPARγ gave a strong signal on PPARγregulated promoters. These signals were significantly higher compared to the signal from the K-19 promoter (Fig. 5A). We also performed qPCR reactions to cover the coding sequences of aP2 using the chromatin fragments obtained in the ChIP experiments. The signal of PARP-2 and PPARγ coding sequences in the immunoprecipitates was strongly decreased compared to the signal of the corresponding promoter. Apparently, both PARP-2 and PPARγ are present on the PPARγ-driven promoters, but not in the coding sequence (Fig. 5B). In addition, our results suggest that PARP-2 possesses specificity towards the PPARγ-driven promoters since the signal from ERβdriven K-19 promoter was significantly lower than that from PPARγ-driven promoters.
Despite of the huge difference in the signal of the specific promoters and the non-specific regions (K19 promoter, coding sequence) we observed some background signal from the non-specific region. It is likely that this represents the real presence of PARP-2 in these regions, which is probably linked to the formaldehyde-induced DNA damage.
To provide proof that the interaction of ERβ with the K19 promoter is basically detectable we complemented 3T3-L1 cells with ERβ and we performed ChIP probing with the K19 primer set. To precipitate chromatin from ERβ-complemented 3T3-L1 cells we used an antibody against ERβ, an anti-MRE11 antibody and a no antibody sample served as negative controls. The precipitate of the ERβspecific antibody gave significantly higher signal then the non-specific MRE11 (2.7 fold increase) as well as with the no antibody control (6.1 fold increase) proving that the K19 primer pair is capable of detecting the K19 promoter if present in the precipitate (Fig. 5C).

DISCUSSION
PPARγ plays an important role in adipose tissue differentiation and function. In PARP-2 knockout mice we have identified a defect of adipose tissue function and a decrease of adipocyte differentiation. In vivo, the adipose tissue depots had smaller weight and histologically showed an adipodegenerative phenotype.
We have detected a mild inflammation in the WAT of the PARP-2 -/mice. The capillaries were dilated and we have detected F4/80 positive cells in the vicinity of the capillaries suggesting the presence of macrophages. The areas more distant from the capillaries are devoid of staining. Similar coloration was not observed in the WAT of the wild type mice. Activated macrophages and adipocytes may secrete proinflammatroy cytokines, such as TNFα that may induce adipocyte cell death (37). Since TNFα expression was not detectable in many mice and if it was detected, its expression was not significantly increased by the absence of PARP-2, it is less likely that inflammation is a leading cause of the adipodegeneration in the PARP-2 -/mice. It is also unlikely that increased lipolysis may contribute to the phenotype in the PARP-2 -/mice, since there was no difference in the expression of HSL between the wild-type and PARP-2 -/mice.
We did observe decreased expression of multiple PPARγ target genes involved in adipocyte function. Expression of both PPARγ isoforms was normal, suggesting effects on PPARγ/RXR transactivation. In vitro, the differentiation of the PARP-2 -/-MEFs into adipocytes was delayed when compared with the differentiation of wild-type MEFs. At the end of the differentiation the expression of both PPARγ 1 and PPARγ 2 was decreased in the PARP-2 -/cells indicating the lack of differentiation. Similarly, the expression of the PPARγ target genes was decreased.
In transfection assays, the ablation of PARP-2 results in the diminution, whereas PARP-2 overexpression raises transactivation by PPARγ. The effect of PARP-2 seems specific for PPARγ, since opposite or no effects were observed for the related PPARα and PPARβ, and the non related ERβ.
PARP-2 achieves these activities because it is part of the PPARγ/RXR transcription complex as shown by ChIP assays, suggesting that PARP-2 could act as a PPARγ/RXR receptor cofactor.
Both members of the PPARγ -RXR nuclear receptor dimer might be the effector behind the phenotype of the PARP-2 -/mice. If PARP-2 would influence directly RXR all PPAR isoforms should respond the same way to the modulation of PARP-2 expression. PPARγ was differentially regulated when compared with PPARα and β, suggesting that PARP-2 acts on PPARγ.
Our report comprises in vitro data and gives first time in vivo evidence that PARP-2 may be considered a cofactor of nuclear receptor transcription.
PARP-1 has been described as a cofactor for numerous transcription factors (reviewed in (40) and (25)), including for some members of the nuclear receptor family, such as the progesterone receptor (41), RXR (42); (43) androgen receptor (44) and the thyroid receptor (42). A recent study, based on in vitro results, suggested that PARP-2 acts as a cofactor of a homeodomain-containing transcription factor, TTF1 that belongs to the Nkx-2 family. Binding of PARP-2 through its E-domain to the C-terminus of TTF1 regulates the expression of the surfactant protein-B in lung epithelial cells.
