Modulation of Poly(ADP-Ribose) Polymerase-1 (PARP-1)-Mediated Oxidative Cell Injury by Ring Finger Protein 146 (RNF146) in Cardiac Myocytes
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Poly(ADP-ribose) polymerase-1 (PARP-1) activation is a hallmark of oxidative stress-induced cellular injury that can lead to energetic failure and necrotic cell death via depleting the cellular nicotinamide adenine dinucleotide (NAD+) and ATP pools. Pharmacological PARP-1 inhibition or genetic PARP-1 deficiency exert protective effects in multiple models of cardiomyocyte injury. However, the connection between nuclear PARP-1 activation and depletion of the cytoplasmic and mitochondrial energy pools is poorly understood. By using cultured rat cardiomyocytes, here we report that ring finger protein 146 (RNF146), a cytoplasmic E3-ubiquitin ligase, acts as a direct interactor of PARP-1. Overexpression of RNF146 exerts protection against oxidant-induced cell death, whereas PARP-1-mediated cellular injury is augmented after RNF146 silencing. RNF146 translocates to the nucleus upon PARP-1 activation, triggering the exit of PARP-1 from the nucleus, followed by rapid degradation of both proteins. PARP-1 and RNF146 degradation occurs in the early phase of myocardial ischemia-reperfusion injury; it precedes the induction of heat shock protein expression. Taken together, PARP-1 release from the nucleus and its rapid degradation represent newly identified steps of the necrotic cell death program induced by oxidative stress. These steps are controlled by the ubiquitin-proteasome pathway protein RNF146. The current results shed new light on the mechanism of necrotic cell death. RNF146 may represent a distinct target for experimental therapeutic intervention of oxidant-mediated cardiac injury.
Poly(ADP-ribose) polymerase-1 (PARP-1) is a ubiquitously expressed enzyme that catalyzes the poly(ADP-ribosyl)ation of acceptor proteins by using nicotinamide adenine dinucleotide (NAD+) as a substrate. The protein consists of an N-terminal DNA-binding domain, an automodification domain and a C-terminal catalytic domain. PARP-1 has low basal enzymatic activity, but its catalytic activity is dramatically stimulated on binding to damaged DNA (single or double strand breaks). Targets of the enzyme include histone proteins and transcription-related factors and PARP-1 itself (via its automodification domain). PARylation can affect the target protein function and its interactions with various proteins and DNA; thereby, PARP-1 plays a key role in the regulation of DNA repair and gene transcription (1,2).
Traditionally, the regulation of nuclear DNA repair and maintenance of genomic integrity was considered the main physiological function of PARP-1. The functional roles of PARP-1 were later extended by the discovery that PARP-1 acts as a coactivator and corepressor of gene transcription, thereby regulating the production of inflammatory mediators (1,2). In response to massive amount of DNA damage, PARP-1 can become so robustly activated that it can lead to a marked depletion of the cellular pool of its substrate (NAD+), culminating in a catastrophic cellular energetic deficit (1,2). Overactivation of PARP-1 has been implicated in a variety of pathophysiological conditions, including ischemia-reperfusion injury, critical illness, pancreatic β-cell injury, diabetic complications and neurodegeneration (1,2). It also plays a role in the pathogenesis of myocardial ischemia reperfusion, where PARP-1 genetic deficiency and pharmacological PARP inhibition exert cardioprotective effects (1, 2, 3, 4, 5, 6).
Energetic failure following PARP-1 activation is not only a result of direct NAD+ consumption, but it is also triggered by mitochondrial dysfunction induced by negatively charged poly(ADP-ribose) (PAR) polymers, which are the principal products of PARP-1 and can be subsequently “liberated” from the PARy-lated proteins by various enzymes including PAR glycohydrolase (7,8). In the early phase of oxidative injury, enzymatic NAD+ consumption appears to be more important, and cell death mostly occurs via necrosis. However, in the late phase of the injury, diminished mitochondrial output and release of proapoptotic molecules from the mitochondria play a dominant role, leading to various forms of programmed cell death (including apoptosis and parthanathos).
