Poly (ADP‐Ribose) Glycohydrolase‐Dependent dePARylation of PCNA Is Essential for DNA Replication

ABSTRACT

Poly (ADP‐ribosyl)ation (PARylation) plays a crucial role in DNA replication, particularly during S phase, where it is detected at replication sites on the lagging strand to facilitate Okazaki fragment processing. However, the role of dePARylation in DNA replication remains elusive. In this study, we demonstrate that poly (ADP‐ribose) glycohydrolase (PARG) is actively involved in degrading poly (ADP‐ribose) at DNA replication sites during S phase. Inhibition of PARG in S‐phase cells leads to the accumulation of DNA single‐strand breaks at replication sites. Furthermore, suppression of dePARylation during S phase impairs DNA replication, which can be rescued by wild‐type PARG but not by a catalytically inactive PARG mutant. Mechanistically, we show that PCNA, a key factor in DNA replication, is PARylated by PARP1 during S phase, which reduces its interaction with FEN1. Timely removal of PARylation from PCNA by PARG restores the interaction between PCNA and FEN1, thereby facilitating DNA replication. Taken together, our findings reveal that PARG promotes DNA replication through the dePARylation of PCNA during S phase, highlighting the critical role of PARG in DNA replication.

The schematic illustration of PARG‐mediated dePARylation in DNA replication. In the normal physiological condition, PARG functions as an eraser to remove PAR from PCNA and promote the interaction between FEN1 and PCNA (A). Interruption of PARG results in the accumulation of PARs in DNA replication sites, represses the interaction between PCNA and FEN1, and suppresses DNA replication in normal unperturbed cells (B).

Abbreviations

DDR

DNA damage response

DSBs

double‐strand breaks

KD

knockdown

PAR

poly (ADP‐ribose)

PARG

poly (ADP‐ribose) glycohydrolase

PARPs

poly (ADP‐ribose) polymerases

PARylation

poly (ADP‐ribosyl)ation

SSBs

single‐strand breaks

WT

wild type

1. Introduction

Poly (ADP‐ribosyl)ation (PARylation) is a dynamic and reversible post‐translational modification in eukaryotes, playing a pivotal role in diverse cellular processes, including DNA damage repair, DNA replication, gene transcription, and chromatin remodeling [1, 2]. In the context of DNA damage response (DDR), poly (ADP‐ribose) (PAR) is synthesized by poly (ADP‐ribose) polymerases (PARPs) at DNA lesion sites, where it modulates protein–protein and protein‐DNA interactions to facilitate DNA damage repair [3]. Beyond DDR, PARylation also occurs at replication forks, regulating fork reversal and the restart of stalled forks [4]. However, excessive accumulation of PAR is cytotoxic and can induce cell death [5]. Therefore, the timely degradation of PAR is essential for maintaining cell viability.

Several enzymes, including poly (ADP‐ribose) glycohydrolase (PARG), ADP‐Ribosylserine hydrolase (ARH3) and terminal ADP‐Ribose protein glycohydrolase 1 (TARG1), have been shown to mediate dePARylation under various conditions [6, 7]. Among them, PARG is the primary dePARylation enzyme, responsible for hydrolyzing approximately 90% of cellular PAR [8, 9, 10]. The role of PARG in cellular metabolism has been extensively studied under exogenous genotoxic stress. PARG is recruited to DNA damage sites, where it regulates the repair of single‐strand breaks (SSBs) and double‐strand breaks (DSBs), and sensitizes cells to alkylating agents and ionizing radiation [11, 12]. Moreover, inhibition of PARG leads to replication fork stalling and persistent replication stress [13]. The interaction between PARG and PCNA, which depends on the PIP‐box and the acetylation of lysine (K409) on PARG, facilitates PARG localization at replication foci during the S phase [11, 14]. Additionally, overexpression of PARG has been shown to prevent NMDA‐induced AIF translocation and cell death, and PARG transgenic mice exhibit reduced infarct volume compared with wild‐type mice [15, 16, 17]. However, the physiological role of PARG‐mediated dePARylation in DNA replication remains poorly understood.

Parg‐null mice are embryonically lethal, and pharmacological inhibition of PARG has been shown to effectively suppress tumor growth in vivo [5, 18], underscoring the critical role of PARG in unperturbed cellular metabolism. Recent studies reported that PARylation is activated by unligated Okazaki fragments at DNA replication sites during the S phase, facilitating the recruitment of DNA replication machinery to these sites [19, 20]. Based on these findings, we propose that PARG‐mediated dePARylation is a crucial mechanism to prevent the trapping of DNA replication machinery at replication forks under normal physiological conditions. Our studies demonstrate that PARP1‐mediated PARylation and PARG‐mediated dePARylation are highly active during the S phase. Specifically, PARG‐mediated dePARylation of PCNA enhances DNA replication by promoting the interaction between PCNA and FEN1. Taken together, our study illustrates the physiological roles of dePARylation in normal unperturbed cells, highlighting its importance in maintaining efficient DNA replication.

