PJ34, a PARP1 inhibitor, promotes endothelial repair in a rabbit model of high fat diet-induced atherosclerosis

ABSTRACT

Objective: Atherosclerosis involves endothelial injury caused by oxidative stress. Endothelial progenitor cells (EPCs) play important roles in preventing the early stages of atherosclerosis. Meanwhile, poly (ADP-ribose) polymerase 1 (PARP1) utilizes nicotinamide adenine dinucleotide (NAD⁺) to repair DNA damage. PARP1 overactivation results in excessive NAD⁺ consumption in the presence of pathological DNA damage. PJ34 is a PARP1 inhibitor that attenuates cellular NAD+ depletion and can prevent endothelial dysfunction. However, few studies have been conducted on its effects on EPCs. In this study, we tried to elucidate the action of PJ34 in rabbit EPCs and tested its effectiveness in rabbit atherosclerosis.

Methods: We analyzed the effect of PJ34 supplementation by inducing oxidative damage by H₂O₂ in vitro and using a rabbit atherosclerosis model induced by a high-fat-diet in vivo. Transwell, immunofluorescence, Matrigel, and western blot analyses, as well as adenoviral vector transfection were used to quantify the levels of reactive oxygen species, proteins, and NAD⁺.

Results: The effects of PJ34 were dependent on SIRT1 levels. In vitro results showed that when oxidative damage attenuated cellular function, PJ34 treatment restored partial functionality. In vivo results confirmed that PJ34 can prevent atherosclerosis in a rabbit model.

1. Introduction

Atherosclerosis begins with endothelial injury and dysfunction caused by oxidative stress and is a major cause of cardiovascular disease [1]. Research has shown that endothelial progenitor cell (EPC) transplantation can effectively treat atherosclerosis in rats [2]. Further, animal experiments have revealed that EPCs play important roles in preventing the early stages of atherosclerosis and restenosis after angioplasty [3]. In recent years, several studies have revealed that the ability of EPCs to replace impaired endothelial cells is closely associated with their abundance and functionality [4]. However, the factors that play important roles in EPC function remain unclear.

Poly (ADP-ribose) polymerase 1 (PARP1) is a genome-stabilizing enzyme capable of repairing damaged DNA. PARP1 consumes the substrate nicotinamide adenine dinucleotide (NAD⁺) and transforms into protein-conjugated chains of poly ADP-ribose (PAR) units within the target protein [5]. Therefore, in many previous studies, PARP1 activity was assessed by a western blot analysis of PAR formation. Sirtuin 1 (SIRT1) is an NAD⁺-dependent deacetylase that targets various transcription factors such as tumor protein p53 [6]. The activities of both PARP1 and SIRT1 are related to the intracellular NAD⁺ levels, and can regulate cellular functions [7]. For example, when intracellular NAD⁺ was depleted, the restoration of NAD⁺ levels protected against diet-induced obesity [8]. However, PARP1 activation can result in excessive NAD⁺ consumption in the presence of pathological DNA damage to repair damaged DNA [9]. Therefore, PARP1 inhibition is of great importance under these circumstances.

PJ34 is a PARP1 inhibitor that attenuates cellular NAD⁺ depletion by inhibiting PARP1 activation [7]. It was reported that PJ34 maintained intracellular NAD⁺ levels to improve hepatic cellular functions [5]. In addition, PJ34 prevented both endothelial dysfunction and hypertension in rats [10]. However, few studies have been performed to examine the effects of PJ34 on EPCs and atherosclerosis.

Here, we treated EPCs obtained from rabbit peripheral blood with PJ34 and confirmed its effects in vitro. Next, we used a rabbit model of atherosclerosis to analyze the effect of PJ34 supplementation on oxidative damage in vivo. The current study provides important experimental evidence supporting the use of PJ34 to improve EPC function and prevent atherosclerosis.