Recent work, using differential display to identify genes induced in the late phase of oxidant injury, led to the discovery of the PAR-interacting protein RNF146. Transgenic RNF146 exerted protection against N-methyl-d-aspartate (NMDA)-induced neural cell death by directly binding to the PAR polymer (9,10). Currently, all available information on the role of RNF146 in the modulation of cell death relates to neuronal injury, although the protein is expressed at high levels in most peripheral tissues. Because our pilot studies showed that RNF146 expression is the highest in the heart and muscle, the goal of the current project was to characterize the role of RNF146 in PARP-1-mediated cell death during oxidative myocyte injury in vitro and during myocardial ischemia-reperfusion injury in vivo.
Materials and Methods
H9c2 rat cardiomyocytes were purchased from American Type Culture Collection (ATCC, Manassas, VA, USA) and maintained in Dulbecco’s modified Eagle medium (DMEM) (Biochrom, Berlin, Germany) supplemented with 4 mmol/L glutamine, 10% fetal bovine serum (FBS) (PAA Laboratories, Westborough, MA, USA), 100 IU/mL penicillin and 100 µg/mL streptomycin (Invitrogen/Life Technologies, Carlsbad, CA, USA) at 37°C in 10% CO2 atmosphere.
H9c2 cardiomyoblasts (10),000/well) were plated on 96-well plates; the following day, the cells were transfected with RNF146 siRNA (1 pmol/well; Silencer Select; assay ID: s158554; Life Technologies) by using Lipofectamine 2000 transfection reagent. Control cells were transfected with Silencer Select negative control #1 siRNA (ID: 4390844; Life Technologies). The knockdown efficiency was evaluated by real-time polymerase chain reaction (PCR) (Taqman assay ID: Rn02534308 using TaqMan Rodent GAPDH Control Reagents [catalog no. 4308313] normalization; Applied Biosystems/Life Technologies) and by Western blotting 48 h after transfection. The cells were exposed to oxidant injury 48 h after transfection.
The complete rat RNF146 cDNA (IMAGE: 7135728; NCBI accession number BC083675) was obtained in pEXPRESS-1 vector from Life Technologies. The coding sequence was excised with EcoRI/XhoI digestion and sub-cloned into pcDNA3.1(+) (Life Technologies) to create the vector pcDNA-RNF146. H9c2 cells were transfected with pcDNA-RNF146 and selected with G418 (500 µg/mL; Invitrogen/Life Technologies) for 4 wks. Individual colonies were picked up and tested for RNF146 overexpression by Western blotting. RNF146 overexpressing clones were expanded, and the presence of the integrated expression cassette was confirmed by PCR. Control clones were generated by transfection of H9c2 cells with β-galactosidase expression vector [pcDNA3.1(+)/myc-His/LacZ; Invitrogen/Life Technologies] and 4-wk-long selection with G418.
Confirmation of Stable Transfection by PCRs
H9c2 cells were transfected with pcDNA-RNF146 and selected with G418 (500 µg/mL, Invitrogen) for 4 wks. Individual colonies were picked up and tested for RNF146 overexpression by Western blotting. RNF146 overexpressing clones were expanded in T75 flasks and used for DNA isolation as described (11). Cells transfected with pcDNA3.1(+)/myc-His/LacZ were selected with G418 and used as controls. Cells were pelleted by centrifuging at 200g for 10 min and lysed in 450 µL DNA lysis buffer (100 mmol/L Tris, pH 8.0, 20 mmol/L ethylenediaminetetraacetic acid [EDTA], 0.8% N-lauroylsarcosine). A total of 175 units RNase A (25 µL; 5PRIME, Gaithersburg, MD, USA) was added, and the samples were incubated at 37°C for 1 h; then 60 mAU Proteinase K (100 µL; 5PRIME) was added and incubated at 55°C overnight. DNA was isolated by subsequent phenol-chloroform extraction and ethanol precipitation. A total of 20 ng DNA was used as template in PCRs by using F1 (5′-CGTGTACGGTGGGAGGTCTA-3′) and R1 (5′-CAGGTCTCACTCGCCTTCTT-3′), F2 (5′-CGTGTACGGTGGGAGGTCTA-3′) and R2 (5′-ATGAAGCGCCCTTTACACAC-3′) or F3 (5′-TAGTGTGTCCCCGTGCATTA-3′) and R3 (5′-GCGATGCAATTTCCTCATTT-3′) primers by using a touchdown PCR protocol. The DNA quality was checked by using PCR primers to amplify an ∼1.2-kb region of the adiponectin 1 (ADIPOR1) gene (ADIPOR1 forward 5′-CGCATCCACACAGAAACTG-3′, ADIPOR1 reverse 5′-TGAGCATGGTCAAGATTCCC-3′).