2. Materials and Methods

2.1. Plasmid Construction

The human full‐length genes encoding PARG, PCNA, FEN1, and PARP1 were amplified from human complementary DNA (cDNA) and subsequently cloned into the PHAGE vector. To generate the catalytically inactive PARG mutant (E755A/E756A), site‐directed mutagenesis was performed on the full‐length PHAGE‐Flag‐PARG plasmid, followed by digestion with the DpnI enzyme to remove the template DNA. The knockdown sequence of PARG (GCAGTTTAGTAATGCTAACAT) was designed and constructed into the PLKO.1 vector. The primers used for plasmid construction are shown in Table S1.

2.2. Cell Culture and Cell Transfection

HEK293T cells were cultured in Dulbecco's modified Eagle's medium (DMEM) (C11995, Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS) (900‐108, Gemini, USA) and 1% penicillin–streptomycin (15140‐122, Gibco). Cells were cultured in a humidified incubator at 37°C and 5% CO₂. HEK293T cells were transfected with the indicated plasmids and then subjected to other treatments at 24 h post‐transfection.

2.3. Cell Synchronization and Cell Cycle Assay

Double thymidine blocks were used to synchronize cells at the G1/S boundary. Briefly, HEK293T cells were seeded into the indicated plates. 2 mM thymidine (T9250, Sigma‐Aldrich, St. Louis, MO) was added in fresh medium and incubated with cells for 16 h. The next day, cells were washed twice with PBS to remove thymidine and then cultured in fresh medium for 8–10 h to allow cell cycle progression. This thymidine treatment cycle was repeated once to achieve optimal synchronization. The cells were treated with protein inhibitors for the indicated time following release. The cells were digested by RNase at 37°C for 30 min. Nuclear DNA was stained with propidium iodine (PI) solution (P4170, Biotopped, Beijing, China) for 30 min at room temperature. The cell cycle data were obtained using flow cytometry (CytoFLEX, Beckman, California, USA).

2.4. Stable Cell Line Construction

The PARG down‐regulated stable cell line was constructed via lentiviral infection. Briefly, HEK293T cells were co‐transfected with PLKO.1‐shPARG plasmid along with psPAX2 and pMD2.G. The replication‐defective lentivirus was collected 48 h post‐transfection and used to infect HEK293T cells in the presence of 10 μg/mL polybrene (H868728, MACKLIN, Shanghai, China). The stable cell line with down‐regulated PARG expression was screened using 1 μg/mL puromycin (P8230, Solarbio, Beijing, China).

2.5. Dot Blot and Western Blot

Total proteins of cells were harvested with NETN 300 lysis buffer [20 mM Tris–HCl (pH 7.4), 300 mM NaCl, 1 mM ethylene diamine tetraacetic acid (EDTA), 1% Triton X‐100] supplemented with protease inhibitor cocktail tablets (04693132001; Roche, Basel, Switzerland) and PARG inhibitor PDD00017273 [21, 22, 23] (5952, Tocris, Bristol, UK). Using a Pierce BCA Protein Assay Kit (PA101‐01, Biomed, Beijing, China) to test the protein concentration of the extracts. The extracts were mixed with SDS loading buffer not including bromophenol blue and boiled at 95°C for 10 min. Proteins (1 μL) were spotted onto a nitrocellulose membrane. The membrane was baked for 30 min at 60°C and then blocked with 5% milk for 1 h at room temperature. After incubation with anti‐PAR monoclonal antibody (4335‐MC‐100, R&D Systems, Minnesota, USA) overnight at 4°C, blots were developed like western blot and captured by a ChemiDoc MP Imaging System (Bio‐Rad). GAPDH was used as an internal control.

For western blot, the protein samples were obtained as described in “Dot blot.” Proteins separated by SDS‐polyacrylamide gel electrophoresis (PAGE) were transferred to polyvinylidene fluoride membranes (IPVH00010, Millipore, MA, USA) and blocked with 5% milk. The membranes were incubated with indicated antibodies (Table S2) and the signals were visualized by chemiluminescent detection.

2.6. Comet Assay

The neutral and alkaline comet assays were performed as previously described [10, 24]. Briefly, cells were washed with PBS three times following treatment. A total of 1 × 10⁵ cells/mL were combined with 1% low‐gelling‐temperature agarose (A8350, Solarbio) at 42°C at a ratio of 1:20 (v/v) and immediately pipetted onto the slides. For the alkaline comet assay, the slides were immersed in the alkaline lysis buffer (2.5 M NaCl, 100 mM EDTA, 10 mM tris–HCl, 1% sarkosyl, and 1% Triton X‐100, pH 10.0) overnight at 4°C. The next day, the slides were washed three times with ddH₂O and were submerged for 10 min in alkaline electrophoresis solution (300 mM NaOH and 1 mM EDTA, pH 13.0) at 4°C before electrophoresis at 25 V (0.6 V/cm) for 45 min. Slides were neutralized in solution (0.4 M Tris–HCL, pH 7.2–7.4) for 5 min and stained with PI (2.5 μg/mL) for 30 min in the dark. The excess stain was washed away with ddH₂O.

For neutral comet assay, the slides were immersed in the neutral lysis buffer [2% sarkosyl, 0.5 M EDTA, and proteinase K (0.5 mg/mL), (pH 8.0)] overnight at 37°C in the dark. The next day, the slides were washed with rinse buffer (90 mM tris buffer, 90 mM boric acid, and 2 mM Na₂EDTA, pH 8.5) three times. Then, the slides were subjected to electrophoresis at 20 V for 45 min (0.6 V/cm) and stained in PI (2.5 μg/mL) for 30 min in the dark. The excess stain was washed away with ddH₂O. Images were obtained using a fluorescence microscope (IX73, Olympus, Tokyo, Japan). Comet tail moment was quantified using CASPlab software (http://casplab.com).