2. Materials and methods

2.1. In vivo animal model

This study was approved by the Ethics Committee of Xinhua Hospital Affiliated with Shanghai Jiao Tong University School of Medicine. All experiments involving animals were conducted in accordance with the Guild for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication, 8^th edition, 2011). Male, New Zealand white rabbits (2500–3000 g, 20 weeks old) were ordered from Songlian Co. (Shanghai, China). The rabbits were housed under a 12:12-h light–dark cycle. Before starting the experimental protocol, the rabbits were subjected to a 2-week acclimation period. Fifteen rabbits were randomly divided into three groups, and each group contained five rabbits. Group 1 was the negative-control group and received a normal diet for 90 days. Groups 2 and 3 were fed a high-fat diet (HFD) for 90 days. Group 3 received PJ34 orally during feeding (20 mg•kg^–1•day^–1). After 90 days, the rabbits were euthanized, and the aortic arch and abdominal aorta of each rabbit were carefully resected.

2.2. In vitro study design

H₂O₂ was used to generate a model of damaged cells. The effects of PJ34 on damaged EPCs were studied in vitro. Cells at passages 3–5 (P3–P5) were incubated for 24 h with normal fresh culture medium and 1 μM PJ34, 100 μM H₂O₂, or 1 μM PJ34 + 100 μM H₂O₂. Subsequently, the culture medium was removed. Cells treated with PJ34 were incubated in fresh culture medium containing 1 μM PJ34 for 24 h, and cells subjected to other treatments were incubated in normal fresh culture medium for 24 h. PAR produced by PARP1 was detected by western blotting to analyze PARP1 activity. Detection of acetylated p53 (ac-p53) levels (regulated by SIRT1-dependent deacetylation) were performed to evaluate SIRT1 activity. The levels of p21, which is regulated by ac-p53 and p53, were also detected by western blotting to further confirm the effects of ac-p53.

2.3. EPC isolation and culture

EPCs were isolated from rabbit peripheral blood by density-gradient centrifugation using Histopaque-1083 separation solution (Sigma). After the rabbits were sacrificed, fresh rabbit peripheral blood was obtained by cardiac puncture and gently added to the separation solution. After density-gradient centrifugation (2000 rpm, 20 min, 4°C), the lowermost layer contained red blood cells, the middle layer contained white blood cells, and the uppermost layer consisted of serum. The white blood cells were withdrawn and resuspended in Endothelial Cell Growth Medium-2 (EGM-2; Lonza). Cells were seeded into 6-well plates and the cells from 10 mL of rabbit peripheral blood were plated in each well. The medium was replenished every three days, and cells were subcultured at a 1:3 ratio. EPCs were detected by immunofluorescence, using antibodies against CD31, CD34, von Willebrand factor (VWF), vascular endothelial growth factor receptor 2 (VEGFR2), and CD133 (Abcam). These markers were further detected by flow cytometry, wherein the antibodies against CD34, CD133, and VEGFR2 were purchased from Invitrogen. Dual staining was performed to assay the uptake of acetylated low-density lipoprotein (ac-LDL) labeled with the fluorescent probe 1,1ʹ-dioctadecyl-3,3,3ʹ,3ʹ-tetramethyl-indocarbocyanine perchlorate (Dil-ac-LDL; Invitrogen) and the binding of fluorescein isothiocyanate (FITC)-conjugated UEA-1 (Sigma). EPCs were incubated with Dil-ac-LDL (10 μg/mL) for 4 h at 37°C and then fixed with 4% paraformaldehyde for 20 min. Next, the cells were incubated with FITC-UEA-1 (10 μg/mL) for 1 h and stained with 4′,6-diamidino-2-phenylindole (DAPI) for 10 min at room temperature.

2.4. Tissue preparation

Excised aortic tissues were fixed in 4% paraformaldehyde for 24 h, then embedded in paraffin. Five micrometer-thick sections were obtained from each paraffin block and stained with hematoxylin and eosin (H&E). The intima thickness (plaque size) of each sample was digitized using a digital camera and analyzed under a research microscope (Olympus). Oil red O staining was performed to confirm the presence of foam cells (fat deposits).

