PARP-1 Inhibition Prevents Oxidative and Nitrosative Stress–Induced Endothelial Cell Death via Transactivation of the VEGF Receptor 2
Objective— PARP-1, a DNA base repair enzyme, is activated by DNA breaks induced by oxidative (ROS) and nitrosative (RNS) stress. By consuming NAD+, PARP-1 activation can lead to ATP depletion and cell death. Studies suggest that inhibiting PARP-1 activity can attenuate pathologies associated with vascular smooth muscle and endothelial dysfunction. PARP-1 inhibition can also activate the prosurvival serine/threonine kinase, Akt. Vascular endothelial growth factor (VEGF) regulates endothelial cell survival via Akt activation downstream of VEGF receptor 2 (VEGFR2) activation. Here we investigated the hypothesis that PARP-1 inhibition protects human umbilical vein endothelial cells (HUVECs) from ROS- and RNS-induced cell death by limiting NAD+ depletion and by activating a prosurvival signaling pathway via VEGFR2 phosphorylation.
Methods and Results— We activated PARP-1 in HUVECs by treatment with hydrogen peroxide (H2O2) and peroxynitrite (ONOO−). Both depleted HUVECs of NAD+ and ATP, processes that were limited by the PARP-1 inhibitor, PJ34. ONOO− and H2O2-induced cell death and apoptosis were attenuated in cells treated with PJ34 or PARP-1 siRNA. PARP-1 inhibition increased Akt, BAD, and VEGFR2 phosphorylation in HUVECs and in PJ34-treated rabbit aortas. The VEGFR2-specific tyrosine kinase inhibitor SU1498 decreased PARP-1 inhibition-mediated phosphorylation of VEGFR2 and Akt, and also reversed survival effects of PJ34. Finally, PARP-1 inhibition protected cells from death induced by serum starvation, evidence for a role in cell survival independent of energy protection.
Conclusions— PARP-1 inhibition prevents ROS- and RNS-induced HUVEC death by maintaining cellular energy in the form of NAD+ and ATP, and also by activating a survival pathway via VEGFR2, Akt, and BAD phosphorylation.
Several studies have shown that inhibiting poly-ADP ribose polymerase-1 (PARP-1) attenuates organ dysfunction in settings such as postmyocardial infarction remodeling,1 ischemia-reperfusion injury,2 diabetic retinopathy,3 septic shock,4 diabetes,3,5 and atherosclerosis.6 A major feature of atherosclerosis includes increased levels of reactive nitrosative species (RNS) and reactive oxygen species (ROS) associated with damage to cell membranes and DNA.7,8 It is not surprising therefore that PARP-1 inhibition was also shown to limit endothelial dysfunction and atherosclerosis in the ApoE−/− mouse.6,9 High levels of ROS and RNS, such as occur in ischemia-reperfusion injury, inflammation, and diabetes mellitus, induce DNA single-strand breaks and activate poly-ADP ribose polymerase-1 (PARP-1).10
PARP-1 is a zinc finger protein that belongs to a family of 18 identified genes that transcribe poly(ADP-ribose) polymerases, enzymes that catalyze the covalent transfer of poly-ADP units from NAD+ to acceptor proteins. PARP-1 has 3 functional domains: a DNA-binding domain (containing a nuclear localization signal), an automodification domain (which acts as an acceptor for poly ADP-ribose units), and a C terminus catalytic domain. Normally, PARP-1 contributes to DNA base excision repair and the maintenance of genomic stability. However, when overactivated by DNA damage induced by ROS/RNS, PARP-1 rapidly uses the substrate β-NAD+ to transfer poly ADP-ribose (PAR) to itself and to nuclear acceptor proteins.11 In an effort to resynthesize NAD+, the cell consumes its ATP pools and reaches an energy crisis, resulting in cell death.
Although PARP-1 inhibition limits cellular energy depletion, recent evidence suggests that it activates a prosurvival signaling cascade through Akt phosphorylation as well.12 Little is known about the upstream mediators of Akt activation. In endothelial cells, VEGF is one of the primary regulators of cell survival,13 via activation of the VEGF receptor (VEGFR2), and subsequent activation of the PI3K/Akt axis.14 Here we investigated, using oxidative (H2O2) and nitrosative (ONOO−) stress, the mechanisms by which inhibiting PARP-1 inhibition protect against EC death. We demonstrate that PARP-1 inhibition attenuates ATP and NAD+ depletion and decreases EC death via a VEGFR2-mediated prosurvival pathway.
