L-arginine synergistic with 5-fluorouracil intervenes in DNA damage repair via the DNA-PKcs/ATM/ATR pathway in hepatocellular carcinoma cells

Abstract

DNA damage repair is a critical physiological process. The combined treatment of L-arginine (L-Arg) and 5-fluorouracil (5-FU) significantly inhibits cell proliferation, enhances nitric oxide (NO) production via inducible nitric oxide synthase (iNOS), and promotes the accumulation of reactive oxygen species (ROS). This heightened oxidative stress triggers DNA damage and apoptosis, as evidenced by a substantial increase in the Bax/Bcl-2 ratio; the activation of caspase-9, caspase-3, and PARP cleavage; and increased level of phosphorylated p53. Moreover, the combination treatment induces G2/M phase arrest, with a significant increase in p-H2AX (Ser 139) (known as γ-H2AX) expression, indicating extensive DNA damage. Mechanistically, the combined treatment modulates DNA damage response pathways by downregulating DNA-PKcs. Concurrently, it enhances the phosphorylation of ATM, ATR, CHK1, CHK2, and BRCA1. Additionally, the L-Arg and 5-FU combination downregulates PI3K/AKT signaling. AZD-7648 (a DNA-PKcs inhibitor) and LY294002 (a PI3K inhibitor) enhance p-ATM and p-ATR activation, resulting in elevated apoptosis and increased γ-H2AX expression. In contrast, the inhibition of ATM/ATR by CGK733 suppresses this response, reducing apoptosis and DNA damage signaling. Additionally, the ROS scavengers NAC and iNOS, when applied separately, restore p-AKT and DNA-PKcs expression; suppress the upregulation of p-ATM, p-ATR, and γ-H2AX; and ultimately reduce apoptosis. These findings are validated in a DEN-induced rat liver cancer model. In summary, 5-FU and L-Arg synergistically increase iNOS/NO-driven ROS accumulation, inducing γ-H2AX-marked DNA damage through dual modulation of repair pathways (inhibiting PI3K/AKT/DNA-PKcs while activating ATM/ATR), ultimately triggering p53-mediated G2/M arrest and apoptosis in hepatocellular carcinoma cells.

Introduction

Liver cancer is one of the most prevalent and lethal malignant tumors and is the second leading cause of cancer-related death worldwide [1]. Among primary hepatic tumors, hepatocellular carcinoma (HCC) accounts for approximately 90% of cases. Standard interventions for early-stage liver cancer include surgical resection, localized ablation procedures, and liver transplantation. However, many patients are diagnosed at an advanced stage of the disease. The principal treatment modalities available encompass immunotherapy, radiotherapy, and chemotherapy. First-line chemotherapeutic agents, including anti-metabolites such as 5-fluorouracil (5-FU), anti-angiogenic agents such as sorafenib, and programmed death inhibitors such as atezolizumab, primarily target deoxyribonucleic acid (DNA) and ribonucleic acid (RNA) [2].

Repairing DNA damage represents one of the body’s most common physiological processes, occurring predominantly through homologous recombination (HR) and non-homologous end joining (NHEJ) [3]. Mammalian cells have evolved sophisticated DNA repair mechanisms to address various forms of DNA damage—including mismatch repair, base excision repair, and nucleotide excision repair [4]. The key components involved in these processes include ataxia-telangiectasia mutated protein (ATM), ATM- and Rad3-related protein (ATR), and the catalytic subunit of DNA-dependent protein kinase (DNA-PKcs). These proteins are integral to the PI3K-associated kinase family and play crucial roles in the HR and NHEJ pathways, which are essential for effective DNA damage repair [5]. Inhibition of DNA-PKcs or activation of ATM/ATR via phosphorylation impedes cellular proliferation while influencing DNA damage correction mechanisms. Phosphorylation events involving these proteins constitute critical steps within the DNA damage response pathway (DDR), a comprehensive signaling framework that cells have developed to endure adverse conditions while ensuring accurate transmission of genetic information to subsequent generations [6]. When DNA-PKcs is inhibited, it may result in a blockade within the NHEJ pathway—one of the primary mechanisms by which cells manage double-strand breaks (DSBs) in their genomic material [7]. Additionally, ATM and ATR activation play critical roles in the homologous recombination (HR) pathway, initiating a cascade of complex phosphorylation events that can arrest the cell cycle at the G1/S or G2/M transitions [ 8, 9] . Consequently, the DNA-PKcs/ATM/ATR pathway represents a promising therapeutic target for cancer treatment. Upon DNA damage, the p53 protein undergoes phosphorylation, stabilizing it and promoting its nuclear accumulation. This activation induces the expression of genes associated with cell cycle arrest and apoptosis [10]. Simultaneously, DNA damage triggers the release of cytochrome c from mitochondria, activating caspase-9. Caspase-9 subsequently activates caspase-3, a key effector enzyme that drives apoptosis through the cleavage of various intracellular substrates, including PARP [11]. PARP cleavage inhibits DNA repair processes, committing the cell to irreversible apoptosis. These molecular events effectively eliminate cells with irreparable DNA damage, thereby preventing malignant transformation.

L-arginine (L-Arg), an essential basic amino acid, plays a crucial role in human physiology. Arginine metabolism primarily involves three pathways: nitric oxide (NO) synthesis, the urea cycle, and polyamine synthesis. Exogenous supplementation with L-Arg can increase NO levels via catalysis by intracellular inducible nitric oxide synthase (iNOS). iNOS expression within cells is not constitutive but rather induced in response to specific stimuli such as pro-inflammatory cytokines or bacterial endotoxins [12]. NO was the first gaseous molecule discovered to participate in cellular signaling transmission. Under the action of iNOS, L-Arg can produce high concentrations of NO, which, at concentrations ranging from 200 to 500 nM, has potent anti-tumor effects [13]. Elevated levels of NO can enhance the expression of p53, promote the release of cytochrome C from mitochondria, and induce protein nitrosylation and nitration. These processes lead to the formation of the cytotoxic agent peroxynitrite, which contributes to the antitumor properties of NO [14]. Although L-Arg does not directly influence nucleotide synthesis, it plays a critical role in DNA repair mechanisms within tumor cells [15]. Specifically, L-Arg modulates the transcription of genes involved in purine and pyrimidine synthesis and affects DNA repair via polyamine synthesis. Consequently, L-Arg supplementation, coupled with endogenous NOS-catalyzed NO production, suppresses neoplastic cell growth through multiple pathways, suggesting a novel therapeutic strategy for cancer treatment. In clinical practice, traditional chemotherapy drugs used for advanced HCC, such as 5-FU, doxorubicin, and mitomycin C (MMC), primarily target DNA damage [16] . However, monotherapy for liver cancer has shown limited efficacy and is associated with severe adverse effects. 5-FU increases iNOS expression in cancer cells, elevating NO level and inducing apoptosis [17]. Concurrent administration enhances the chemotherapeutic effect of 5-FU on BT-20 and MCF-7 breast cancer cells, particularly in p53-positive (wild-type) cells, where the effect is more pronounced [18]. When L-Arg level is sufficient, 5-FU induces iNOS expression in HCC cells ( e.g., BEL-7402 cells in vitro), significantly increasing intracellular NO level and promoting both apoptosis and necrosis in BEL-7402 cells, thereby increasing tumor cell sensitivity to 5-FU-based chemotherapy [19].