2.7. EdU Staining and Immunofluorescence

EdU (5‐Ethynyl‐2′‐deoxyuridine) is a thymine nucleoside analogue and could replace thymine in newly synthesized DNA during DNA replication. EdU staining kit (C10310, Ribobio, Guangzhou, China) was used to evaluate DNA replication activity according to the manufacturer's instructions. Briefly, the cells were incubated with 10 μM EdU for 2 h in a humidified incubator at 37°C and 5% CO₂. After washing twice with PBS, the cells were fixed at room temperature in 4% paraformaldehyde for 30 min and permeabilized in 0.5% Triton X‐100 for 10 min on a decoloration shaker. The cells were then stained with an Apollo dyeing solution at room temperature in the dark for 30 min. The nuclei were labeled with 4′,6‐diamidino‐2‐phenylindole (DAPI) (S36939; Invitrogen, Carlsbad, CA). Images were obtained using a fluorescence microscope (IX73, Olympus).

After fixed and permeabilized like EdU staining, the cell slides were blocked with 8% goat serum for 1 h. After washing with PBS, the cell slides were incubated with indicated primary antibodies at 4°C overnight. The next day, the cell slides were washed three times with PBS and incubated with fluorescence‐labeled secondary antibody at 37°C for 1 h in the dark. The nuclei were visualized by DAPI. Images were obtained using a fluorescence microscope (IX73/LSM900, Olympus/ZEISS).

2.8. Immunoprecipitation and PARylation Assay

After treatment, cells were lysed in immunoprecipitation (IP) buffer [20 mM Tris–HCl (pH 7.4), 300 mM NaCl, 1 mM EDTA, 1% Triton X‐100] supplemented with protease inhibitor cocktail tablets (04693132001, Roche, Basel, Switzerland). The extracts were clarified by centrifugation (12,000 g, 10 min). 10% of the supernatant was applied as input, while the remaining lysate was incubated with Protein G beads (101 243, Invitrogen, CA, USA) and the corresponding antibodies for protein pull‐down at 4°C overnight. The beads were then washed twice with high‐concentration IP buffer [20 mM Tris–HCl (pH 7.4), 300 mM NaCl, 1 mM EDTA, 1% Triton X‐100] and three times with low concentration IP buffer [20 mM Tris–HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1% Triton X‐100]. Finally, the bound proteins were eluted in loading buffer by heating at 95°C for 10 min.

For the PARylation assay, cells were suspended in 80 μL of IP buffer containing protease inhibitor. The supernatants were mixed with 10 μL of 10% SDS and heated at 95°C to disrupt noncovalent protein–protein interactions. The heat‐treated samples were then diluted with 900 μL of IP buffer and subjected to ultrasonication using a Uibra‐Cell Ultrasonic Processor (Sonics, Newtown, CT). Subsequent steps followed the same protocol as described for the IP assay.

2.9. Statistical Analysis

In this study, all data were analyzed using SPSS (version 19.0, IBM, Armonk, NY) and are presented as mean ± SD. Differences between two groups were assessed using a two‐tailed Student t test. For comparisons involving multiple groups, one‐way analysis of variance (ANOVA) was employed, followed by Bonferroni post hoc analysis (data showing a normal distribution and homogeneity of variance) or Tamhane's T2 (M) analysis (data showing variance inhomogeneity). For data with a non‐normal distribution, nonparametric tests were used, including the Mann–Whitney U test for two‐group comparisons and the Kruskal‐Wallis H test for multiple group comparisons. p values less than 0.05 were considered significant.

3. Results

3.1. PARylation and dePARylation Are Dynamically Regulated in the S Phase

To investigate the dynamics of PARylation during the S phase, 293 T cells were synchronized at the G1/S boundary using thymidine and subsequently released for various time periods. Cell cycle arrest induced by thymidine treatment was confirmed by flow cytometry (Figure S1A). Using dot blot and western blot analyses, we observed significant fluctuations in PARylation levels during the S phase, with a peak occurring 2 h post‐release from the thymidine block, followed by a gradual decline to baseline levels (Figure 1A and Figure S1B). Twelve hours post‐release from the thymidine block, PAR signals initiated a new phase of periodic fluctuations (Figure 1A). Treatment with the PARP inhibitor Olaparib resulted in a progressive downregulation of PARylation levels (Figure 1B), suggesting a short half‐life of PARylation and rapid dePARylation activity during the S phase. Given that PARylation and dePARylation are primarily mediated by PARP1 and PARG, respectively, we next examined whether PAR metabolism in the S phase is regulated by these enzymes. Treatment with Olaparib (PARP inhibitor) resulted in a significant reduction in PARylation levels in 293 T cells (Figure 1C), whereas inhibition of PARG using PDD00017273 led to an accumulation of PARylation (Figure 1E). Immunofluorescence staining further demonstrated that ADP‐ribosylation (ADP‐r) signals were markedly reduced in S‐phase cells treated with Olaparib (Figure 1D), while inhibition of PARG increased ADP‐r signals (Figure 1F). Collectively, these findings underscore the dynamic interplay between PARylation and dePARylation during DNA replication in the S phase.