Materials and Methods
PARP-1 inhibitor PJ34 was purchased from Sigma. PARP-1 siRNA and anti–PARP-1 polyclonal antibody were from Santa Cruz Biotechnology. SU1498 and ATP luciferase assay kit were purchased from Calbiochem/EMD Biosciences. Antiphosphotyrosine 4G10 (pY-4G10) was from Upstate Biotechnologies. Antibodies to phospho-Akt (Ser-473), Flk-1 (VEGFR2), Akt, phospho-Bad (Ser-136), and Bad were from Cell Signaling Technologies. The NAD+/NADH Assay kit was purchased from BioAssay Systems.
HUVECs were isolated from human umbilical veins and seeded onto gelatin-coated 60-mm dishes maintained in Medium 200 (Cascade Biologics) with low serum growth supplement and 5% FBS as previously described.15 Cells were used at passages 2 to 5.
To make Na+ONOO−, 0.6 mol/L NaNO2 and 0.7 mol/L acidified H2O2 were mixed via 2 15-mL syringes connected by rubber tubing and a stopcock. The mixture was quickly quenched in a beaker with 1.5 mol/L NaOH. The resulting yellow solution was stirred with MnO2 until the disappearance of all bubbles, decanted, and placed in −20°C for 2 h. The yellow top layer was scraped with a spatula into an Eppendorf tube and the concentration was determined by diluting it 1:1000 in 10 mmol/L NaOH and reading absorbance in a spectrophotometer (extinction coefficient=16700 mol/L−1).
PARP-1 Activity Assay
PARP-1 activity was assayed (R&D Systems) based on the incorporation of biotinylated ADP-ribose onto histone proteins. Cell lysates from HUVECs containing 50 μg of protein were loaded into a 96-well plate coated with histones and biotinylated poly ADP-ribose, allowed to incubate for 1 hour, treated with strep-HRP, and read at 450 nm in a spectrophotometer.
Cell Death Assays
HUVECs were grown to confluence in 60-mm dishes, serum-starved overnight, treated with H2O2, and ONOO−, fixed with 3.7% formaldehyde 8 h after treatment, and stained with Hoescht dye. Cells were examined under a fluorescent microscope and counted for DNA condensation in 3 fields. For serum starvation, HUVECs were grown to confluence in 60-mm dishes and serum-starved at the indicated concentrations for 24 h. Cells were trypsinized, stained with trypan blue, and counted in a hemocytometer under an upright microscope. Cells were counted in 3 fields, and the average cell number was calculated.
Measurement of NAD+
HUVECs were cultured in 60-mm plates and treated with varying concentrations of H2O2 and ONOO− for 3 h. After exposure, cells were washed with cold PBS and pelleted. They were homogenized with NAD extraction buffer (BioAssay Systems), heated at 60°C for 5 min, and diluted in assay buffer (BioAssay Systems). Samples were vortexed and spun at 14 000 rpm for 5 min. 40 μL of sample was added to each well of 96-well microtiter plate. 80 μL of the working reagent (50 μL assay buffer, 1 μL alcohol dehydrogenase, 10 μL 1v% ethanol, 14 μL PMS, and 14 μL MTT) was added quickly to each well. The reaction was measured at 565 nm at 37°C for 15 min. The difference in optical density between 0 and 15 min was used to calculate NAD+ with a prepared standard curve (0 to 10 μmol/L of NAD+). The calculated NAD+-levels for each experiment were average values of at least 4 measurements.
Assay of Cellular ATP
ATP was measured using the ATP luciferin assay kit (Calbiochem). Cells were cultured in a 96-well microtiter plate and treated with varying concentrations of H2O2 and ONOO− for 3 h. Culture medium was removed and cells were washed twice with PBS and treated with 100 μL nucleotide-releasing reagent. After 5 min at room temperature, 1 μL of luciferin-luciferase nucleotide-monitoring reagent was added and the plate loaded into a luminometer and read 1 min later. ATP levels were calculated against uninduced controls and expressed in relative light units.
HUVECs were seeded into a 96-well plate at a density of 105 cells in 120 mL of media and cultured for 24 h. They were treated with 200 μL of H2O2 or 150 μL ONOO− for varying times. LADH activity was assayed using a kit (Cayman Chemical). Briefly, lactate and INT (a tetrazolium salt) were added to each well. The conversion of lactate to pyruvate and subsequent NADH production was monitored by the colorimetric reduction of INT at 490 nm.