RNF146 expression was also measured at the mRNA level. Total RNA was isolated from RNF146 overexpressing cells and pcDNA3.1(+)/myc-His/LacZ transfected controls by TRizol reagent (Invitrogen/Life Technologies). A total of 2 µg RNA was treated with DNase (Epicentre), and reverse transcription was carried out by using a High Capacity cDNA Archive kit (Applied Biosystems/Life Technologies) following the manufacturer’s instructions. RNF146 overexpression was confirmed by RNF146 realtime PCR (Taqman assay ID: Rn02534308 using TaqMan Rodent GAPDH Control Reagents [catalog no. 4308313] normalization) and by an exon-spanning assay (RNF146 forward primer: 5′-GTGCCTGTGGGATCTGTGAT-3′, RNF146 reverse primer: 5′-CAGGTCTCACTCGCCTTCTT-3′ and FAM/TAMRA labeled RNF146 probe: 5′-GGCTGTGGTGAAATTGATCACTCAC-3′).
Transient Transfection of 293T Cells with the RNF146 Expression Vector
The 293T cells were purchased from American Type Culture Collection (ATCC) and maintained in DMEM supplemented with 2 mmol/L glutamine, 10% FBS (Invitrogen/Life Technologies), 100 IU/mL penicillin and 100 µg/mL streptomycin (Invitrogen/Life Technologies) at 37°C in 10% CO2 atmosphere. Cells were transiently transfected with pcDNA-RNF146 by a linear polyethyleneimine-based transfection reagent (ExGen500, Fermentas, Vilnius, Lithuania).
Cells were exposed to H2O2 to induce PARP-1 activation and cell death, and viability was measured by the 3-(4),5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) assay after 3 h. In addition, PARP-1 activation was confirmed by Western blotting (as described below) by using an antibody against PAR at 5 min after H2O2.
Oxidative Stress-Induced Injury and Viability Assays
H9c2 cells (10),000/well) were plated on 96-well plates, and 48 h later, they were exposed to H2O2 in fresh culture medium for 3 h. Cell viability was measured by the MTT and LDH assays as described (12). (Cells exposed to siRNAs were subjected to H2O2 injury 48 h after transfection.) The PARP inhibitor PJ34 (3 µmol/L; Sigma-Aldrich, St. Louis, MO, USA) was added to the cells simultaneously with H2O2. Cell culture supernatant (30 µL) was saved for assaying the lactate dehydrogenase (LDH) release, and MTT was added to the cells at a final concentration of 0.5 mg/mL for 1 h. The converted formazan dye was dissolved in isopropanol and measured photometrically on a Synergy Mx plate reader (BioTek, Winooski, VT, USA). Viability was calculated by using a calibration curve created by measuring the MTT converting capacity of serial dilutions of H9c2 cells.
LDH activity was measured by mixing the supernatant with freshly prepared LDH assay reagent [85 mmol/L lactic acid, 1,040 mmol/L NAD+, 224 mmol/L N-methylphenazonium methyl sulfate, 528 mmol/L 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyl-2H-tetrazolium chloride, 200 mmol/L Tris, pH 8.2]. The changes in absorbance at 492 nm were measured kinetically for 15 min, and LDH activity in the supernatant is shown as the Vmax value.