In our previous research, the combination of L-Arg and 5-FU inhibited key enzymes involved in aerobic glycolysis, including GLUT1, PKM2, and LDHA, suppressing glucose metabolism via the iNOS/NO/AKT pathway [20]. Cancer metabolism is characterized by elevated glycolysis and glucose uptake, which sustain the rapid proliferation of cancer cells. However, the combination of L-Arg and 5-FU inhibits glycolysis, leading to reduced ATP production and increased reactive oxygen species (ROS) generation. This disrupts the cellular redox balance, induces DNA damage and strand breaks, and ultimately triggers apoptosis in cancer cells [21]. The evidence indicates that glucose metabolism influences the efficacy of 5-FU treatment: glycolytic inhibition sensitizes lung cancer cells to 5-FU, whereas the Warburg effect in KRAS G12D glycolytic tumor organoids enhances 5-FU toxicity by altering nucleotide pools. Therefore, targeting glycolysis may improve the therapeutic efficacy of 5-FU in cancer treatment [22].

The current study focused on elucidating the molecular mechanisms by which L-Arg, in combination with 5-FU, interferes with DNA damage repair in liver cells via the DNA-PKcs/ATM/ATR pathway. Human hepatic carcinoma cell lines, including HepG2 and HuH-7, were used to assess markers associated with DNA damage, apoptosis, and the expression levels of proteins involved in the DNA damage response. Furthermore, a rat model of primary liver cancer induced by DEN was used to examine pathological changes in liver tissue and assess proteins related to DNA damage repair and apoptosis. This research investigated the role of L-Arg in conjunction with 5-FU in restoring DNA damage in liver cancer cells, providing novel theoretical insights for liver cancer therapy.

Materials and Methods

Reagents

An injectable formulation of arginine HCl was obtained from Xinyi Jinzhu Pharmaceutical Company (Shanghai, China). The 5-fluorouracil injectable solution was purchased from Xudong Haipu Pharmaceutical Company (Shanghai, China), while the 5-FU compound itself was sourced from Merck KGaA (Darmstadt, Germany). Diethylnitrosamine (DEN) was acquired from Yien Chemical Technology Company (Shanghai, China). HUVEC cell complete medium was from Pricella Biotechnology Company (Wuhan, China). Propidium iodide (PI) and a reactive oxygen species (ROS) detection kit were procured from Labgic Technology Company (Beijing, China). RNase reagent, hematoxylin-eosin (HE) staining kit, a streptavidin-biotin (SP) immunohistochemistry kit, and N-acetylcysteine (NAC) were obtained from Solarbio Science & Technology Company (Beijing, China). 4-Amino-5-methylamino-2′,7′-difluorofluorescein diacetate (DAF-FM DA), LY294002 and a comet assay kit were obtained from Beyotime Biotechnology Company (Shanghai, China). CGK733 was obtained from Ambeed Biotechnology Company (Shanghai, China), and AZD-7648 was obtained from Bide Pharmatech Ltd. (Shanghai, China). A one-step TUNEL in situ apoptosis detection kit (Red, Elab Fluor® 594), an Annexin V-FITC/PI dual staining kit, and a prestained protein marker (8–180 kDa) were purchased from KeyGEN Biotechnology Company (Nanjing, China). Color-coded prestained protein markers (8–200 kDa and 55–320 kDa) were acquired from Sevier Biotechnology Co., Ltd. (Wuhan, China). Rabbit anti-rat polyclonal antibodies for iNOS, caspase-3, caspase-9, PARP, DNA-PKcs, β-actin, histone H3, and Alexa Fluor 488 were obtained from Proteintech Biotechnology Company (Wuhan, China). Rabbit monoclonal antibodies against p-H2AX (Ser139) (γ-H2AX), p-BRCA1, p-ATM, p-CHK1, p-ATR, and p-CHK2, as well as a mouse monoclonal antibody for p-P53, along with horse anti-mouse and goat anti-rabbit secondary antibodies conjugated to HRP, were procured from Cell Signaling Technology (Danvers, USA). Both the iNOS shRNA plasmid and the negative control shRNA plasmid were acquired from Genechem Company (Shanghai, China). Additionally, the transfection reagent Lipofectamine 3000 was purchased from Thermo Fisher Scientific (Waltham, USA).

Cells and animals

HepG2, HuH-7, HeLa, and NIH3T3 cells were acquired from Servicebio Company (Wuhan, China). AML12 cells were acquired from Zhong Qiao Xin Zhou Biotechnology Company (Shanghai, China). KYSE-150, A549, and HUVECs cells were acquired from Wuhan Pricella Biotechnology Company (Wuhan, China). The cells cultured under conditions of 95% humidity and 5% CO ₂ and maintained at 37°C. The HUVECs cells were cultured in HUVEC cell complete medium containing 10% fetal bovine serum. Other cells were grown in DMEM supplemented with 10% fetal bovine serum. Upon reaching approximately 80%–85% confluency, the cells were trypsinized using a solution containing 0.25% EDTA for passaging or further experimental procedures. SPF-grade SD rats weighing 180–220 g and aged 6 weeks were obtained from Liaoning Changsheng Biotechnology Company (Benxi, China). The animals were housed in a controlled environment with a relative humidity of 40%–70% and a temperature range of 22°C–26°C and were provided ad libitum access to food and water. Following a one-week adaptation period, the rats were subjected to subsequent experiments. The animal study was reviewed and approved by Medical Ethics Committee of the First Affiliated Hospital of Henan University of Science and Technology (2024-0307). All animal handling and experimental procedures strictly adhered to institutional guidelines.