2.5. Western blot analysis

Cellular proteins were extracted using a RIPA lysis buffer. Approximately 30 μg of protein was loaded in separate wells, resolved by 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and transferred to polyvinylidene difluoride membranes (Millipore). The proteins of interest were sequentially incubated with primary and secondary antibodies, followed by detection using enhanced chemiluminescence (Millipore). Antibodies against SIRT1, p53, and p21 were ordered from Biosynthesis Biotechnology. Anti-ac-p53, anti-β-actin, anti-mouse, and anti-rabbit antibodies were purchased from Cell Signaling Technology. An anti-PAR antibody was purchased from Invitrogen. The anti-β-actin antibody served as an internal control.

2.6. Cell viability assay

A Cell Counting Kit-8 (CCK-8, Dojindo) was used to analyze cell viability. Cells (1 × 10⁴/well) were suspended in 24-well plates. After various treatments, CCK-8 (10 μL/well) was added to each well, the cells were incubated for 3 h, and absorbance was measured at 450 nm.

2.7. Cell migration assay

Transwell plates (Corning) were used to evaluate cellular migration after treatment. Fresh EGM-2 (600 μL) was added to the bottom chambers, and 5 × 10⁴ cells suspended in 200 μL of serum-free medium were added to the top chamber and incubated at 37°C for 24 h. Transmigrated cells were fixed in 4% paraformaldehyde and stained with crystal violet.

2.8. Matrigel angiogenesis assay

Fifty microliters of Matrigel™ (BD Biosciences) was added to 96-well plates. After incubation at 37°C for 30 min, cells (2 × 10⁴/well) were seeded onto the Matrigel™ at 37°C for 6 to 8 h.

2.9. Senescence-associated beta-galactosidase (SA-β-Gal) assay

A SA-β-Gal Staining Kit (Cell Signaling Technology) was used to identify cellular senescence. After treatment, EPCs were fixed at 20°C for 15 min. Next, the EPCs were incubated in SA-β-Gal staining solution at 37°C for 12 h.

2.10. Cellular reactive oxygen species (ROS) measurement

A Reactive Oxygen Species Assay Kit (Sigma) was used to measure ROS production. Cells were seeded in a 96-well plate and pre-treated dichloro-dihydro-fluorescein diacetate (DCFH-DA) at 37°C for 30 min. The cells were then subjected to various treatments. After 6 h, the absorbance was measured using a fluorescence enzyme-labeling device.

2.11. NAD⁺ measurement

An NAD/NADH Assay Kit (Abcam) was used to evaluate intracellular NAD/NADH levels. We prepared the standard solution according to the manufacturer’s protocol and set the standard curve. The intracellular NAD/NADH levels were calculated based on the standard curve and the NAD⁺ = total NADH − NAD.

2.12. Adenovirus transfection

Adenoviral vectors expressing green fluorescent protein (Ad-GFP) and SIRT1 short hairpin RNA (Ad-sh-SIRT1) were purchased from Hanheng Biotechnology. Cells were transfected with these vectors for 6 h in serum-free medium and then incubated with fresh medium for 48 h. The transfection efficiency was analyzed by western blotting and Ad-GFP was used as a negative control.

2.13. Statistical analysis

All values are expressed as the mean ± standard error. Comparisons between two groups were analyzed by the independent samples t-test. P values <0.05 were considered statistically significant. All experiments were performed at least three times.

3. Results

3.1. EPC identification

The cells isolated from rabbit blood exhibited monolayer growth and morphological characteristics typical of EPCs. P1 cells were small and displayed typical cobblestone-like morphology, while P12 cells were larger and more irregular in shape after repeated subculture. (Figure 1(a)). Immunofluorescence confirmed the cells as EPCs based on their expression of the cell-surface antigens CD31, CD34, VWF, VEGFR2, and CD133, which are standard EPC markers (Figure 1(b)) [11,12]. By flow cytometric analysis, the positive expression rates of CD34, VEGFR2, and CD133 were 84.0%, 95.5%, and 81.3%, respectively (Figure 1(c)). Specifically, VEGFR2 is an endothelial cell marker, whereas CD34 and CD133 are progenitor cell markers [13]. By examining Dil-Ac-LDL uptake and FITC-UEA-1 binding, we further verified the cells as EPCs (Figure 1(d)) [11,12].

Figure 1.