Oxygen-Glucose Deprivation Injury
Oxygen-glucose deprivation (OGD) injury was conducted as previously described (13,14). RNF146 overexpressing and control H9c2 cells (10),000/well) were plated on 96-well plates and cultured for 4 d. Culture medium was replaced with DMEM containing no glucose before the induction of hypoxia. Culture plates were placed in gas-tight incubation chambers (Billups-Rothenberg, Del Mar, CA, USA), and the chamber atmosphere was replaced by flushing the chamber with 95% N2/5% CO2 mixture at 30 L/min flow rate for 10 min. The hypoxia was maintained by clamping and incubating the chambers for 8 h at 37°C. All assay plates were subjected to hypoxia-included wells exposed to glucose-free medium (OGD) and medium containing 4.5 g/L glucose (hypoxia CTL). Cells were also exposed to glucose-free medium or maintained in glucose-containing medium (4.5 g/L glucose) under normoxia (normoxia CTL). After 8 h, glucose and serum concentration was restored by supplementing the culture medium with glucose and FBS, and the cells were incubated for 16 h at 37°C at 5% CO2 atmosphere. MTT converting capacity (“viability”) of the cells, ATP content and LDH release was measured after 8 h of hypoxia/normoxia and after the 16-h recovery period.
ATP concentration was determined by the commercially available CellTiter-Glo® Luminescent Cell Viability Assay (Promega, Madison, WI, USA). The cells were lysed in 100 µL CellTiter-Glo reagent according to the manufacturer’s recommendations, and the luminescent signal was recorded for 1 s on a high-sensitivity luminometer (Synergy 2; Biotek, Winooski, VT, USA). The assay is based on ATP requiring luciferin-oxyluciferin conversion mediated by a thermostable luciferase that generates a stable “glow-type” luminescent signal.
Cells were lysed in denaturing loading buffer (20 mmol/L Tris, 2% SDS, 10% glycerol, 6 mol/L urea, 100 µg/mL bromophenol blue, 200 mmol/L β-mercaptoethanol), and mouse tissue samples were homogenized in radioimmunoprecipitation assay buffer (150 mmol/L NaCl, 1% NP40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate, 50 mmol/L Tris, pH 8.0) supplemented with protease inhibitors (Complete Mini EDTA-free; Roche, Indianapolis, IN, USA). Lysates were sonicated, boiled and resolved on 4–12% NuPage Bis-Tris acrylamide gels (Invitrogen/Life Technologies) and then transferred to nitrocellulose. Membranes were blocked in 10% nonfat dried milk and probed overnight with anti-RNF146 (Abnova, Walnut, CA, USA), PARP-1 (Cell Signaling Technology, Danvers, MA, USA), PAR (catalog number 528815, lot number D00057484; Calbiochem, San Diego, CA, USA), ubiquitin (Cell Signaling Technology) or heat shock protein 70 (HSP70, Enzo Life Sciences, Farmingdale, NY, USA) antibodies (1:1,000). After incubation with peroxidase-conjugated secondary antibodies, the blots were detected on a charge-coupled device (CCD) camera-based detection system (GBox; Syngene USA, Frederick, MD, USA) with enhanced chemiluminescent substrate. To normalize signals, membranes were stripped in 62.5 mmol/L Tris, 2% SDS, 100 mmol/L β-mercaptoethanol at 60°C for 20 min, blocked and reprobed with antibodies against actin (Santa Cruz Biotechnology, Santa Cruz, CA, USA). The signals were quantitated by using the Genetools analysis software (Syngene USA).
PARP-1 was activated in confluent H9c2 cells grown in 6-cm culture dishes by exposure to H2O2 in fresh culture medium. Control cells were pretreated with PARP inhibitor PJ34 (3 µmol/L, 30 min) and received fresh culture medium without H2O2. Cells were lysed and scraped in lysis buffer (100 mmol/L HEPES, 200 mmol/L NaCl, 40 mmol/L EDTA, 4 mmol/L EGTA, 100 mmol/L NaF, 20 mmol/L β-glycerophosphate, 2 mmol/L Na3VO4, 2% Triton X-100) supplemented with protease inhibitors (Complete Mini EDTA-free) on ice. Cell lysates were centrifuged at 16,000 × g for 10 min, and the cleared lysate was used for direct immunoprecipitation by using the Dynabeads Protein G Immunoprecipitation Kit (Life Technologies). Antibodies recognizing the N terminus of RNF146 (Abnova), residues surrounding Gly623 of PARP-1 (Cell Signaling Technology) or ubiquitin (Cell Signaling Technology) were incubated with Protein G Dynabeads for 1 h at 4°C to allow them to bind to magnetic beads. The antibody-Dynabeads complexes were incubated with 2× diluted cell lysates for 1 h at 4°C. The captured complexes were thoroughly washed and eluted under denaturing conditions. Samples were analyzed by Western blotting by using antibodies recognizing different parts of the proteins: the C terminus of RNF146 (Aviva Systems Biology, San Diego, CA, USA) and the region surrounding Gly215 of PARP-1 (Cell Signaling Technology) and protein-A-HRP conjugate (Amersham/GE Healthcare, Life-Sciences, Pittsburgh, PA, USA).