MTT assay

For MTT evaluation, HepG2 and HuH-7 cells were routinely trypsinized, and the cell suspension density was adjusted to 1.0 × 10 ⁵ cells/mL. These cells were then plated and treated with combined doses of L-Arg at concentrations of 5, 10, and 20 mM, along with 5-FU at concentrations of 0.05, 0.1, and 0.2 mM. At 44 h and 68 h post-treatment, 20 μL of preprepared MTT solution (Servicebio, Wuhan, China) was added to each well, followed by incubation for 4 h. The supernatant was subsequently discarded, and DMSO was added to dissolve the formazan crystals. The plate was gently shaken for 10 min in the dark. The absorbance (OD) was measured at 490 nm via a microplate reader. Cell viability (%) was calculated via the following formula: Cell viability (%) = OD (drug-treated)/OD (control) × 100%. The combination index (CI) for the combined treatment was calculated via CalcuSyn software on the basis of the degree of cell inhibition. A CI value greater than 1 indicates antagonism, a CI value less than 1 indicates synergy, and a CI value equal to 1 represents an additive effect. On the basis of the optimal concentrations determined in the preliminary experiments, additional studies were subsequently performed to evaluate the effects of these concentrations on three additional tumor cell lines (KYSE-150, A549, and HeLa) as well as normal cells (HUVECs, AML12, and NIH3T3), employing the same experimental protocols described earlier.

Colony formation determination

HepG2 and HuH-7 cells were seeded at a density of 500 cells per dish. After the cells adhered and spread, they were divided into four treatment groups: control, L-Arg (20 mM), 5-FU (0.2 mM), and a combination of L-Arg and 5-FU. Following a 48-h incubation period, the culture medium was replaced by fresh medium. Once distinct colonies became visible, the old medium was removed, and the cells were fixed in 4% paraformaldehyde for 20 min after rinsing with PBS. The cells were subsequently stained with a 0.5% crystal violet solution for 10 min, followed by a PBS rinse to eliminate excess stain. The colonies were air-dried and imaged. The colony-forming efficiency was quantified via ImageJ software according to the following formula: colony-forming efficiency (%) = (number of colonies/number of cells seeded) × 100%.

NO fluorescence probe assay

HepG2 and HuH-7 cells were seeded in culture plates at a density of 5 × 10 ⁶ cells per milliliter. Following cell adhesion, the designated drugs were administered to the cells for 48 h. Subsequently, the cells were incubated with a diluted NO fluorescent probe (DAF-FM DA, 5 μM) at 37°C in the dark for 30 min. Finally, the fluorescence intensity was measured by flow cytometry for each group to evaluate the intracellular NO level. The excitation and emission wavelengths were set at 495 nm and 515 nm, respectively.

DAPI staining assay

HepG2 and HuH-7 cells were seeded at a density of 1.0 × 10 ⁵ cells per milliliter. The designated drugs were subsequently added to each group, and the cultures were maintained for 48 h. The cells were then fixed with 4% paraformaldehyde and permeabilized with 0.5% Triton X-100 for 20 min. DAPI staining solution (100 μL per well) was added, followed by incubation in the dark for 10 min. Finally, the cells were washed and imaged under a fluorescence microscope.

Annexin V-FITC/PI dual-staining detection

HepG2 and HuH-7 cells were maintained at a concentration of 1 × 10 ⁵ cells per milliliter in culture dishes. Once the cell density reached 80%, the cells were treated with L-Arg (20 mM) or 5-FU (0.2 mM) either individually or in combination at the indicated concentration (L-Arg: 20 mM + 5-FU: 0.2 mM). The cells were harvested 48 h after drug treatment. These cells were rinsed with PBS and then resuspended in binding solution. Then, Annexin V conjugated with FITC staining solution was introduced and mixed, and propidium iodide was subsequently added. The mixture was kept in the dark for 10 min. Finally, flow cytometry was used to measure the percentage of apoptotic cells.

PI staining for cell cycle analysis

HepG2 and HuH-7 cells were treated with L-Arg (20 mM) or 5-FU (0.2 mM), either individually or in combination, at the indicated concentrations (L-Arg: 20 mM + 5-FU: 0.2 mM). The cells were harvested 48 h post-treatment. The cell suspension was gently mixed with pre-chilled 70% ethanol and fixed on ice for 1 h. After centrifugation, the cells were washed with 250 μL of cold PBS at 4°C. RNase solution was subsequently added, and the cells were incubated at 37°C for 30 min. PI staining solution was subsequently added, and the cells were further incubated at 37°C for 30 min. Finally, flow cytometry analysis was performed to assess the degree of cell cycle arrest in each group.

TUNEL staining assay

Both HepG2 and HuH-7 cells were cultured at a density of 1×10 ⁵ cells per dish. The cells were treated with L-Arg (20 mM) or 5-FU (0.2 mM), either individually or in combination, at the specified concentrations (L-Arg: 20 mM + 5-FU: 0.2 mM). Following a 48-h exposure to the indicated treatments, the cells were fixed with 4% paraformaldehyde. Subsequently, cell permeabilization was achieved by incubating the dishes with 0.5% Triton X-100 solution for 20 min. A TdT enzyme reaction mixture was then prepared and added to each dish at a volume of 50 μL, followed by incubation in the dark at 37°C for 1 h. Next, a streptavidin-TRITC labeling solution was prepared at a ratio of streptavidin-TRITC:labeling buffer=1:9 and applied to each dish for a 30-min incubation in the dark at 37°C. Afterward, the dishes were washed multiple times with PBS, and the nuclei were stained with DAPI, followed by an additional wash. Finally, anti-fade mounting medium was applied prior to imaging via a laser confocal microscope.

Immunofluorescence assay

Both HepG2 and HuH-7 cells were seeded at a density of 1 × 10 ⁵ cells per dish. Following a 48-h drug treatment, the cells were fixed with 4% paraformaldehyde for 20 min. Next, permeabilization was performed via incubation with 0.5% Triton X-100 for 20 min, followed by washing with PBS. The cells were subsequently blocked with 10% goat serum for 30 min to reduce non-specific binding. The primary antibody, diluted in 1% BSA, was then added, and the dishes were incubated overnight at 4°C. After the samples were washed with PBS, an Alexa Fluor 488-conjugated secondary antibody was applied, and the samples were incubated in the dark. The cells were then stained with DAPI for 10 min before being imaged via a confocal fluorescence microscope.