HUVECs were transiently transfected with siRNA designed for PARP-1 knockdown by the manufacturer (Santa Cruz Biotechnology) in Opti-MEM I Reduced Serum Medium (Invitrogen) using Lipofectamine 2000. Experiments were performed 48 h after the transfection. 10 ng of PARP-1 siRNA and scrambled siRNA were used.
Immunoprecipitation and Western Blotting
HUVECs were washed twice in cold PBS and harvested in lysis buffer containing 20 mmol/L Tris pH 7.5, 150 mmol/L NaCl, 1 mmol/L EDTA, 1 μmol/L EGTA, 1% Triton X-100, 2.5 μmol/L sodium pyrophosphate, 1 μmol/L β-glycerolphosphate, 100 mmol/L NaVO4, 1 mol/L NaF, and protease inhibitor. The protein concentration of the lysates was determined using the Bradford method (Bio-Rad). Equal amounts of protein were incubated with a specific antibody overnight at 4°C with gentle rotation. Then protein A/G PLUS-agarose (Santa Cruz) was added and incubated for additional 2 h. Then the beads were washed extensively with lysis buffer, and immune complexes were eluted in SDS-PAGE sample buffer. Total immune complex samples or protein samples from total cell lysate were separated by SDS-PAGE, transferred to a nitrocellulose membrane, and incubated with appropriate primary antibodies. After washing and incubating with secondary antibodies (LiCor), immunoreactive proteins were visualized with the Odyssey LiCor Infrared Imaging System.
Perfusion Organ Culture
Animal experiments were performed according to the guidelines of the NIH and the American Heart Association for the care and use of laboratory animals and were approved by the University of Rochester Animal Care Committee. Male New Zealand White rabbits (2 to 3 kg; Covance Research Products Princeton, NJ) were anesthetized with ketamine (50 mg/kg, i.v.) and xylazine (2 mg/kg, i.v.). Arterial segments from the descending thoracic aorta were isolated and cannulated at a constant pressure (80 mm Hg) as described previously.15 To obtain a physiological fluid viscosity (0.04 poise), 5% dextran (Sigma-Aldrich) was added. Vessels were perfused with PBS containing 10 ng/mL VEGF or 10 mmol/L PJ34 for 30 min. To harvest endothelial cells, vessels were opened longitudinally and 0.2 mL of lysis buffer (20 mmol/L Tris-HCl, pH 8.0, 0.05% Triton X-100, 150 mmol/L NaCl, 2mmol/L EDTA, 50 mmol/L sodium fluoride, 2 mmol/L sodium orthovanadate, and protease inhibitor cocktail [Sigma]) was applied to the endothelial surface at room temperature for 6 min and collected.
Group differences were analyzed using the standard Student t test. All values are expressed as mean±SE. P<0.05 was considered statistically significant.
Hydrogen Peroxide and Peroxynitrite Increase PARP-1 Activity, Which Is Inhibited by PJ34
Because both H2O2 and ONOO− have DNA-damaging capacities, we studied their ability to activate PARP-1. HUVECs were exposed to H2O2 and ONOO− for 3 h, and PARP-1 activity was assayed. H2O2 activated PARP-1 in a dose-dependent manner with an EC50 of 200 μmol/L, and a maximum 8-fold increase; ONOO− at equimolar concentrations was 2.5 times more effective (Figure 1A). To inhibit PARP-1 we used PJ34, a water-soluble phenanthridinone-derived PARP-1 inhibitor.16 PJ34 inhibited both H2O2 and ONOO−-induced PARP-1 activity in a dose-dependent manner with an IC50 of ≈6.5 μmol/L (supplemental Figure IA).
Hydrogen Peroxide and Peroxynitrite Induce NAD+ Depletion That Is Reversed PARP-1 Inhibition
After confirming that H2O2 and ONOO− activate PARP-1 whereas PJ34 inhibits this enzyme in HUVECs, we examined their effects on energy consumption. PARP-1 uses NAD+ as a substrate to generate branching polymers of ADP-ribose, leading to ATP and NAD+ loss. To monitor intracellular NAD+, we used an enzymatic assay based on alcohol dehydrogenase activity. We found that NAD+ levels dropped 68% after a 3-hour exposure to H2O2 (Figure 1B) and 77% after a 3-hour exposure to ONOO− (Figure 1B). This NAD+ loss was limited to 20% for H2O2 and 40% for ONOO− with PJ34 (10 μmol/L) pretreatment (Figure 1B).