H9c2 cells were plated (50),000/well) on Lab-Tek™ II eight-well chamber slides (Nalge Nunc, Rochester, NY, USA) and cultured until they reached confluency. They were exposed to H2O2 (1 mmol/L, 30 min) or vehicle and fixed in 4% neutral buffered formalin for 15 min. Cells were permeabilized with 0.2% Triton X-100, blocked with 2.5% horse serum (Vector Laboratories, Burlingame, CA, USA) and probed with PARP-1 (1:100; Cell Signaling Technology) or RNF-146 antibody (1:100, Abnova, Walnut, CA) followed by incubation with Alexa Fluor 546-labeled secondary antibody (Invitrogen/Life Technologies). The cells were subsequently stained with nuclear stain Hoechst 33342 and Alexa Fluor 488 phalloidin conjugate, and after extensive washing, coverslips were applied with Mowiol. (Alexa Fluor 488 secondary label was used to detect RNF146 when no phallodin staining was performed.) Images were taken on an Eclipse 80i fluorescent microscope (Nikon Instruments, Melville, NY, USA) with a Coolsnap HQ2 14-bit CCD camera (Photometrics, Tucson, AZ, USA). PARP-1 staining intensity was evaluated by a NIS Elements software package (Nikon Instruments). The nuclear area was recognized on a Hoechst 33342 channel, and applying this binary layer to the PARP-1 channel fluorescence intensity was measured in the nuclear and extranuclear areas. The ratio of nuclear and extranuclear areas remained unchanged after H2O2 exposure on the images evaluated.
Balb/c male mice (n = 10) were anesthetized and myocardial ischemia was produced by occlusion of the left anterior descending (LAD) coronary artery as described (15). After 30 min of LAD coronary artery occlusion, the ligature was removed, and reperfusion was visually confirmed. The hearts were reperfused for 2 h and were processed for histology in half of the animals. The ischemic area (area at risk) was separated from the nonischemic parts and processed separately. Mouse heart samples were fixed in 4% neutral buffered formalin and embedded in paraffin. The 4-µm sections were cut and the slides were stained with hematoxylin and eosin, with Masson trichrome stain technique or subjected to immunohisto-chemistry for PARP-1, PAR and RNF146. In the remaining animals, reperfusion was allowed for 3 h and the heart samples were used for quantitative analysis by Western blotting. Sham-operated control mice (n = 10) were subjected to anesthesia and surgical procedures except for LAD coronary artery occlusion and samples were processed simultaneously.
Immunohistochemical staining was performed as previously described (16,17). Mouse heart samples were fixed in 4% neutral buffered formalin and embedded in paraffin. The 4-µm sections were cut and picked up on adhesive slides. Endogenous peroxidase activity was suppressed on deparaffinized and rehydrated sections by treating slides with H2O2 (0.6% in methanol, 15 min). Antigenic epitopes were retrieved by microwaving the sections in citrate buffer (0.2 mol/L citrate, pH 3.0, for PAR or 10 mmol/L citrate, pH 6.0, for all other antigens). Sections were blocked with normal horse serum (blocking serum) or with M.O.M. diluent (Vector Laboratories). Ig and protein blocking reagents (Vector Laboratories) in PBS contained 0.2% Triton X-100. Antibodies against PARP-1 (1:100; Bethyl Laboratories, Montgomery, TX, USA), PAR (1:200; Calbiochem) or RNF146 (1:100; Abnova) were applied in blocking serum or M.O.M. diluent, and sections were incubated overnight at 4°C. Sections were washed in PBS containing 0.2% Triton X-100 (wash buffer) and incubated with biotinylated anti-rabbit or anti-mouse antibodies (1:200), respectively. Then sections were washed and incubated with ABC peroxidase reagent (Vectastain Elite ABC kit; Vector Laboratories) for 30 min, and peroxidase sites were revealed with 3,3′-diaminobenzidine-tetrahydrochloride (DAB) and H2O2 (DAB substrate kit; Vector Laboratories). Slides were washed and counterstained with Hematoxylin QS (Vector Laboratories), dehydrated in an ascending alcohol series, cleared in xylene and coverslipped with Permount (Thermo Fisher Scientific Inc., Waltham, MA, USA). Negative control sections were simultaneously stained with the omission of primary antibodies.