Comet assay

HepG2 and HuH-7 cells were seeded at a concentration of 1×10 ⁵ cells per well. Following exposure to the respective drugs, the cells from each group were harvested. Pre-melted 1% normal melting point agarose gel was applied at 30 μL per well onto pre-warmed comet assay slides maintained at 45°C, immediately covered with a coverslip, and solidified in a refrigerator set to 4°C for 10 min. Once the gel solidified, the coverslips were carefully removed, and 0.7% low melting point agarose gel containing the previously collected cells from each group was quickly layered over the first gel layer and refrigerated for an additional 10 min. The slides were subsequently immersed in lysis buffer (lysis buffer:DMSO = 9:1) at 4°C. After lysis, the slides were transferred to pre-prepared alkaline electrophoresis buffer. Electrophoresis was then performed in the dark at low voltage (25 V) in an ice bath. Following electrophoresis, the slides were neutralized by immersion in Tris-HCl buffer (0.4 M), pH 7.5, at 4°C for 5 min per cycle. Next, the PI staining solution was added to the gel, which was subsequently incubated in darkness for 20 min, after which it was washed with ultrapure water prior to being sealed with a coverslip and imaged under a laser scanning confocal microscope. The extent of DNA damage in the cells was quantified via CASP software on the basis of the percentage of tail DNA (% tail DNA). The degree of DNA damage was classified as follows: less than 5% tail DNA indicated no observable damage; 5%–20% indicated minor damage; 20%–40% indicated moderate damage; 40%–95% indicated significant damage; and greater than 95% indicated severe damage.

ROS detection assay

HepG2 and HuH-7 cells were seeded at a density of 1×10 ⁵ cells/mL. The cells were subsequently treated with NAC (3 mM) or L-Arg+5-FU (L-Arg: 20 mM, 5-FU: 0.2 mM), either individually or in combination. After 48 h of exposure to the indicated treatments, the treatment medium was removed, and an appropriate volume of diluted H2DCFDA working solution was added to fully cover the cells. The cells were incubated in a 37°C cell culture incubator in the dark for 30 min. Following incubation, the cells were washed 1–2 times with serum-free culture medium to thoroughly remove unincorporated H2DCFDA. Finally, the cells were analyzed via a fluorescence microscope or examined via flow cytometry.

shRNA transient transfer assay

The cells were seeded into 6-well plates. A transfection mixture was prepared by diluting Lipofectamine 3000 reagent in high-glucose DMEM without serum or antibiotics, along with plasmids encoding either iNOS-targeting shRNA or control shRNA. The sense sequence of the iNOS shRNA was 5′-CCAGAAGCAGAATGTGACCAT-3′, while the sense sequence for the negative control shRNA was 5′-TTCTCCGAACGTGTCACGT-3′. This mixture was incubated at room temperature for 20 min before being added to the appropriate wells for transfection. The cells were subsequently incubated in a CO ₂ incubator for 8 h, after which the medium was replaced by complete medium supplemented with 10% fetal bovine serum (FBS). The cells were then cultured for an additional 24 h prior to treatment with the respective drugs for 48 h.

Establishment of a primary HCC model in rats

Sixty clean-grade SD rats were randomly divided into five groups, each consisting of twelve rats. The groups were as follows: normal control group (intraperitoneal injection of 0.9% saline, 10 mL·kg ^–1), model group (intraperitoneal injection of DEN, 50 mg·kg ^–1), L-Arg group (intraperitoneal injection of L-Arg, 700 mg·kg ^–1), 5-FU group (intraperitoneal injection of 5-FU, 20 mg·kg ^–1), and combination group (intraperitoneal injection of both L-Arg and 5-FU, 700 mg·kg ^–1 +20 mg·kg ^–1). Except the normal control group, all the other groups received intraperitoneal injections of DEN twice weekly during the first four weeks, followed by once weekly for the subsequent 16 weeks. Starting from the fifth week, after each weekly DEN injection, the corresponding drugs were administered intraperitoneally for 16 consecutive weeks. At the end of the experiment, the rats were anesthetized with intraperitoneal sodium pentobarbital, and their liver tissues were harvested for weighing and assessment of nodule formation. A portion of the liver tissue was washed with 0.9% saline and fixed in 4% paraformaldehyde for the preparation of paraffin-embedded and frozen sections. Another portion of each fresh liver sample was stored at -80°C for subsequent immunoblot analysis.

Hematoxylin and eosin (H&E) staining

Liver specimens, measuring approximately 1 cm × 1 cm × 0.5 cm, were fixed in 4% paraformaldehyde, dehydrated through a graded alcohol series, and embedded in paraffin. Sections were cut at a thickness of 5 μm. Five slides from each group were baked, deparaffinized, and stained with hematoxylin. Excess stain was removed by rinsing with running water before the slides were differentiated in hydrochloric acid alcohol. The slides were subsequently blued with running water and stained with eosin. After excess stain was removed, the slides were dried and prepared for microscopic examination.

Immunohistochemical staining

The paraffin-embedded sections were deparaffinized and subjected to antigen retrieval via sodium citrate buffer (0.01 mol·L ^–1) for 10 min. This process involved boiling in a microwave on medium heat for 5 min, pausing, reheating to boiling after 10 min, and then cooling to 57°C, followed by two washes with PBS. Endogenous peroxidase activity was blocked with 3% hydrogen peroxide, followed by rinsing with distilled water. Goat serum blocking solution was applied for 20 min without washing. Primary antibodies were added, and the samples were incubated overnight at 4°C. A biotin-labeled goat anti-rabbit working solution was applied, and the samples were incubated at room temperature. Streptavidin-HRP reagent was subsequently added, followed by a 30-min incubation at room temperature. After a final PBS rinse, the sections were treated with DAB solution in the dark, and the reaction was stopped with distilled water. The cell nuclei were counterstained with hematoxylin, followed by dehydration, clearing, and mounting. The samples were examined and imaged under a microscope. Brownish-yellow or brown granules indicated positive expression, which was analyzed semi-quantitatively using Image-Pro Plus 6.0 software.

Immunoblot analysis

The cells or tissues were collected in centrifuge tubes, and an appropriate volume of Western and IP cell lysis buffer containing PMSF or nuclear protein extraction buffer was added to extract total and nuclear proteins. DNA-PKcs, p-ATM, p-ATR, and p-BRCA1 were separated via 4% or 6% separation gels, whereas the other proteins were separated via 12% or 15% separation gels. All the above proteins were concentrated via conventional concentration gel systems. The concentration of the separation gel was adjusted on the basis of the actual molecular weight of the protein, and the separation gel at the location of the target protein was excised according to the electrophoresis results for subsequent experiments. The proteins were subsequently separated by SDS-PAGE at the appropriate concentration and transferred onto a PVDF membrane. The membrane was blocked with 5% skim milk at room temperature for 1 h, followed by incubation with the primary antibody overnight at 4°C. The next day, after washing with TBST, the membrane was incubated with the corresponding secondary antibody at room temperature for 1 h. After another TBST wash, the proteins were visualized via the enhanced chemiluminescence (ECL) method.