FIGURE 1.

PARylation and dePARylation are dynamically regulated in S phase. (A) Analysis of PAR levels during cell cycle progression. HEK293T cells were treated with 2 mM thymidine for 16 h and released from thymidine for 8 h, then treated with thymidine for the second time to synchronize cells at G1/S boundary, cells were released from thymidine for indicated time before lysed and applied to dot blot assay. PAR level was measured by PAR specific antibody, and GAPDH was used as the loading control. Shown on the left is the image of dot blot assay as well as the indication of the time released from thymidine. The density of the dot was quantified by Image lab and the PAR level in cells released from thymidine for different time was normalized to that in asynchronous cells (right), data from 3 independent experiments were included. (B) The half‐life of PAR in S phase. Cells were synchronized at S phase with double thymidine treatment, after releasing the cells from thymidine for 2 h, the synchronized cells were treated with 2 μM Olaparib for 5/20/60 min prior to being lysed and applied to dot blot assay. Shown on the left is the dot blot image and shown on the right is the statistical analysis, data from 3 independent experiments were included. (C, E) The effects of PARP or PARG inhibitor on PAR level in S phase cells. Shown is the western blot indicating PAR level in S phase cells treated with DMSO and 2 μM Olaparib (C) or 2 μM PDD00017273 (E). The synchronized cells that were pretreated with DMSO, Olaparib or PDD0017273 for 2 h were released from thymidine for 0 or 2 h to be lysed and applied to western blot assay. (D, F) The effects of PARP or PARG inhibitor on ADP‐r profile in S phase cells. The synchronized cells that were pretreated with DMSO, Olaparib or PDD0017273 for 2 h were released from thymidine for 2 h to stain with ADP‐r antibody and PCNA antibody. All nuclei were stained with DAPI (blue) (Scale bar, 10 μm). Cells stained with PCNA indicate the cells in S phase. Shown on the left are the representative images and shown on the right is the statistical analysis. All “Ctrl” represents asynchronous cells. All data are shown as the mean ± SD. *p < 0.05.

3.2. Unligated Okazaki Fragments Induce PARylation in S Phase

Previous studies have demonstrated that PARylation during the S phase is primarily driven by unligated Okazaki fragments, a byproduct of lagging‐strand DNA synthesis, rather than DNA replication stress or DNA damage [19, 20]. To further investigate the mechanism underlying active PARylation during the S phase, we analyzed the expression levels of PARP1 and PARG. Our results showed no significant changes in the expression of either PARP1 or PARG during the S phase (Figure 2A). We hypothesized that the PARylation observed in the S phase might be triggered by SSBs resulting from unligated Okazaki fragments. To test this hypothesis, we quantified SSBs and DSBs in S‐phase cells by comet assays. The results revealed a significant increase in SSBs during the S phase, as evidenced by elevated tail moments under alkaline conditions compared to neutral conditions (Figure 2B). Furthermore, inhibition of Okazaki fragment formation using emetine led to a reduction of SSBs in S‐phase cells (Figure 2C). However, a subset of SSBs persisted following emetine treatment, potentially attributable to other endogenous or exogenous stimuli. Notably, the number of XRCC1 foci, which facilitate the repair of endogenous stochastic SSBs and also exhibited during S phase [25, 26, 27, 28], was significantly reduced in emetine‐treated cells (Figure 2D). Dot blot analysis showed that cells treated with emetine exhibited lower levels of PAR synthesis compared to the mock‐treated controls (Figure 2E). These results provide compelling evidence that the SSBs generated from unligated Okazaki fragments are a significant source of PAR during S‐phase DNA replication.

FIGURE 2.

Unligated Okazaki fragments induce PARylation in S phase. (A) The protein level of PARP1 and PARG at different times post‐release from double thymidine block treatment in HEK293T cells by western blot. Samples were addressed as described for Figure 1A. (B) SSBs and DSBs levels in S phase cells assessed by comet assay under alkaline and neutral conditions, respectively. After releasing cells from thymidine for 0/2/4 h after synchronizing, cells were harvested and subjected to the comet assay. Shown in the upper panel are the representative images of cell nuclei stained with propidium iodide (PI) after electrophoresis; Scale bar, 100 μm. Shown on the lower panel is the statistical analysis of the comet tail quantified by CASPlab software under alkaline and neutral conditions, data of 100 cells from three independent experiments were included. (C) Emetine treatment significantly reduced SSBs in S phase cells. S phase cells that were released from thymidine for 1 h were treated with DMSO or 2 μM emetine for another 1 h prior to alkaline comet assay. Shown on the left are the representative images of alkaline comet assay (Scale bar, 100 μm), shown on the right is the statistical analysis. Data of 100 cells from three independent experiments were included. (D) The effects of emetine on XRCC1 foci. HEK293T cells that were released from thymidine for 1 h after synchronizing were treated with DMSO or 2 μM emetine for another 1 h prior to staining with XRCC1 antibody. Shown on the left are the representative images of XRCC1 foci, and shown on the right is the statistical analysis. Data of 50 cells from 3 independent experiments were included (Scale bar, 10 μm). (E) The image (left) and the statistical analysis (right) of dot blot assay of PAR level in HEK293T cells after DMSO, emetine (2 μM) or Olaparib (2 μM) treatment for 1 h. All “Ctrl” represents asynchronous cells. All data are shown as the mean ± SD. *p < 0.05.