Slides were viewed on an Eclipse 80i fluorescent microscope (Nikon Instruments) under bright-field illumination, and images were captured with a digital firewire camera system (DS-Vi1 Color digital camera; Nikon Instruments). Five images (area of 251 × 188 µm) were taken of all slides, and the positively stained and negative nuclei were counted. The percent positivity of nuclei is shown on the graphs.
One-way analysis of variance (ANOVA) was used to detect differences between groups or unpaired two-tailed Student t test to compare two groups. Post hoc comparisons were made using the Tukey test. A value of p < 0.05 was considered statistically significant. The correlation analysis was performed by linear regression analysis. All statistical calculations were performed by using Prism 4 analysis software (GraphPad Software, La Jolla, CA, USA). Data are shown as mean ± SEM (standard error of the mean) values.
RNF146 Protects Cardiomyocytes Against Oxidant-Induced Cell Death
RNF146 Directly Interacts with PARP-1, and the Levels of Both Proteins Decrease in Oxidative Stress
As mentioned in the Introduction, RNF146 was initially recognized as a cytoplasmic PAR binding protein that can capture PAR polymers and protects neurons against the PAR polymer-induced cell death (parthanathos). In our cardiomyocyte model, however, RNF146 had a protective effect in the early (necrotic) phase of oxidant injury without blocking the activation of PARP-1. Because the intracellular NAD+ pools are compartmentalized (9,23,24), the mechanisms by which PARP-1 can exhaust the cytoplasmic NAD+ compartment remain incompletely understood. Still, a significant degree of cellular NAD+ and ATP depletion occurs after PARP-1 activation in oxidant injury, and depletion of cytosolic NAD+ is required for cell death, since restoration of cytosolic NAD+ content is sufficient to prevent PARP-1-mediated cell death (25). We hypothesized that RNF146 may uncouple PARP-1 activation from the loss of NAD+ in the cytoplasm by interfering with a PARylated “signaling molecule.” Because PARP-1 is the most abundantly PARylated protein in oxidative stress, we tested whether RNF146 can interact with PARP-1. Under basal conditions, PARP-1 is primarily nuclear and, in some cells, mitochondrial (26, 27, 28), whereas RNF146 is cytoplasmic. No interaction is expected between the two proteins. We tested whether the localization of RNF146 may change in oxidative stress, since a nuclear translocation of the overexpressed RNF146 was observed by us in cardiomyocytes (Figure 4C) and by others in neurons (8). We found that endogenous RNF146 also translocates to the nucleus upon H2O2 stimulation (Figure 6A).
To test the potential of a direct interaction between RNF146 and PARP-1, coimmunoprecipitation assays were performed. Immunoprecipitation by PARP-1 antibody pulled down RNF146, whereas immunoprecipitation with RNF146 antibody pulled down PARP-1 in both RNF146 overexpressing and normal control cells (Figure 6B). Interestingly, the interaction between the two proteins did not appear to be enhanced when the auto-PARylation of PARP-1 increased. RNF146 overexpression increased the amount of PARP-1 in the precipitate either using RNF146 or PARP-1 antibody for pull-down.