Statistical analysis

All experimental data are expressed as the mean ± standard deviation (SD) and were tested for a normal distribution. Data that conformed to a normal distribution were analyzed via one-way analysis of variance (ANOVA). Statistical analyses were performed via GraphPad Prism 10 software, and Dunnett’s t test is typically employed for comparing multiple groups against a single control group. A P value less than 0.05 was considered statistically significant, whereas a P value less than 0.01 was considered highly significant.

Results

Impact of combined L-Arg and 5-FU on cell viability in tumor and normal cells

MTT assay was used to evaluate the effects of the combination of L-Arg and 5-FU on HepG2 and HuH-7 cell survival rates at 48 h and 72 h. As illustrated in Figure 1A, treatment with L-Arg and 5-FU at various concentrations significantly decreased cell survival rates. Moreover, as both the concentration and exposure time increased, the survival rates progressively decreased. In HepG2 cells, co-treatment with L-Arg at 20 mM and 5-FU at 0.2 mM for 48 h resulted in the lowest CI value of 0.056, indicating the strongest synergistic effect ( Figure 1B). Similarly, in HuH-7 cells, the same concentrations of L-Arg and 5-FU for 48 h produced the lowest CI value of 0.027, demonstrating the optimal synergistic interaction ( Figure 1B). Therefore, subsequent experiments were performed under the following conditions: L-Arg at 20 mM and 5-FU at 0.2 mM for 48 h. Additionally, the number of colony-forming units in the group treated with the combination of L-Arg and 5-FU was markedly reduced ( Figure 1C). At this predetermined concentration, L-Arg or 5-FU alone significantly inhibited the proliferation of three tumor cell lines (KYSE-150, A549, and HeLa). However, compared with single agent treatment, combined treatment with L-Arg and 5-FU had a more pronounced inhibitory effect on HepG2 and HuH-7 cells ( Figure 1D). Conversely, at the same dosage, the inhibitory effects on three non-tumor cell lines (HUVECs, AML12 cells, and NIH3T3 cells) were minimal ( Figure 1E). These results demonstrated that the synergistic combination of L-Arg and 5-FU selectively enhanced the anti-proliferative effects on tumor cells, particularly HepG2 and HuH-7 cells, while exhibiting minimal cytotoxicity to non-tumor cell lines.

Figure 1 .

Co-administration of L-Arg and 5-FU influences the proliferation of both tumor and normal cells

(A) The combined effects of L-Arg and 5-FU on the viability of HepG2 and HuH-7 cells were assessed at 48 h and 72 h via MTT assay. (B) The combination index (CI) for L-Arg and 5-FU was analyzed via CalcuSyn software on the basis of the MTT assay data. (C) The impact of L-Arg and 5-FU on colony-forming ability was evaluated through colony formation assay. (D) Effects of L-Arg and 5-FU on the viability of KYSE150, A549, and HeLa tumor cells were assessed at 48 h via MTT assay. (E) The effects of L-Arg and 5-FU on the viability of normal HUVECs, AML12 cells, and NIH3T3 cells were assessed at 48 h via MTT assay. Data are presented as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. *P < 0.05, **P < 0.01, compared with the control group; # P < 0.05, ## P < 0.01, compared L-Arg with 5-FU group.

Impact of combined L-Arg and 5-FU on NO levels and iNOS expression in HepG2 and HuH-7 cells

NO levels were quantified via DAF-FM DA fluorescent probes. Compared with no treatment, treatment with L-Arg alone or in combination with 5-FU significantly increased NO levels ( Figure 2A). Notably, co-administration of L-Arg and 5-FU resulted in a significantly greater increase in NO levels than did either L-Arg or 5-FU treatment alone, suggesting a synergistic effect between L-Arg and 5-FU in enhancing intracellular NO production. As shown in Figure 2B, iNOS expression was significantly upregulated in both the 5-FU-treated group and the L-Arg+5-FU combination group compared with the untreated control group. Moreover, iNOS expression was markedly greater in the L-Arg+5-FU group than in the L-Arg-only group, indicating that the combination of L-Arg and 5-FU has a more pronounced effect on iNOS expression than does L-Arg alone. These results demonstrated that both 5-FU alone and in combination with L-Arg can effectively increase iNOS expression in HepG2 and HuH-7 cells.

Figure 2 .

Co-administration of L-Arg and 5-FU affectes NO levels and iNOS levels in HepG2 and HuH-7 cells

(A) The influence of L-Arg and 5-FU on NO levels was quantified via DAF-FM DA fluorescent probes. (B) The impact of L-Arg and 5-FU on iNOS protein levels was evaluated by immunoblot analysis. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. *P < 0.05, **P < 0.01, compared with the control group; ## P < 0.01, compared L-Arg with 5-FU group.

Impact of combined L-Arg and 5-FU on apoptosis in HepG2 and HuH-7 cells

DAPI staining was used to visualize the morphological changes in the nucleus. The nuclei of the cells in the control group exhibited typical oval or nearly round shapes ( Figure 3A). In contrast, cells treated with L-Arg, 5-FU, or their combination presented characteristic apoptotic features, including nuclear fragmentation, crescent-shaped nuclei, nuclear condensation, and high chromatin condensation. These apoptotic features were most prominent in the L-Arg + 5-FU group, suggesting a synergistic effect of the two agents in inducing apoptosis. Apoptosis was further evaluated via Annexin V/FITC-PI staining. The apoptosis rate was significantly greater in the L-Arg + 5-FU group than in the other treatment groups ( Figure 3B). Immunoblotting analysis was conducted to evaluate the levels of proteins associated with apoptosis in HepG2 and HuH-7 cells. The results demonstrated that the levels of Bax were significantly increased, whereas those of Bcl-2 were markedly decreased, leading to a substantial increase in the Bax/Bcl-2 ratio in the co-administration group ( Figure 3C,D). Additionally, the cleavage of caspase-9, caspase-3, and PARP, as well as the phosphorylation of p53, was significantly increased in the combined L-Arg and 5-FU group ( Figure 3E,F). These findings suggested that co-administration treatment effectively activated apoptosis-related proteins in HepG2 and HuH-7 cells.

Figure 3 .