3.3. Suppression of dePARylation Inhibits DNA Replication

Given that previous studies have established PARP1 as a critical sensor of unligated Okazaki fragments during DNA replication [19, 20], we sought to determine whether PARG also plays a pivotal role in this process. As illustrated in Figure 3A, cells were synchronized at the G1/S boundary using a double thymidine block, pretreated with the PARG inhibitor PDD00017273 for 1 h, and then released for the indicated time to analyze SSBs formation and DNA replication. Additionally, PARG knockdown cells were utilized to further evaluate SSBs formation and DNA replication (Figure S2A). Alkaline comet assays revealed that suppression of PARG significantly increased SSBs during the S phase, as evidenced by elevated tail moments compared to DMSO‐treated controls or scramble‐transfected cells (Figure 3B and Figure S2B). Furthermore, the percentage of EdU‐positive cells was markedly reduced following PARG inhibition, indicating that disruption of PARG impairs DNA replication (Figure 3C and Figure S2C). Collectively, these data suggest that inhibition of PARG‐mediated dePARylation during the S phase impedes DNA replication, underscoring the importance of the balance between PARylation and dePARylation for the successful completion of DNA replication.

FIGURE 3.

Suppression of dePARylation inhibits DNA replication. (A) Schematic diagram showing the details of the method used. HEK293T cells were treated with 2 mM thymidine for 16 h, after washing cells with PBS for twice, cells were cultured with fresh medium for 8 h to release cells back into cell cycle. Cells were treated with 2 mM thymidine for 16 h again to ensure that all the cells will be captured in late G1 or early S phase. The synchronized cells were released for the indicated time prior to harvest for comet assay or EdU staining. (B) PARG inhibition leads to increased SSBs in S phase. Shown are the representative images (left) and statistical analysis (right) of alkaline comet assay representing SSBs level in S phase cells treated with DMSO or PDD00017273 (2 μM). The synchronized cells that were pretreated with DMSO or PDD0017273 for 1 h were released for 0/2/4 h to harvest for comet assay. (Data of 100 cells from three independent experiments were included. Scale bar, 100 μm). “Ctrl” represents asynchronous cells. (C) PARG interruption suppressed DNA replication activity. Shown are the representative images (left) and statistical analysis (right) of EdU staining in S phase HEK293T cells. Cells were treated with 10 μM EdU for 2 h in the absence or presence of PDD00017273 (2 μM). The synchronized cells that were pretreated with DMSO or PDD0017273 for 1 h were released for 4 h to harvest. The white arrows represent the unreplicated DNA. Scale bar = 10 μm, data from 20 randomly picked images were included in each group. All data are shown as the mean ± SD. *p < 0.05.

3.4. PARylation of PCNA Suppresses Its Interaction With FEN1

Since it has been shown that PARG is recruited to DNA replication sites through a PCNA‐dependent pathway [11], we investigated whether PARG participates in PCNA‐dependent DNA replication. First, we examined the PARylation of PCNA during the S phase and found that PCNA was highly PARylated 2 h after the release from thymidine treatment (Figure 4A). Treatment with Olaparib significantly decreased the PARylation of PCNA (Figure 4B), indicating that PARP1 is the major ADP‐ribosyltransferase responsible for PCNA PARylation.

FIGURE 4.

PARylation of PCNA suppresses its interaction with FEN1. (A) The PARylation level of PCNA in S phase. HEK293T cells were synchronized in early S phase and released from thymidine for 2 and 4 h before being lysed. For the PARylation assay, the cells were suspended with 80 μL IP buffer. The supernatants were added to 10 μL 10% SDS and heated at 95°C to disrupt noncovalent protein–protein interactions. The heat‐treated samples were mixed with 900 μL IP buffer for the IP assay. Cell lysates were immunoprecipitated with anti‐HA antibody followed by being subjected to SDS‐PAGE and immunoblotted with the indicated antibodies. “Ctrl” represents asynchronous cells. Shown on the left is the western/dot blot image and shown on the right is the statistical analysis; data from 3 independent experiments were included. (B) PARylation of PCNA significantly decreased in cells treated with olaparib (2 μM). Cell lysates synchronized in S phase were immunoprecipitated with anti‐HA antibody and samples were addressed as described in (A). Shown on the left is the western/dot blot image and shown on the right is the statistical analysis. (C) Overexpression of PARP1 impairs the binding of FEN1 to PCNA. Flag‐PARP1, Myc‐FEN1, and HA‐PCNA were transfected into HEK293T cells as indicated. 24 h later, co‐immunoprecipitation was performed with HA antibody. Shown is the image (left) and statistical analysis (right) of the co‐immunoprecipitation assay showing the interaction between PCNA and FEN1 in HEK293T cells transfected with Flag‐PARP1 plasmid or not. (D) The main PARylation sites of PCNA in S phase. Cells transiently expressing HA‐tagged PCNA wild‐type (HA‐PCNA‐WT) or its mutants, HA‐PCNA‐D120A, HA‐PCNA‐D122A, HA‐PCNA‐E124A, and HA‐PCNA‐D120A/D122A/E124A (HA‐PCNA‐3A), were lysed and immunoprecipitated with anti‐HA antibody followed by being separated with SDS‐PAGE and immunoblotted with indicated antibodies, and the density of ADP‐r was quantified with Image lab. Shown are the image (top) and the statistical analysis (bottom) of the relative level of ADP‐r in immunoprecipitates. (E) Interaction between wild‐type PCNA or PCNA mutants and FEN1 in HEK293T cells. Cell lysates from cells transiently expressing Myc‐FEN1 and HA‐PCNA‐WT or HA‐PCNA‐mutants were subjected to co‐immunoprecipitation with HA antibody, and the density of Myc‐FEN1 was quantified with Image lab. Shown are the image (left) and the statistical analysis (right) of the relative level of Myc‐FEN1 in immunoprecipitates. All data are shown as the mean ± SD. *p < 0.05.