RNF146 was reported to act as an E3 ubiquitin ligase, which is the protein that confers substrate specificity in the ubiquitin-proteasome system: it targets specific protein substrates for degradation (10,29,30). To test whether PARP-1 is ubiquitinated, we also performed the precipitation with anti-ubiquitin antibody that also pulled down PARP-1 from the RNF146 overexpressing cells and to a smaller extent from control cells (Figure 6C). The amount of detectable PARP-1 was lower in the samples exposed to H2O2 (1 mmol/L, 30 min), and free ubiquitin completely disappeared from them.
RNF146 Undergoes Nuclear Translocation in Myocardial Ischemia-Reperfusion Injury In Vivo and PARP-1 Is Consumed
The main findings and conclusions of the present study are the following: (a) RNF146, a cytoplasmic E3-ubiquitin ligase, acts as a direct interactor of PARP-1 in cardiac myocytes in vitro. (b) RNF146 modulates oxidative cell death; its overexpression protects against cell injury, whereas cell injury is augmented after RNF146 silencing in cardiac myocytes in vitro. (c) RNF146 translocates to the nucleus concomitantly with oxidant-induced PARP-1 activation, resulting in the nuclear-to-cytoplasmatic translocation of PARP-1, followed by the degradation of both proteins in cardiac myocytes in vitro. (d) The same processes also occur in vivo, where PARP-1 and RNF146 degradation occurs in the early phase of myocardial ischemia-reperfusion injury.
The original working model that stipulated that oxidant-mediated DNA injury promotes necrotic type cell death via PARP-1 overactivation and associated metabolic catastrophe due to NAD+ and ATP depletion and mitochondrial dysfunction (31, 32, 33, 34, 35) has been further advanced and refined over the last two decades in several different ways. First, the above-mentioned mechanisms have been investigated in the context of caspasemediated PARP-1 degradation, a well-known phenomenon that occurs in oxidatively stressed and dying cells. It has been demonstrated that caspase activation, and the consequent proteolytic PARP-1 degradation, in fact, serves to protect cells from an overwhelming degree of cell necrosis, and the intracellular energetic pools that are saved in the cell after PARP-1 cleavage are then used to execute apoptosis (36, 37, 38). Apoptosis, in this context, can be viewed as a mode of cell death that is more favorable than necrosis, because the various intracellular cell constituents, many of which serve as damage-associated molecules, are not released into the extracellular environment. A further refinement in the concept of PARP-1-mediated cell death was related to the discovery that a mitochondrial isoform of PARP-1 also exists (at least in some cell types) and may play an active role in oxidative cell injury (26, 27, 28). Another important finding was the recognition that PAR (the product of PARP) can be released into the cytoplasm from the nucleus and can directly act on the mitochondria to induce the release of apoptosis-inducing factor (AIF), which in turn induces nuclear DNA damage (39, 40, 41, 42). Whereas this mechanism was originally recognized in neurons, subsequently, it was demonstrated in oxidatively stressed cardiac myocytes (43,44). The PARP-1 activation/PAR/AIF-mediated cell death, which has elements of both necrosis and apoptosis, is now also considered a distinct mode of cell death (parthanathos) (8,39). A systematic search for downstream regulators of parthanathos identified RNF146 (also termed “Iduna”) (9). In an independent line of studies, RNF146 (in this study, it was also called “dactylidin”) was simultaneously identified as a differentially expressed protein in the vulnerable regions of the brain in Alzheimer’s disease; in this study, its E3 ubiquitin ligase function was also suggested (18).