In addition to NAD+ content, we also studied ATP as a measure of energy consumption. We used an assay based on luciferase-catalyzed oxidation of D-luciferin in the presence of ATP and oxygen, whereby the amount of ATP is quantified by the amount of light produced. To inhibit PARP-1, we used both PARP-1 siRNA and PJ34. PARP-1 siRNA successfully decreased PARP-1 expression in HUVECs to 15% of control at 10 nmol/L (supplemental Figure IB). We treated HUVECs with H2O2 at varying concentrations for 3 h and found that at all concentrations of H2O2, PARP-1 siRNA, and PJ34 significantly increased ATP content (Figure 1C).
ROS- and RNS-Induced HUVEC Death Can Be Attenuated by PARP-1 Inhibition
To rule out the possibility of cell leakage contributing to ATP and NAD+ depletion, we assayed HUVECs for LDH release after H2O2 and ONOO− treatment (supplemental Figure IIA). There was no significant LDH leakage 3 h after ROS/RNS treatment. Because NAD+ depletion and ATP consumption are associated with cell death,10 we assayed the effect of PARP-1 inhibition on H2O2 and ONOO−-mediated cell death. PARP-1 siRNA-transfected HUVECs experienced a ≈60% reduction in cell death compared to scrambled siRNA-transfected controls (Figure 2A). In addition, when apoptosis was measured using Hoescht stain, PARP-1 siRNA reduced the number of apoptotic nuclei after 8 h by ≈25% (Figure 2B). PJ34 also reduced HUVEC death induced by H2O2 (67% decrease, Figure 2A) and by ONOO− (60% decrease, supplemental Figure IIB). Cells transfected with PARP-1 siRNA, and then treated with PJ34 exhibited a decrease in apoptosis (Figure 2B) that did not differ significantly from the effects of PJ34 or PARP-1 siRNA alone. This result suggests that PARP-1 inhibition attenuates apoptosis primarily through the inhibition of PARP-1 ADP-ribosylation activity.
PARP-1 Inhibition Increases Akt and BAD Activation in Response to ROS and RNS
Although our data support the hypothesis that PARP-1 inhibition limits energy consumption and mitigates ROS- and RNS-induced cell death, we were interested in the possibility of other PARP-1–mediated survival mechanisms. Specifically, PARP-1 inhibition increases Akt phosphorylation in neuronal and liver cells exposed to oxidative stress.12 To investigate the relevance of this signaling pathway in endothelial cells, we treated HUVECs with H2O2 and PJ34 and measured Akt phosphorylation. Akt phosphorylation was enhanced nearly 9-fold by PJ34 (Figure 3A). To confirm that this result was specific to PARP-1 inhibition, we also transfected cells with PARP-1 siRNA. Similar to PJ34, PARP-1 knockdown increased Akt phosphorylation by ≈9-fold after H2O2 exposure (supplemental Figure IIIA).
Because PARP-1 inhibition increased Akt phosphorylation, we hypothesized that PARP-1 inhibition would also increase the phosphorylation of downstream Akt substrates, such as BAD.17 We confirmed a 7-fold increase in BAD phosphorylation (pBAD) in PJ34-treated HUVECs stimulated with H2O2 (Figure 3A). A 7-fold increase in pBAD was also observed when we used PARP-1 siRNA to inhibit PARP-1 (supplmental Figure IIIB).
We next determined whether PARP-1 inhibition activated Akt in the setting of nitrosative stress. We treated HUVECs with ONOO− and PJ34 and assayed for pAkt and pBAD. Similar to the results with H2O2, ONOO− in conjunction with PJ34 increased Akt phosphorylation by 6-fold and BAD phosphorylation by 8-fold (Figure 3B).
PARP-1 Inhibition Increases the Association of BAD With 14-3-3
When phosphorylated on Ser136, BAD associates with the multifunctional phosphoserine-binding protein 14-3-3.18 This association inhibits BAD-induced cell death. To confirm that the phosphorylation of BAD elicited by PJ34 was part of this signaling cascade, we assayed for the association of BAD with 14-3-3 after H2O2 and PJ34 treatment (supplemental Figure IIIC). Under control conditions, the basal association of BAD and 14-3-3 was enhanced 2-fold by PJ34. This interaction was increased slightly by H2O2, but rose significantly to nearly 7-fold when cells were pretreated with PJ34. These data indicate that PARP-1 inhibition can phosphorylate BAD and increase its association with 14-3-3, thus inhibiting its apoptotic effects.