Co-administration of L-Arg and 5-FU modulates apoptosis and the expression levels of apoptosis-related proteins in HepG2 and HuH-7 cells

(A) The apoptotic morphological changes induced by combined L-Arg and 5-FU treatment were evaluated via DAPI staining. (B) The rate of apoptosis resulting from the combination of L-Arg and 5-FU was quantified via Annexin V/FITC-PI staining. (C,D) The effects of L-Arg and 5-FU on the protein expression levels of Bcl-2 and Bax were assessed. (E,F) Immunoblot analysis revealed the effects of L-Arg and 5-FU on the phosphorylation of p53, as well as the cleavage of caspase-9, caspase-3, and PARP. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. *P < 0.05, **P < 0.01, compared with the control group; # P < 0.05, ## P < 0.01, compared L-Arg with 5-FU group.

Impact of combined L-Arg and 5-FU on DNA content and DNA damage repair in HepG2 and HuH-7 cells

Propidium iodide (PI) staining was employed to quantify the DNA content. As shown in Figure 4A, compared with that in the control group, the proportion of cells in the G2/M phase was markedly greater, whereas the proportion in the G0/G1 phase was significantly lower in the L-Arg plus 5-FU group. These results suggest that the coadministration of L-Arg and 5-FU effectively induces apoptosis and arrests cells in the G2/M phase. The comet assay was utilized to evaluate the effects of coadministration on DNA damage. Compared with the control group, moderate DNA damage was observed in both the L-Arg group and the 5-FU group, as indicated by a comet tail DNA percentage ranging from 5% to 20%. In contrast, severe DNA damage was evident in the coadministration treatment group, with a comet tail DNA percentage ranging from 40% to 95% ( Figure 4B). These findings indicate that the combined use of L-Arg and 5-FU can cause substantial DNA damage in cells. The impact of combined treatment on DNA fragmentation in HepG2 and HuH-7 cells was assessed via tetramethylrhodamine (TRITC) labeling. Red fluorescence, indicative of DNA breaks, was significantly increased in the coadministration group ( Figure 4C). Moreover, the levels of γ-H2AX were significantly elevated in the combined L-Arg and 5-FU group ( Figure 4D,E), suggesting that the combined treatment induced significant DNA fragmentation and apoptosis.

Figure 4 .

Co-administration of L-Arg and 5-FU modulates DNA damage and the cell cycle distribution in HepG2 and HuH-7 cells

(A) Cell cycle distribution following treatment with L-Arg and 5-FU was analyzed via PI staining. (B) DNA migration after combined treatment with L-Arg and 5-FU was quantified via comet assay. (C) DNA fragmentation induced by the combination of L-Arg and 5-FU was assessed via TUNEL assay. (D) The impact of combined L-Arg and 5-FU treatment on γ-H2AX levels was evaluated via immunofluorescence analysis. (E) Immunoblot analysis further confirmed the effects of L-Arg and 5-FU on γ-H2AX expression levels. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. *P < 0.05, **P < 0.01, compared with the control group; ## P < 0.01, compared L-Arg with 5-FU group.

Impact of the coadministration of L-Arg and 5-FU on the regulation of DNA damage repair-associated proteins in HepG2 and HuH-7 cells

Immunofluorescence and immunoblot analyses were conducted to examine the effects of combined treatment on DNA-PKcs expression. The level of DNA-PKcs was reduced in the combination group ( Figure 5A). In contrast, the expression levels of phosphorylated ATM, ATR, CHK1, CHK2, and BRCA1 were significantly increased ( Figure 5B). Moreover, AZD-7648, a DNA-PKcs inhibitor, markedly potentiated the upregulation of p-ATM, p-ATR, γ-H2AX, and cleaved-caspase-3 proteins induced by the combination of L-Arg and 5-FU, thereby increasing apoptosis in HCC cells ( Figure 5C). In contrast, CGK733, an ATM/ATR inhibitor, substantially abrogated the upregulation of the p-ATM, p-ATR, and cleaved-caspase3 proteins as well as the reduction in γ-H2AX expression triggered by the combination of L-Arg and 5-FU, resulting in reduced apoptosis in HCC cells ( Figure 5D). However, it had a minimal effect on DNA-PKcs. This selective action underscores the crucial role of ATM/ATR in mediating the apoptotic effects induced by the combination of L-Arg and 5-FU.

Figure 5 .

Co-administration of L-Arg and 5-FU modulates the regulation of DDR-related proteins in HepG2 and HuH-7 cells

(A) The impact of combined L-Arg and 5-FU treatment on DNA-PKcs levels was assessed via immunofluorescence assays. (B) Immunoblot analysis was performed to evaluate the effects of L-Arg and 5-FU on the expression levels of DNA-PKcs, p-ATM, p-ATR, p-CHK1, p-CHK2, and p-BRCA1. (C) The influence of the DNA-PKcs inhibitor AZD7648 on the protein levels of DNA-PKcs, p-ATM, p-ATR, γ-H2AX, and cleaved caspase-3 was investigated. The cells were pre-treated with AZD7648 (10 μM) for 1 h before subsequent treatments. (D) The effects of the ATM/ATR inhibitor CGK733 on the protein levels of DNA-PKcs, p-ATM, p-ATR, γ-H2AX, and cleaved caspase-3 were examined. Cells were pre-incubated with CGK733 (10 μM) for 1 h prior to further treatments. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. *P < 0.05, **P < 0.01, compared with the control group; ## P < 0.01, compared L-Arg with 5-FU group.

Impact of co-administration of L-Arg and 5-FU on the regulation of PI3K/AKT pathway components in HepG2 and HuH-7 cells

Immunoblot analysis was employed to evaluate the effects of combined treatment with L-Arg and 5-FU on the PI3K/AKT signaling pathway. Compared with those in the control group, the levels of p-PI3K and p-AKT were significantly lower in the combination group ( Figure 6A,B). Treatment with LY2940029 (a PI3K inhibitor) further reduced DNA-PKcs expression in the combination group but increased the expressions of p-ATM and p-ATR ( Figure 6C,D). These findings suggested that the combined treatment concurrently regulated DNA damage repair through the DNA-PKcs/ATM/ATR pathway and significantly suppressed the expressions of proteins associated with the PI3K/AKT signaling pathway.

Figure 6 .

Co-administration of L-Arg and 5-FU modulates PI3K/AKT pathway components in HepG2 and HuH-7 cells

(A,B) The effects of combined L-Arg and 5-FU treatment on the levels of phosphorylated PI3K (p-PI3K) and phosphorylated AKT (p-AKT) were evaluated by immunoblot analysis. (C,D) The influence of LY294002 on the expression levels of key components of the PI3K/AKT pathway in the L-Arg and 5-FU treatment groups was assessed. Cells were pre-treated with LY294002 (10 μM) for 1 h before being exposed to L-Arg and 5-FU. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. **P < 0.01, compared with the control group; ## P < 0.01, compared L-Arg with 5-FU group.