Identification of EPCs from rabbit peripheral blood. (a) EPCs at P1 and P12. (b) EPCs characterized by immunofluorescence of CD31, CD34, VWF, VEGFR2, and CD133. (c) Flow cytometric analysis of CD34, VEGFR2, and CD133 expression. (d) Uptake of ac-LDL and binding of FITC-UEA-1.

3.2. Protein levels in EPCs with replicative senescence

In this study, we repeatedly subcultured EPCs from rabbit blood for up to 12 passages. Senescence was confirmed in SA-β-gal assays showing that positivity increased with repeated subculture (Figure 2(a)). By western blot, our results suggested that SIRT1 level attenuated with repeated subculture while PAR, ac-p53, p53, ac-p53/p53, and p21 levels increased (Figure 2(b)).

Figure 2.

Protein levels in EPCs with replicative senescence. (a) SA-β-gal staining at P3, P6, P9, and P12. (b) The expression levels of SIRT1, PAR, ac-p53, p53, and p21 analyzed by western blotting at P3, P6, P9, and P12. *P < 0.05 versus P3, **P < 0.01 versus P3, and ***P < 0.001 versus P3.

3.3. Cellular modeling and PJ34 efficiency

To verify the effects of PJ34 on EPCs, we first used H₂O₂ to induce cellular damage in young EPCs (P3–P5). Our western blot results suggested that H₂O₂ attenuated SIRT1 levels and increased PAR, ac-p53, p53, ac-p53/p53, and p21 levels, similar to that in replicative senescence (Figure 3(a)). Based on the cellular status observed under a microscope and protein expression results, 100 μM H₂O₂ was used to perform subsequent experiments.

Figure 3.

Determining the concentration of H₂O₂ and PJ34. (a) The expression levels of SIRT1, PAR, ac-p53, p53, and p21 analyzed by western blotting after treatment with various concentrations of H₂O₂. (b) PAR expression analyzed by western blotting after treatment with various concentrations of PJ34. *P < 0.05 versus control, **P < 0.01 versus control, and ***P < 0.001 versus control.

To determine the appropriate PJ34 concentration, we used various concentrations of PJ34 to inhibit PARP1 activation, and the inhibition was confirmed by decreased PAR synthesis (Figure 3(b)). Based on the cellular status observed under a microscope and protein expression results, 1 µM PJ34 was selected for subsequent experiments.

3.4. PJ34 promoted SIRT1 activity and improved EPC functionality

We used untreated EPCs as a negative control. The group treated with PJ34 alone was used as a treatment control. EPCs treated with H₂O₂ alone were considered impaired. The group treated with H₂O₂ + PJ34 was used to confirm the effects of PJ34 on impaired EPCs.

Our results showed that in the groups treated with H₂O₂ alone and H₂O₂ + PJ34, ROS levels increased 6 h after treatment (Figure 4(a)). Intracellular NAD⁺ levels were significantly attenuated in the group treated with H₂O₂ alone by the end of the experiment. Intracellular NAD⁺ levels of EPCs treated with H₂O₂ + PJ34 were partially restored versus the group treated with H₂O₂ alone, although they did decrease compared to that in the control (Figure 4(b)). Moreover, EPC function containing cell viability, angiogenesis ability, and cellular migration ability were attenuated significantly in the group treated with H₂O₂ alone. However, attenuated function was partially restored in the group treated with H₂O₂ + PJ34 (Figure 4(c–e)). Western blot analysis revealed that PAR levels decreased significantly in both groups treated with PJ34. SIRT1 levels were attenuated compared to those in the control group, and no difference was found between the groups treated with H₂O₂. In addition, ac-p53 levels and the ac-p53/p53 ratio (regulated by SIRT1), and the P21 levels increased in the H₂O₂-treated groups but were partially restored after treatment with H₂O₂ + PJ34 (Figure 4(f)).

Figure 4.