PARP-1 Inhibition Activates the VEGF Receptor 2
To determine the mechanism by which PARP-1 inhibition stimulates Akt and BAD phosphorylation, we studied proteins upstream of Akt involved in survival pathways. VEGF is a well-established vascular survival factor, and its activity can be modulated by poly-ADP ribosylation (PARsylation).19 Recent data have also implicated PARP-1 inhibition in angiogenesis,20 a process mediated in part by VEGFR2. We therefore studied the effects of PARP-1 inhibition on VEGFR2 phosphorylation to define the mechanism of Akt phosphorylation.
Cells were treated with VEGF, H2O2, or PJ34 and lysates were probed for phosphotyrosine using the 4G10 antibody. Phosphorylation of a 230-kDa band increased 22-fold after 15 min of VEGF treatment (Figure 4, lane 2), but was attenuated by pretreatment with the VEGFR2-specific tyrosine kinase inhibitor SU1498 (Figure 4, lane 3). Phosphorylation of a band of identical molecular weight to the VEGFR2 similarly increased after 15 min of PJ34 exposure alone (Figure 4, lane 4), which was also decreased by SU1498 pretreatment (Figure 4, lane 5). Akt phosphorylation was increased by PJ34 (Figure 4, lane 4), but attenuated by SU1498 (Figure 4, lane 5) indicating that PJ34-mediated Akt phosphorylation correlates with VEGFR2 phosphorylation (Figure 4, lane 2).
Next, we immunoprecipitated HUVEC lysates with antiphosphotyrosine (4G10) and probed for VEGFR2. Treatment with VEGF increased immunoprecipitation of a 230-kDa band consistent with the VEGFR2×10-fold (supplemental Figure IVA, lane 2), which was decreased by SU1498 (supplemental Figure IVA, lane 3). PJ34 treatment for 15 min increased VEGFR2-immunoreactive protein by 7-fold, which was reversed by SU1498 (supplemental Figure IVA, lanes 4 and 5, respectively). Similarly, PARP-1 siRNA strongly increased VEGFR2 immunoprecipitation, which was prevented by pretreatment with SU1498 (supplemental Figure IVA, lanes 6 and 7).
To confirm the in vivo relevance of this signaling cascade, we isolated arterial segments from the descending thoracic aorta of rabbits and perfused them ex vivo with the PARP-1 inhibitor, PJ34, for 30 min. We found that PJ34 induced phosphorylation of VEGFR2 and Akt, similar to treatment with VEGF (Figure 5).
PARP-1 Inhibition Confers Survival Advantage to Endothelial Cells Independent of Energy Protection
To investigate whether the observed VEGFR2 phosphorylation played a role in the prosurvival effects of PARP-1 inhibition, we treated HUVECs with H2O2, PJ34, and the SU1498 and counted cell number after 24 h. Inhibiting VEGFR2 limited the prosurvival effects of PARP-1 inhibition by ≈40% (Figure 6), indicating that signaling through VEGFR2 confers protection to HUVECs from oxidative stress–induced death. To confirm that PARP-1 inhibition could confer a survival effect independent of energy protection, we induced HUVEC death using serum starvation. After pretreating HUVECs with varying concentrations of PJ34 and serum-starving cells for 24 h, we found that PARP-1 inhibition conferred a statistically significant survival advantage at 10 μmol/L and above (supplemental Figure IVB). To prove further the energy independent effects of PARP-1, we studied knockdown of PARP-1 with siRNA. Similar to PJ34, PARP-1 siRNA protected HUVECs from serum starvation–induced death (supplemental Figure IVC).
The major findings of this study are that PARP-1 inhibition stimulates tyrosine phosphorylation of VEGFR2 and activation of Akt, thereby promoting endothelial cell survival. VEGF is an established regulator of EC apoptosis13 and growth,21 whereas VEGFR2 transactivation by H2O2 and fluid shear stress22 have been implicated in EC survival. Here we found that PARP-1 inhibition, using either pharmacological inhibition (PJ34) or RNA knockdown (PARP-1 siRNA), increased tyrosine phosphorylation of VEGFR2 (Figure 4) and attenuated ROS- and RNS-induced cell death (Figure 6). In contrast, the VEGFR2-specific tyrosine kinase inhibitor SU1498 prevented PJ34-mediated VEGFR2 phosphorylation and limited cell survival (Figures 4 and 6⇑).