Effects of ROS on DNA damage repair and apoptosis in L-Arg- and 5-FU-treated HepG2 and HuH-7 cells

The intracellular ROS levels were quantitatively detected via the fluorescent probe 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA). The results demonstrated that the combination of L-Arg and 5-FU significantly increased ROS levels, whereas NAC, an ROS inhibitor, mitigated the increase in ROS induced by the combination therapy ( Figure 7A,B). Upon intervention with NAC, the expression of p-AKT and DNA-PKcs induced by the combination therapy was elevated, whereas the levels of p-ATM and p-ATR were reduced ( Figure 7C). Consequently, γ-H2AX expression decreased, leading to a reduction in cleaved caspase-3 expression. These results showed that L-Arg and 5-FU increased ROS levels, driving AKT inhibition, DNA-PKcs suppression, ATM/ATR activation, DNA damage, and apoptosis.

Figure 7 .

The role of reactive oxygen species (ROS) in DNA damage and apoptosis induction during the coadministration of L-Arg and 5-FU

(A) Flow cytometric analysis of ROS changes under combined treatment with L-Arg and 5-FU. (B) Fluorescence microscopy-based evaluation of ROS modulation induced by the coadministration of L-Arg and 5-FU. The cells were preincubated with carboxy-PTIO, an NO scavenger (50 μM), for 1 h before subsequent treatments. (C) Effects of the ROS inhibitor N-acetylcysteine (NAC) on the protein expression levels of p-AKT, DNA-PKcs, p-ATM, p-ATR, γ-H2AX, and cleaved caspase-3. The cells were pre-treated with NAC (3 mM) for 1 h prior to subsequent treatments. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. *P < 0.05, **P < 0.01, compared with the control group; # P < 0.05, ## P < 0.01, compared L-Arg with 5-FU group.

Impact of iNOS on apoptotic and DNA damage repair-related protein expression in HepG2 and HuH-7 cells

Immunoblot analysis was performed after transient transfection with iNOS shRNA plasmids to assess protein expression levels. iNOS expression was significantly decreased in the co-administration group transfected with iNOS shRNA than in the negative control combination group ( Figure 8A,B). Furthermore, the expression of Bcl-2 and DNA-PKcs markedly increased in the iNOS shRNA combination group, whereas the expressions of phosphorylated ATM, ATR, p53, and cleaved caspase-3 significantly decreased. These results indicated that coadministration suppressed the DNA damage repair process and promoted apoptosis, which was closely associated with elevated iNOS expression.

Figure 8 .

iNOS affects the levels of apoptosis-related and DNA damage repair-related proteins in HepG2 and HuH-7 cells

(A) HepG2 cells. (B) HuH-7 cells. Data are expressed as the mean ± standard deviation, with n = 3. The normal distribution of the data was verified prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA), and Dunnett’s t test was used for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. **P < 0.01, compared with the control group; # P < 0.05, ## P < 0.01, compared L-Arg with 5-FU group.

Impact of L-Arg and 5-FU on pathological modifications and the expressions of apoptotic proteins in rat liver tissue

In the evaluation of rat liver tissue via HE staining, the effects on primary liver cancer pathology were observed. In the normal control group, the central vein was clearly visible and surrounded by radially arranged hepatic cords composed of hepatocytes of uniform size and rich cytoplasm ( Figure 9A). Conversely, in the DEN-induced model group, the liver lobules exhibited an irregular structure, with the liver cells clustering together. Additionally, variations in liver cell size were noted, along with nuclear division phenomena such as enlarged, multinucleated, and multipolar nuclei. The nuclei of the liver cells were large and deeply stained blue, indicating significant changes in cellular morphology. Necrosis and infiltration of inflammatory cells were evident in the hepatic tissue, whereas a decrease in nuclear fission was observed in the L-Arg and/or 5-FU groups. These findings suggest that the combined treatment can effectively mitigate the pathological changes in liver tissue induced by DEN in a primary liver cancer model in rats.

Figure 9 .

Co-administration of L-Arg and 5-FU induces pathological changes and modulates the protein levels of iNOS, cleaved caspase-3, and cleaved PARP in rat liver tissue

(A) Pathological alterations in rat liver tissue were assessed in the L-Arg plus 5-FU group via hematoxylin and eosin (HE) staining. (B) The modulation of iNOS levels in rat liver tissue in the L-Arg plus 5-FU group was evaluated. (C) The effect of L-Arg plus 5-FU on cleaved caspase-3 levels in rat liver tissue was analyzed. (D) The influence of L-Arg plus 5-FU on cleaved PARP levels in rat liver tissue was examined. Data are expressed as the mean ± standard deviation, with n = 12. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software. **P < 0.01, compared with the normal control group; ΔΔ P < 0.01, compared with the DEN model group. ## P < 0.01, compared L-Arg with 5-FU group.

Immunohistochemical staining was employed to examine the impact of L-Arg and 5-FU on the expressions of iNOS, cleavage of caspase-3, and PARP in rat liver tissue. Compared with the model group, the co-administered group presented significantly increased iNOS expression ( Figure 9B–D). Similarly, the cleavage of both caspase-3 and PARP was markedly increased in the coadministration group. These results confirmed that the combined treatment significantly elevated iNOS expression and enhanced the cleavage of caspase-3 and PARP in rat liver tissue, thereby promoting apoptosis.

Impact of combined L-Arg and 5-FU on the expression of DNA damage repair proteins, PI3K/AKT pathway proteins and apoptosis-related proteins in rat liver tissue

Both immunofluorescence and immunoblot analyses were performed to assess the levels of DNA damage repair proteins, PI3K/AKT pathway proteins, and apoptosis-associated proteins in rat liver tissue. The results demonstrated that the levels of phosphorylated AKT, DNA-PKcs, and Bcl-2 were significantly reduced, whereas the expressions of phosphorylated ATM, ATR, and p53, as well as the cleavage of caspase-9 and caspase-3, were markedly increased in the co-administration group ( Figure 10A–C). These findings suggested that the combination of L-Arg and 5-FU enhanced iNOS activity and suppressed the DNA-PKcs repair pathway, which subsequently activated the ATM/ATR signaling cascade, ultimately promoting apoptosis in HCC cells.

Figure 10 .