PJ34 treatment improved H₂O₂-induced EPC function damage. (a) ROS production assessed after 6 h of H₂O₂ treatment. (b) Intracellular NAD⁺ levels. (c) Cell viability evaluated by performing CCK-8 assays. (d) Angiogenesis ability evaluated on Matrigel. (e) Cellular migration ability analyzed in Transwell assays. (f) Expression levels of SIRT1, PAR, ac-p53, p53, and p21, as determined by western blot analysis. *P < 0.05 versus control, **P < 0.01 versus control, and ***P < 0.001 versus control; ^#P < 0.05 versus H₂O₂ group, ^##P < 0.01 versus H₂O₂ group, and ^###P < 0.001 versus H₂O₂ group.

Interestingly, SIRT1 levels did not change, but SIRT1 activity increased in accordance with the increased ac-p53/p53 ratio.

3.5. PJ34 improved EPC functionality through increased SIRT1 activity

To confirm the mechanism of PJ34, Ad-sh-SIRT1 was used to silence SIRT1 expression (Figure 5(a)). After transfection for 48 h, EPCs were treated with H₂O₂ and PJ34 as described in Section 3.4. Our results suggested elevated ROS levels in the group treated with H₂O₂ and the maintenance of intracellular NAD⁺ levels in the group treated with PJ34 was observed (Figure 5(b,c)) as described in Section 3.4. PJ34, however, did not restore cellular functionality in terms of cell viability, angiogenesis ability, and migration ability (Figure 5(d–f)). Western blot analysis revealed that PJ34 inhibited PARP1 activation in accordance with the PAR levels as described in Section 3.4. However, the ac-p53 and p21 levels elevated by H₂O₂ were not restored significantly by PJ34 (Figure 5(g)).

Figure 5.

PJ34 treatment had no effect on H₂O₂-induced EPC function damage after silencing SIRT1. (a) SIRT1 expression analyzed 48 h after Ad-sh-SIRT1 transfection. (b) ROS production assessed after 6 h of H₂O₂ treatment. (c) Intracellular NAD⁺ levels. (d) Cell viability evaluated by performing CCK-8 assays. (e) Angiogenesis ability evaluated on Matrigel. (f) Cellular migration ability analyzed in Transwell assays. (g) Expression levels of SIRT1, PAR, ac-p53, p53, and p21 analyzed by western blotting. *P < 0.05 versus control, **P < 0.01 versus control, and ***P < 0.001 versus control; ^#P < 0.05 versus H₂O₂ group, ^##P < 0.01 versus H₂O₂ group, and ^###P < 0.001 versus H₂O₂ group.

3.6. PJ34 prevented atherosclerosis in a rabbit model

To confirm the effects of PJ34 on atherosclerosis in vivo, rabbits were fed an HFD to generate an atherosclerosis model. Our results suggested that between the three groups tested, the intimal layer of the HFD rabbits was significantly thicker and the plaque area was significantly larger, compared to that in the control group. In the group treated with HFD + PJ34, the intimal layer was thicker than that in the control group, but thinner than that in the HFD group. The plaque area was larger than that in the control group, but smaller than that in the HFD group. The presence of foam cells (fat deposits) was confirmed by oil red O staining (Figure 6(a)).

Figure 6.

PJ34 inhibited HFD-induced atherosclerosis. (a) H&E staining was used to detect the intima thickness and plaque area. Oil red O staining was used to further confirm the presence of foam cells (fat deposits). (b) Proposed mechanism. *P < 0.05 versus control, **P < 0.01 versus control, and ***P < 0.001 versus control; ^#P < 0.05 versus HFD group, ^##P < 0.01 versus HFD group, and ^###P < 0.001 versus HFD group.

Thus, our findings suggest that in vitro, PJ34 inhibited PARP1 activation, preserved intracellular NAD⁺ levels, promoted SIRT1 activity, and prevented EPC impairment induced by H₂O₂. In vivo, PJ34 prevented HFD-induced atherosclerosis in the rabbits (Figure 6(b)).

4. Discussion and conclusion

Atherosclerosis begins with endothelial cell injury, and EPCs play an important role in repairing these injured endothelial cells [2]. Therefore, we focused on EPCs in this study. Our results demonstrated that PJ34 prevents oxidative impairment in EPCs derived from rabbit blood in vitro. Moreover, animal experiments confirmed that PJ34 prevents atherosclerosis in rabbits in vivo.