PARP-1 inhibition has been reported to be beneficial in the settings of inflammation,2,4 diabetes,23,24 sepsis,4 shock, and ischemia-reperfusion.2 A recent study demonstrated that PARP-1 inhibition promoted plaque stability in ApoE−/− mice by inhibiting foam cell apoptosis.6 The PJ34 protective mechanism is thought to be attributable to the maintenance of mitochondrial membrane potential and intracellular NAD+. Indeed, we found that PJ34 and PARP-1 siRNA both significantly limited ROS- and RNS-induced NAD+ depletion, ATP loss, and EC death (Figures 1 and 2⇑). However, studies suggest that PARP-1 inhibition can also regulate prosurvival signaling pathways by increasing Akt phosphorylation.2,12
We found that PARP-1 inhibition increased Akt phosphorylation in HUVECs, with or without ROS/RNS stimulation (Figure 3). We also found that PJ34 and PARP-1 siRNA increased phosphorylation of BAD and its association with 14-3-3. BAD is a proapoptotic protein of the Bcl-2 family. When active, Akt phosphorylates BAD on Ser136, causing BAD to dissociate from the Bcl-2/Bcl-X complex and lose its proapoptotic function. In agreement with previously published data, ONOO− inhibited Akt and BAD phosphorylation,25 an effect that was partially reversed by PJ34 (Figure 3).
Our data also indicate that PARP-1 inhibition can attenuate serum starvation–induced HUVEC death (supplemental Figure IVB and IVC), arguing that the survival advantage conferred by PARP-1 inhibition is partially independent of energy protection. PJ34 and PARP-1 siRNA enhanced cell survival under conditions of serum starvation by nearly 50%, presumably by activating a survival signaling cascade involving VEGFR2-Akt-BAD.
Taken together, these data imply that PARP-1 constitutively acts, either directly or indirectly, as a negative regulator of VEGFR2 and Akt phosphorylation. The question remains how a nuclear localized enzyme such as PARP-1 can exert effects on the signaling cascade of a membrane-bound receptor. Although there is little documentation of cytoplasmic PARP-1, a recent report concludes that the HIV viral protein R recruits PARP-1 to a cytoplasmic glucocorticoid receptor complex and prevents its nuclear localization.26 In a similar fashion, it is possible that a discrete pool of cytoplasmic PARP-1 in endothelial cells changes its direct association with cytoplasmic or membrane proteins depending on its catalytic activity, thereby regulating VEGFR2 phosphorylation.
Another possibility is that poly-ADP ribosylation (PARsylation) of a cytoplasmic protein associated with the VEGFR2 signaling cascade could regulate VEGFR2 phosphorylation. VEGFR2 signaling is a complex process involving ligand binding and recruitment of adaptor and scaffolding proteins such as PI-3 kinase, Grb2, Gab1, and SHP2 to phosphotyrosines on the transmembrane receptor. These proteins, when PARsylated, may influence VEGFR2 phosphorylation. A recent example is the recruitment of tankyrase, a PARP-1 family member scaffold, from the cytoplasm to the membrane after PARsylation.27
PARP-1 inhibition may prove beneficial in limiting the initiation and progression of vascular disease because of its abilities to protect against ROS/RNS-induced energy depletion, to stimulate prosurvival signaling pathways, and to prevent NFκB activation and the transcription of inflammatory genes.4,28,29 A therapeutic concern may be the effect of chronic PARP-1 inhibition on genomic stability because of the role of PARP-1 in DNA repair and genomic stability. However, PARP-1 knockout mice are viable and only susceptible to genetic damage when exposed to high levels of radiation.30 DNA base excision repair involves a number of enzymes, many of which likely compensate for PARP-1, including PARP-2. Therefore, we believe that inhibiting PARP-1 will have minimal effects on genomic stability under physiological conditions. The long-term effects of PARP-1 inhibition on endothelial function will undoubtedly involve changes in gene expression and inflammation, consequences that deserves careful consideration in the future.
In conclusion, we believe that PARP-1 inhibition will improve endothelial function by preventing energy depletion, enhancing a survival pathway via VEGFR2/Akt/BAD phosphorylation, and by limiting inflammation via NFkB, thus making PARP-1 a valid therapeutic target for vascular disease.