Previous studies have identified 13 potential PARylation sites on PCNA, including E93, D94, D120, D122, E124, E130, E132, D188, E197, E200, D256, E257, and E259 (Figure S3A). Among these, three sites (D120, D122 and E124) are located at the interface between PCNA and FEN1, an exonuclease that removes RNA primers during Okazaki fragment processing [29, 30] (Figure S3B). PARylation of PCNA at these sites is likely to create steric hindrance, impairing the interaction between PCNA and FEN1, potentially resulting in DNA replication defects [30, 31, 32]. Consistent with this presumption, immunoprecipitation results confirmed that overexpression of PARP1 impaired the binding of FEN1 to PCNA (Figure 4C). Immunofluorescence staining also demonstrated that the co‐localization of PCNA and FEN1 in the nucleus was decreased (Figure S3C). To further validate the functional importance of these three interface residues, we generated alanine substitution mutants (D120A, D122A, E124A, and a triple mutant 3A) and assessed their PARylation status. As shown in Figure 4D, wild‐type PCNA (PCNA‐WT) exhibited robust PARylation, whereas the D120A, D122A, E124A, and 3A mutants showed significantly reduced PARylation levels. We then evaluated the impact of these mutations on the interaction between PCNA and FEN1. Consistent with our hypothesis, PCNA‐D122A and PCNA‐E124A exhibited stronger interactions with FEN1, while PCNA‐D120A showed a weaker interaction (Figure 4E and Figure S3D). Structural modeling revealed that PCNA‐D120 forms a salt bridge with R355 of FEN1 (Figure S3B‐c). Disruption of this salt bridge in the PCNA‐D120A mutant likely explains the decreased interaction between PCNA‐D120A and FEN1. Moreover, under PDD00017273 treatment, PCNA‐D122A and PCNA‐E124A mutants promoted DNA replication, as measured by EdU incorporation, whereas PCNA‐D120A inhibited DNA replication compared to PCNA‐WT (Figure S3E). These findings indicate that PARylation of PCNA suppresses the interaction between PCNA and FEN1.

3.5. PARG‐Mediated dePARylation Facilitates the Interaction Between PCNA and FEN1

To investigate the role of PARG in DNA replication, we employed shRNA‐mediated knockdown of PARG and examined the detailed molecular events between PARG and PCNA. Under PARG inhibition, PARylation of PCNA was significantly enhanced (Figure 5A), which subsequently decreased the interaction between PCNA and FEN1 (Figure 5B). Similarly, treatment with a PARG inhibitor markedly weakened the PCNA‐FEN1 interaction and their co‐localization in the nucleus (Figure 5C and Figure S4A). To confirm whether the enzymatic activity of PARG is required in this process, we reconstituted PARG‐knockdown cells with either wild‐type PARG [PARG (WT)] or a catalytically inactive mutant PARG [PARG (E755A/E756A)]. While PARG (WT) restored the interaction between PCNA and FEN1, PARG (E755A/E756A) failed to do so (Figure 5D and Figure S4B). Collectively, these data suggest that PARG‐mediated dePARylation is an immediate downstream step in PARP‐dependent DNA replication, promoting the interaction between PCNA and FEN1 at DNA replication sites.

FIGURE 5.

Disruption of PARG inhibits the interaction between PCNA and FEN1. (A) PARylation of PCNA significantly increased in cells treated with PDD00017273 (2 μM). Cell lysates synchronized in S phase were immunoprecipitated with anti‐HA antibody before being subjected to SDS‐PAGE followed by immunoblotting with the indicated antibodies. Shown are the image (left) and the statistical analysis (right) of the proportion of PARylated PCNA when PARG is inhibited. (B) PARG deficiency impairs the binding of FEN1 to PCNA. Myc‐FEN1 and HA‐PCNA were transfected into HEK293T cells and PARG knockdown cells as indicated. 24 h later, co‐immunoprecipitation was performed with HA antibody. The density of Myc‐FEN1 was quantified with Image Lab. Shown is the image (left) and statistical analysis (right) of the co‐immunoprecipitation assay showing the interaction between PCNA and FEN1 in control cells and PARG knockdown cells. (C) Inhibition of PARG impairs the interaction between FEN1 and PCNA. Myc‐FEN1 and HA‐PCNA were transfected into HEK293T cells. 24 h later, the cells were treated with DMSO or PDD00017273 (2 μM) for 1 h and lysed for co‐immunoprecipitation assay. The density of Myc‐FEN1 was quantified with Image Lab. Shown is the image (left) and statistical analysis (right) of the co‐immunoprecipitation assay showing the interaction between PCNA and FEN1 in control cells and PARG inhibition cells. (D) Interaction between PCNA and FEN1 in PARG knockdown cells transfected with PARG (WT) or PARG (E755A/E756A). The indicated plasmids were transfected into PARG deficient HEK293T cells. The density of Myc‐FEN1 was quantified with Image Lab. Shown is the image (left) and statistical analysis (right) of the co‐immunoprecipitation assay showing the interaction between PCNA and FEN1 in the indicated cells. Data are shown as the mean ± SD. *p < 0.05, n.s, no significance.