The initial functional role of RNF146 was first demonstrated in neuronal models of cell death induced by NMDA receptor agonists. RNF146 was found to be neuroprotective against glutamate NMDA receptor-mediated excitotoxicity both in vitro and in vivo and against stroke (9). The major mechanism responsible for the protective effects was attributed to its ability to directly interfere with the PAR polymer (and its downstream effects such as AIF release and cell death), because the protective effects of RNF146 were attenuated by mutation at its PAR polymer-binding site (9). However, subsequent work revealed a more complex role of RNF146 in its interactions with PARP-1. For example, Zhang et al. (30) demonstrated that the interaction of RNF146 with PAR promotes the degradation of PARylated proteins. This effect was attributed to the interaction of PAR with the WWF domain of RNF146; the enhanced degradation of PARylated proteins (as shown with several PARylated proteins, such as axin, tankyrase and PARP-1) was found to occur through ubiquitination and subsequent degradation through the proteasome (10,46). Another line of work demonstrated that the E3 ligase function of RNF146 is increased by PAR binding, and this PAR binding leads to the ubiquitination of the associated proteins (as demonstrated with PARP-1, X-ray repair cross-complementing protein 1 [XRCC1], DNA ligase III and Lupus Ku autoantigen protein p70 [Ku70]). The PAR-dependent, RNF146-mediated ubiquitination of PARP-1 has been demonstrated to target PARP-1 for proteasomal degradation (47,48). It is noteworthy that the ubiquitination of PARP-1 has already been noted in prior studies as well (49), although without exploring the potential role of RNF146 in this process.
Curiously, the subcellular compartmentalization of PARP-1 was not addressed in these prior studies, even though it was generally assumed that PARP-1 is primarily nuclear, and RNF146 is exclusively cytoplasmatic. The previous state-of-the-art of parthanathos specified that PAR (that is, the product of PARP-1), but not PARP-1 enzyme itself, undergoes nuclear-to-cytoplasmatic translocation. The results of the current study (admittedly, in a different cell type and cell injury model, which makes direct comparisons difficult) confirm and extend some of these previous observations and may even resolve some of the above-mentioned dilemmas. First, the studies in the current report confirm the direct interaction of PARP-1 and RNF146. (Although in our experiments the intensity of the pull-down did not increase when the auto-PARylation of PARP-1 increased, this does not necessarily exclude the possibility that this interaction occurs through PAR, since PARP-1 has some degree of basal auto-PARylation. However, other mechanisms [for example, a direct protein-protein interaction] may also be possible). Second, the results of the current study confirm the RNF146-mediated ubiquitination of PARP-1 and are consistent with the model whereby the RNF146-induced ubiquitination directs PARP-1 into the proteasome, thereby facilitating its degradation. Third, on the basis of novel results of the current study, we conclude that the interaction of RNF146 and PARP-1, at least in oxidatively stressed cardiomyocytes, involves the cytoplasmatic-to-nuclear translocation of RNF146, as well as the nuclear-to-cytoplasmatic exit of PARP-1. The present report provides evidence for these phenomena both in an in vitro oxidatively stressed cardiomyocyte model and in an in vivo myocardial ischemiareperfusion model. We hypothesize that this mechanism (somewhat similarly to the caspase-mediated cleavage of PARP-1 mentioned earlier [36, 37, 38] and by the recently recognized degradation of PARP-1 by ADP-ribosyl-acceptor hydrolase 3 ) serves as a protective function during oxidant-mediated cell death, since it limits the degree of PARP-1 overactivation and the associated cellular energetic catastrophe and cell necrosis. Also, this mechanism may partially restore cellular energetic pools, perhaps in an attempt of the cell to switch the mode of cell death to a more regulated form (for example, apoptosis).
Activation of the ubiquitin-proteasome system was observed in ischemia-reperfusion injury, and proteasome inhibitors have been proposed to exert a cardioprotective role in myocardial infarction (51). However, in agreement with our results, no beneficial effect of proteasome inhibition was observed in short-term reperfusion injury (52). Thus, we hypothesize that the positive effects of proteasome inhibition may be related to the inhibition of inflammatory pathways.
On the basis of several lines of data in neurons (9,53) and on the basis of the current data in cardiac myocytes, we conclude that the amount of RNF146 in a cell directly and remarkably affects the fate of the cell during oxidative injury; high levels of RNF146 are protective, whereas low levels of RNF146 (for example, after siRNA-mediated silencing) exacerbate cell injury. Whereas many mechanistic details remain to be further defined, we propose that pharmacological upregulation/therapeutic induction of RNF146 may represent a future approach for experimental therapeutic intervention of oxidant-mediated cardiac injury.
The authors declare that they have no competing interests as defined by Molecular Medicine, or other interests that might be perceived to influence the results and discussion reported in this paper.
This work was supported by the National Institutes of Health (to C Szabo) (R01GM056687).
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