TTF1-mediated transcription encompasses similar mechanisms including chromatin modification, and involves some of the same cofactors such as the SRCs as described for PPARγcoupled transcription. This suggests that similar molecular mechanisms exist both in the case of PPARγ-and TTF1-mediated transcription (25). Our results hence confirm the observation of Maeda and colleagues, that PARP-2 is a cofactor of some transcription factors, and extend these conclusions by showing that PARP-2 is involved in nuclear receptor-mediated transcriptional control in vivo. Recent evidence has suggested that the interaction between PARP-1 and the promoter of target gene could be mediated via double strand breaks, which are produced by activation of a nuclear receptor followed by the unwinding of DNA by topoisomerase II (45). Our results do provide evidence that interaction with DNA is important for the interaction between PPARγ and PARP-2. ChIP assays that depend on DNA binding show strong interaction.
In contrast, immunoprecipitation experiments performed on cell extracts showed only a weak interaction between PARP-2 and PPARγ (data not shown), which was abrogated by low concentrations of NP40 (>0.1%). Furthermore, like is the case for PARP-1, the NH 2 -terminus of PARP-2, comprising its DNA-binding domain, seems also to play the most important role in the interaction with PPARγ. Consistent with this observation a nuclear receptorbinding consensus sequence ( 113 LIQLL 117 ) was present in the E domain of PARP-2.
Concerning the mode of action of PARP-2, it is possible that not only the physical presence but also the activity of PARP-2 is necessary for the nuclear receptor function. Poly(ADP-ribosyl)ation is reported to increase throughout the differentiation process of 3T3-L1 cells (46). Interestingly this poly(ADPribosyl)ation activity is not completely inhibited by PARP-1 depletion (47), suggesting the involvement of other member(s) of the PARP family, such as PARP-2. Indeed both PARP-1 and PARP-2 are reported to poly(ADP-ribosyl)ate histones (13). Similarly to histone acetylation, poly(ADP-ribosyl)ation of the high mobility group of proteins and histones, loosens chromatin structure enabling transcription initiation (48). Consistent with this line of thinking, there is molecular and in vivo evidence that the enzymatic activity of PARP-1 is necessary for efficient gene transcription and inhibition of PARP activity impairs the transcription of a number of different genes, including different chemokines and inflammation-related genes (e.g. iNOS, TNFα, ICAM-1, IL-8, MIP-1α, IL−12) (49), (40), (50).
PARP-2 specifically occupies the promoter of PPARγ target genes, since it bound efficiently to the regulatory sequence, whereas binding to the corresponding coding sequences was strongly decreased. Despite this rather specific binding, we observed a background signal rising most likely from non-coding regions or from non-PPARγdependent promoters, such as that of the K19 gene, which is under the control of ERβ. When comparing the specific to the above mentioned non-specific signal it is at least 10-100 fold increased, which can be considered as a significant difference. It is likely that the non-specific presence of PARP-2 on the K19 promoter and in the non-coding regions is explained by the fact that PARP-2 binds to the DNA-damage sites created by the formaldehyde treatment during the crosslinking of the cells. The crosslinking-related DNA damage is present throughout the entire genome, equally affecting coding regions and promoters, thus theoretically providing a background signal throughout the genome.
The present study indicates that PARP-2 modulates the activity of PPARγ/RXR nuclear receptor complex, a key transcription factor involved in the pathogenesis of several important diseases such as obesity, insulin resistance, type II diabetes atherosclerosis and lipodystrophy. Since many of these diseases affect a large part of the population and have high costs to society, our data linking the activation of PPARγ and PARP-2, open up the possibility to modulate PPARγ activity via PARP-2. It is therefore tempting to speculate that the various PARP inhibitors that are currently being developed and being tested in clinical trials (51) could also be useful in the metabolic disease arena. Cells were scraped from day 2 daily. These samples were analyzed by Western blotting. PARP-2 was depleted by the pSuper-siPARP-2 construct, but was unmodified by the pSuper-scrPARP-2 construct. Whereas the transfection with pBabe-PARP-2 resulted on day 3 and day 4 in a robust induction of PARP-2 expression, the transfection with pBabe alone did not modify PARP-2 expression. . PPARγ 2 and PARP-2 occupy together the PPARγ-dependent promoters. A, PARP-2 is present on PPARγ driven promoters (aP2, CD36) as demonstrated in ChIP assays. The PARP-2 signal is increased in the PPARγ-driven promoters compared to the non-related, ERβ-driven K-19 promoter. Error bars represent SD. ###, *** p<0.001; #, * p<0.05. B, Similar ChIP assay was performed to compare the presence of PARP-2 on the promoters versus the coding sequence of the same gene. PARP-2 is present preferentially on the promoter rather then the coding sequence of the aP2 gene. Error bars represent SD. *** p<0.001; * p<0.05. C, As a positive control, the K19 primer set was tested for the ability to detect the presence of ERβ in ERβ transfected 3T3-L1 cells. The α-ERβ signal was significantly higher then the signal of the negative control α-MRE11 and the no antibody control. These data confirm that the K19 primer set is capable of the detection of the presence of the ERβ receptor on the K19 promoter. Error bars represent SD. ###, *** p<0.001; #, * p<0.05. Table 1. Oligonucleotides used to generate pSuper-siPARP-2 and pSuper-scrPARP-2 The interfering sequences are in bold.