3.6. The Enzymatic Activity of PARG Is Required for DNA Replication

Based on PARG's critical role in modulating PCNA‐FEN1 binding, we hypothesized that its enzymatic activity would be essential for both preventing SSBs accumulation from unligated Okazaki fragments and maintaining normal DNA replication progression. To address this, we employed PARG‐knockdown cells complemented with either PARG (WT) or PARG (E755A/E756A). Expressions of PARG (WT) and PARG (E755A/E756A) were confirmed by western blot analysis (Figure 6A). Alkaline comet assays revealed that SSBs resulting from unligated Okazaki fragments during DNA replication were significantly reduced in PARG‐knockdown cells expressing PARG (WT), whereas cells expressing PARG (E755A/E756A) failed to rescue this phenotype (Figure 6B). Moreover, we observed that, compared with the PARG‐knockdown group, the compromised DNA replication activity, as assessed by EdU incorporation, was markedly improved in cells expressing PARG (WT), whereas DNA replication in cells expressing PARG (E755A/E756A) remained similarly impaired as the PARG‐knockdown cells (Figure 6C). In conclusion, these findings demonstrate that the enzymatic activity of PARG is indispensable for DNA replication.

FIGURE 6.

Enzymatic activity of PARG is required for DNA replication. (A) The expression of Flag‐PARG (WT) and Flag‐PARG (E755A/E756A) in PARG knockdown cells by western blot assay. GAPDH served as the internal control. (B) Representative images (left) and statistical analysis (right) of alkaline comet assay representing SSBs level of cells expressing wild‐type PARG and its enzymatic dead mutant (E755A/E756A) (scale bar, 100 μm). PARG‐knockdown cells reconstituted with wild‐type PARG and its enzymatic dead mutant (E755A/E756A) were synchronized by double thymidine block and released for the indicated time. “Ctrl” represents asynchronous cells. Data of 100 cells from three independent experiments were included. (C) Representative images (left) and statistical analysis (right) of EdU staining in cells complemented with wild‐type PARG and PARG mutant (E755A/E756A). The control cells, PARG‐knockdown cells, PARG knockdown cells complemented with PARG (WT) and PARG knockdown cells transfected with PARG (E755A/E756A) were released from thymidine for 2 h after synchronizing and were treated with 10 μM EdU for another 2 h prior to EdU staining (n = 20 in each group; scale bar, 10 μm). The white arrows represent DNA that is not replicated. Data are shown as the mean ± SD. *p < 0.05.

4. Discussion

PARylation is a reversible post‐translational modification that plays a critical role in various cellular processes [2, 33]. Over the past decade, PARPs have emerged as promising therapeutic targets in cancer treatment [34, 35]. Given the DNA replication vulnerabilities induced by PARG inhibitor in ovarian cancer cells and the fact that the catalytic pocket of PARG can be easily inhibited by small molecules, PARG inhibitors represent a potential therapeutic strategy for cancer [13]. Notably, Hanzlikova et al. demonstrated that PARP1 acts as a key sensor of unligated Okazaki fragments during DNA replication [19], promoting us to investigate whether PARG is also essential for this process. In this study, we identified PARG‐mediated dePARylation as an active and dynamic process during the S phase, with PARG serving as the primary enzyme responsible for removing PAR from PARylated proteins during this phase. Furthermore, we demonstrated that PARG regulates DNA replication by removing PAR from PCNA, thereby facilitating the binding of PCNA to FEN1 (Figure 7A). Disruption of PARG led to the accumulation of PAR at DNA replication sites, impaired the PCNA‐FEN1 interaction, and ultimately suppressed DNA replication in normal unperturbed cells (Figure 7B).

FIGURE 7.

Working Models. (A) The schematic illustration of the interaction between PCNA and FEN1 in the normal physiological condition. In this model, PARG functions as an eraser to remove PAR from PCNA and promote the interaction between FEN1 and PCNA. (B) Schematic illustration of impaired PCNA binding with FEN1 after suppression of dePARylation of PCNA. Inhibition of PARG results in PAR accumulation on PCNA at DNA replication sites and represses the interaction between PCNA and FEN1.