Similar to our results, a decreased SIRT1 level accompanying senescence in endothelial cells was reported by Yan et al. [14]. Further, the increased p53 and p21 levels observed in this study were consistent with the results reported by Rossman et al., who also studied endothelial cells [15]. In our study, we used H₂O₂ to damage cells, as is commonly done, such as in the study on mice pancreatic islets by Ahangarpoura et al. [16]. In addition, we confirmed that the PARP1 inhibitor PJ34 blocked PAR activation by PARP1 and thereby preserved intracellular NAD⁺ levels, consistent with the results reported by Mohamed et al. who worked on skeletal muscle from mice [7]. In our study, the preservation of intracellular NAD⁺ levels resulted in the maintenance of higher cellular function levels in the H₂O₂-treated group following PJ34 treatment. Similarly, it was reported that PARP1 inhibition rescued the short lifespan of Caenorhabditis elegans [17].

Mechanistically, H₂O₂ inhibited SIRT1 levels compared to those in the control group, which was consistent with results reported by Liu et al [18]. SIRT1 levels were stable in the groups treated with either H₂O₂ or H₂O₂ + PJ34, but the levels of proteins regulated by SIRT1 (such as ac-p53) were partly restored by treatment with H₂O₂ + PJ34. The change in SIRT1 activity was consistent with the observed cellular functions. However, the effects of PJ34 were not observed after Ad-sh-SIRT1 transfection. Many previous studies reported that SIRT1 activation protects endothelial cells from dysfunction [19]. Our results suggested that in rabbits, PJ34 improved cellular function by enhancing SIRT1 activity. We speculate that increased levels of the SIRT1 cofactor NAD⁺ were unable to promote SIRT1 expression, but instead enhanced its biological activity. This was reflected by the improvement in deacetylation levels. More importantly, our results showed that PJ34 treatment prevented atherosclerosis in rabbits, which may be attributed to PARP1 inhibition and the preservation of intracellular NAD⁺ levels.

Novel results from this study demonstrated that PARP1 and SIRT1 levels were unchanged by PJ34 treatment, but that PARP1 activity was inhibited and SIRT1 activity was enhanced by PJ34 treatment. Both PARP1 and SIRT1 regulate metabolism and utilize NAD⁺. Therefore, in vitro, PJ34 may improve EPC functionality by preserving intracellular NAD⁺ levels and increasing SIRT1 activity, without increasing SIRT1 expression. These observations can be expanded to other molecular studies. In addition, our data showing the role of PJ34 in improving EPC functionality may contribute to the development of cellular therapies for atherosclerosis.

There were several limitations to this study. Typically, results obtained for endothelial cells are similar to those obtained for EPCs. Therefore, endothelial cells may also play an important role in the PJ34-mediated molecular events described in this study; however, we did not investigate endothelial cells here. Similarly, NAD⁺ is ubiquitously produced in cells and is associated with multiple cellular functions involving many regulatory and signaling pathways. Therefore, other signaling pathways may affect the pathway studied here, and should be examined in future research. Lastly, while the use of rabbits in this model gives an idea of how this pathway may work in humans, there are obvious dietary and metabolic differences that prevent the direct application of these results in human patients.

Funding Statement

This work was supported by the National Natural Science Foundation of China [81471399]; Science and Technology Commission of Shanghai [18441905200].

Disclosure statement

No potential conflict of interest was reported by the authors.

Ethics approval

This study was approved by the Ethics Committee of Xinhua Hospital affiliated with Shanghai Jiao Tong University School of Medicine. All experiments involving animals were conducted in accordance with the Guild for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication, 8^th edition, 2011).

Consent for publication

The authors provide consent for publication.

Availability of data and materials

Datasets generated and analyzed during this study are available from the corresponding author by reasonable request.

Authors’ contributions

Siyuan Zha designed and performed all the experiments. Fei Wang provided experimental guidance. Zhiyuan Ma and Zhen Li collected human umbilical cord blood. Fang Liu and Qing Cao performed the data analysis. Siyuan Zha wrote and edited the manuscript. All authors have read and approved the manuscript.