Original received July 5, 2007; final version accepted January 15, 2008.
Palfi A, Toth A, Hanto K, Deres P, Szabados E, Szereday Z, Kulcsar G, Kalai T, Hideg K, Gallyas F Jr, Sumegi B, Toth K, Halmosi R. PARP inhibition prevents postinfarction myocardial remodeling and heart failure via the protein kinase C/glycogen synthase kinase-3β pathway. J Mol Cell Cardiol. 2006; 41: 149–159.
Oumouna-Benachour K, Hans CP, Suzuki Y, Naura A, Datta R, Belmadani S, Fallon K, Woods C, Boulares AH. Poly(ADP-ribose) polymerase inhibition reduces atherosclerotic plaque size and promotes factors of plaque stability in apolipoprotein E-deficient mice: effects on macrophage recruitment, nuclear factor-κB nuclear translocation, and foam cell death. Circulation. 2007; 115: 2442–2450.
Stocker R, Keaney JF Jr. Role of Oxidative Modifications in Atherosclerosis. Physiol Rev. 2004; 84: 1381–1478.
Bennett MR. Reactive oxygen species and death: oxidative DNA damage in atherosclerosis. Circ Res. 2001; 88: 648–650.
Tapodi A, Debreceni B, Hanto K, Bognar Z, Wittmann I, Gallyas F Jr, Varbiro G, Sumegi B. Pivotal role of Akt activation in mitochondrial protection and cell survival by poly(ADP-ribose)polymerase-1 inhibition in oxidative stress. J Biol Chem. 2005; 280: 35767–35775.
Jin ZG, Ueba H, Tanimoto T, Lungu AO, Frame MD, Berk BC. Ligand independent activation of VEGF receptor 2 by fluid shear stress regulates activation of endothelial nitric oxide synthase. Circ Res. 2003; 93: 354–363.
Yamawaki H, Lehoux S, Berk BC. Chronic physiological shear stress inhibits tumor necrosis factor-induced proinflammatory responses in rabbit aorta perfused ex vivo. Circulation. 2003; 108: 1619–1625.
Asahara T, Takahashi T, Masuda H, Kalka C, Chen D, Iwaguro H, Inai Y, Silver M, Isner JM. VEGF contributes to postnatal neovascularization by mobilizing bone marrow-derived endothelial progenitor cells. Embo J. 1999; 18: 3964–3972.
Jin ZG, Wong C, Wu J, Berk BC. Flow shear stress stimulates Gab1 tyrosine phosphorylation to mediate Akt and eNOS activation in endothelial cells. J Biol Chem. 2005; 280: 12305–12309.
Soriano FG, Pacher P, Mabley J, Liaudet L, Szabo C. Rapid reversal of the diabetic endothelial dysfunction by pharmacological inhibition of poly(ADP-ribose) polymerase. Circ Res. 2001; 89: 684–691.
Zou MH, Hou XY, Shi CM, Nagata D, Walsh K, Cohen RA. Modulation by peroxynitrite of Akt- and AMP-activated kinase-dependent Ser1179 phosphorylation of endothelial nitric oxide synthase. J Biol Chem. 2002; 277: 32552–32557.
Muthumani K, Choo AY, Zong WX, Madesh M, Hwang DS, Premkumar A, Thieu KP, Emmanuel J, Kumar S, Thompson CB, Weiner DB. The HIV-1 Vpr and glucocorticoid receptor complex is a gain-of-function interaction that prevents the nuclear localization of PARP-1. Nat Cell Biol. 2006; 8: 170–179.
Yeh TY, Meyer TN, Schwesinger C, Tsun ZY, Lee RM, Chi NW. Tankyrase recruitment to the lateral membrane in polarized epithelial cells: regulation by cell-cell contact and protein poly(ADP-ribosyl)ation. Biochem J. 2006; 399: 415–425.
Hassa PO, Covic M, Hasan S, Imhof R, Hottiger MO. The enzymatic and DNA binding activity of PARP-1 are not required for NF-κB coactivator function. J Biol Chem. 2001; 276: 45588–45597.
Hassa PO, Haenni SS, Buerki C, Meier NI, Lane WS, Owen H, Gersbach M, Imhof R, Hottiger MO. Acetylation of PARP-1 by p300/CBP regulates coactivation of NF-κB-dependent transcription. J Biol Chem. 2005; 280: 40450–40464.