Accumulating evidence indicates that PARG also regulates DNA replication under exogenous genotoxic stress. In response to DNA damage, PAR chains are rapidly generated at DNA lesions to recruit DNA damage repair factors [8]. PARG subsequently degrades these PAR chains, facilitating the loading of repair factors and promoting the repair of DNA lesions in the subsequent stages [36]. Failure to degrade PAR chains can trap the DDR machinery at lesion sites, thereby blocking DNA repair [10, 37]. Under persistent replication stress, PARG‐deficient cells exhibit elevated γ‐H2AX expression, sustained CHK1‐S345 phosphorylation, S‐phase arrest, and increased replication fork collapse [38]. However, during unperturbed S phase, PARylation functions as a signaling mechanism to repair unligated Okazaki fragments, and the retention of PAR at replication sites must be precisely regulated [19]. PARG ensures the timely removal of PAR from replication forks, enabling complete and efficient replication. These observations highlight the importance of PAR metabolism in DNA replication, where the balance between PARP1 and PARG activities is crucial for efficient DNA replication. Consistent with recent findings that replication fork progression is impaired by the accumulation of post‐replicative ssDNA gaps in unperturbed PARG‐depleted cells [39], our study further demonstrates that the retention of PAR on PCNA due to PARG inhibition prevents the recruitment of FEN1, a key DNA replication factor, to replication sites. Beyond its effects on FEN1 recruitment, our data reveal that PARylated PCNA accumulation also significantly impairs its binding to DNA ligase I, the critical factor mediating nick sealing between Okazaki fragments (data not shown). These lead to an increase in unligated Okazaki fragments at replication forks, ultimately inhibiting DNA replication in unperturbed S‐phase cells. Our data elucidate the direct target and molecular mechanism by which PARG regulates DNA replication and provide insights into the embryonic lethality associated with PARG deletion.

In our study, we identified PCNA as a direct target of PARG in the regulation of DNA replication. During the S phase of normal unperturbed cells, PARG is recruited to DNA replication sites by PCNA, where it promptly removes PAR from PCNA. This dePARylation facilitates the interaction between PCNA and FEN1, thereby promoting efficient DNA replication. As a central coordinator of the DNA replication machinery, PCNA functions as a docking platform for proteins involved in DNA replication, DNA repair, cell cycle regulation, and cell survival. Post‐translational modifications of PCNA play key roles in DNA metabolism. For instance, mono‐ or poly‐ubiquitination of PCNA at the Lys164 residue promotes the restart of stalled replication forks, while SUMOylation at Lys164 inhibits PCNA ubiquitination. Additionally, SUMOylation at Lys127 may interfere with the binding of DNA polymerase to PCNA [40, 41]. However, the regulatory role of PARylation on PCNA remains poorly understood. Our findings reveal that PCNA is dynamically PARylated by PARP1 and dePARylated by PARG. PARylation of PCNA inhibits its binding with FEN1, highlighting a regulatory mechanism by which PARG‐mediated dePARylation facilitates DNA replication and expands the functional repertoire of PCNA post‐translational modifications.

PCNA plays an essential role in DNA metabolism and has emerged as a promising non‐oncogenic target for cancer therapy. Small molecules that disrupt PCNA‐protein interactions, particularly when combined with inhibitors targeting oncogenic drivers, may significantly yield synergistic anti‐tumor effects [42]. This strategy is further supported by the oncogenic properties of PARG, which is frequently overexpressed in multiple cancer types, including breast cancer, lung squamous cell carcinoma, and hepatocellular carcinoma, where elevated PARG expression correlates with poor clinical outcomes [43, 44, 45]. Importantly, PARG inhibitors have demonstrated potent antitumor activity both in vitro and in vivo [10, 13, 44]. Our study reveals an additional therapeutic dimension: PARG inhibitors not only suppress the oncogenic activity of PARG itself, but also indirectly impair PCNA's interaction with key replication factors like FEN1. We therefore propose that combining PARG inhibitors with other agents targeting DNA replication machinery may offer a powerful dual‐pronged approach for cancer therapy, particularly for malignancies dependent on efficient DNA replication and repair.

In conclusion, our study identifies PARG as a critical regulator of DNA replication. PARG‐mediated dePARylation of PCNA promotes its interaction with FEN1, and inhibition of PARG impairs this process, leading to suppressed DNA replication. These findings elucidate the essential role of PARG in DNA replication and provide mechanistic insights into how PARG regulates this fundamental cellular process.

Author Contributions

Zhenzhen Yan and Chen Wu conceived and designed the experiments; Zhenzhen Yan performed the experiments, analyzed the data, and made figures in the manuscript; Qianxi Feng, Zichang Qiao, Yaguang Wang, and Shuai Guo constructed all the vectors, performed the comet assay, and western blot; Xiuhuan Jiang, Xiaoru Han, and Xinying Cheng did the immunofluorescence staining; Xiaoyun Yang and Xiuhua Liu made structure modeling; Zhenzhen Yan, Chen Wu, and Hongyan Li wrote and revised the manuscript. All authors read and approved the final manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: fsb270959‐sup‐0001‐Figures.docx.

Table S1: fsb270959‐sup‐0002‐Tables.docx.

Yan Z., Feng Q., Jiang X., et al., “Poly (ADP‐Ribose) Glycohydrolase‐Dependent dePARylation of PCNA Is Essential for DNA Replication,” The FASEB Journal 39, no. 16 (2025): e70959, 10.1096/fj.202403378R.

Data Availability Statement

The data that support the findings of this study are available in the methods and results sections of this article.