Co-administration of L-Arg and 5-FU modulates the expression levels of proteins associated with DNA damage repair, the PI3K/AKT signaling pathway, and apoptosis in rat liver tissue

(A) Immunofluorescence staining was used to analyze the effect of combined L-Arg and 5-FU treatment on DNA-PKcs levels. (B) Immunoblot analysis was used to evaluate effects of co-administration of L-Arg and 5-FU on the levels of DNA-PKcs, phosphorylated ATM (p-ATM), phosphorylated ATR (p-ATR), and phosphorylated AKT (p-AKT) in rat liver tissue. (C) The influence of coadministration of L-Arg and 5-FU on the levels of phosphorylated p53 (p-p53), Bcl-2, cleaved caspase-3, and cleaved caspase-9 in rat liver tissue was assessed via immunoblot analysis. Data are expressed as the mean ± standard deviation, with n = 12. The normal distribution of the data was confirmed prior to analysis. Statistical evaluations were performed via one-way analysis of variance (ANOVA) followed by Dunnett’s t test for pairwise comparisons between groups. Analyses were conducted via GraphPad Prism 10 software.**P < 0.01, compared with the normal control group; ΔΔ P<0.01, compared with the DEN model group. ## P < 0.01, compared L-Arg with 5-FU group.

Discussion

In terms of cellular biology, unrepaired DNA DSBs can lead to genomic instability, potentially reducing cell viability. Certain chemotherapeutic agents induce DNA damage [23]. For instance, 5-FU, a commonly used antimetabolite in tumor therapy, exerts its antitumor effects by damaging genetic material (DNA/RNA) and disrupting tumor cell function. iNOS, which is expressed in various tumor cells, catalyzes the production of NO from L-Arg [24]. Elevated NO levels indicate antitumor activity. In this study, the combined treatment of L-Arg and 5-FU had significant inhibitory effects on other tumor cell lines, with the most pronounced effect on HepG2 and HuH-7 liver cancer cells. At equivalent concentrations, minimal effects were observed on normal cells. This disparity may arise from potential off-target effects or the specificity of the drug’s mechanism of action. The heightened sensitivity of liver cancer cells could stem from their distinct metabolic or molecular features, increasing their vulnerability to drugs [25]. Conversely, the limited effect on normal cells indicates a degree of selectivity, which is beneficial for minimizing adverse effects.

Concurrently, 5-FU can increase the expression of iNOS in HepG2 and HuH-7 cells. When combined with L-Arg, 5-FU significantly increased NO production in these cells. Apoptosis suppresses and eliminates tumor cells through characteristic features such as cell shrinkage, nuclear condensation, chromatin margination, and nuclear disintegration [26]. Upon receiving apoptotic signals, Bax/Bak-mediated cytochrome C release activates caspase-9 and caspase-3, leading to PARP cleavage and apoptotic cell death [27]. Tumor apoptosis and DNA damage are closely interconnected processes. DNA damage, often induced by factors such as radiation or chemical agents, can initiate a series of cellular responses, including apoptosis, to maintain genomic stability [28]. When DNA damage occurs, cell cycle checkpoints arrest the damaged cell cycle, providing time for DNA repair. In this study, co-administration of the treatment significantly induced nuclear alterations in cells, resulting in classic apoptotic features and markedly increasing the apoptosis rate, while also causing G2/M phase arrest in HepG2 and HuH-7 cells. In the combination groups, the cleavage of caspase-9, caspase-3, and PARP, as well as the levels of phosphorylated p53 and Bax, were notably increased. Therefore, co-administration induces apoptosis and promotes DNA damage in HepG2 and HuH-7 cells.

Following DNA damage, tumor cells initiate a self-repair mechanism. When the capacity for self-repair is insufficient to counteract the extensive DNA damage induced by drugs, tumor cells undergo programmed cell death or apoptosis [29]. DNA damage can occur in various forms, including base modifications, DNA cross-linking, single-strand breaks (SSBs), and DSBs. H2AX, a member of the histone H2A family, becomes rapidly phosphorylated at serine 139 upon DSB occurrence, converting the histone variant H2AX into γ-H2AX, also referred to as p-H2AX. The results demonstrate that combination treatment induces a pronounced comet tailing phenomenon, with the DNA content in the comet tails exceeding 40%. These findings indicate that simultaneous treatment with L-Arg and 5-FU significantly induces DNA damage in HCC cells. Furthermore, this combination markedly promotes DNA fragmentation and substantially increases γ-H2AX expression. Consequently, the combined use of L-Arg with 5-FU causes significant DNA damage within the cells. DNA-PKcs, ATM, and ATR belong to the PI3K family of protein kinases and repair damaged DNA through distinct pathways. DNA-PKcs is involved in multiple DNA damage repair processes and is closely associated with NHEJ development. The DNA-dependent protein kinase catalytic subunit recruits and phosphorylates NHEJ-related proteins. By suppressing DNA-PKcs expression, the activity of NHEJ-associated proteins in cells can be reduced, thereby increasing tumor cell susceptibility to DNA-damaging agents and inhibiting tumor cell proliferation [ 6, 30] . ATM serves as a key kinase that responds to DNA DSBs via two mechanisms: non-homologous end joining and homologous recombination repair [31]. After a DNA double-strand break occurs, the MRE11-RAD50-NBS1 (MRN) complex recognizes and binds to the break site, recruiting ATM to initiate the DNA damage response [32]. The histone protein H2AX is subsequently rapidly phosphorylated into γ-H2AX, which serves as a signaling platform to recruit additional repair factors, thereby amplifying the DNA damage response at the lesion site. Research has demonstrated that cooperation between ATM and the MRN complex is critical for ensuring efficient DNA repair [ 8, 33] . Upon DSB formation, ATM becomes activated and subsequently phosphorylates CHK2, leading to its activation. Activated CHK2 phosphorylates cdc25A, resulting in its degradation and preventing the removal of inhibitory phosphates from CDK2, thereby halting the progression of S phase [ 34, 35] . Additionally, both CHK2 and ATM can phosphorylate p53, stabilizing it and promoting the transcription of p21, which inhibits CDK2 activity and exacerbates cell cycle arrest. At stalled replication forks, single-stranded DNA (SS-DNA) activates ATR, which phosphorylates and activates CHK1, further contributing to cell cycle checkpoint control [36]. CHK1 phosphorylates CDC25C, thereby rendering it inactive. This inactivation prevents the removal of inhibitory phosphates from CDK1, effectively blocking the transition from the G2 phase to the M phase. Furthermore, DNA-PKcs, ATM, and ATR are all capable of phosphorylating H2AX. Studies have demonstrated that DNA-PKcs is predominantly associated with H2AX phosphorylation during apoptosis, ATM is primarily involved in H2AX phosphorylation during double-strand break processes, and ATR acts as the main kinase responsible for H2AX phosphorylation in response to single-strand damage and replication-induced formation of γ-H2AX [37]. The current findings revealed that co-administration of the treatment significantly reduced DNA-PKcs expression while increasing the expressions of phosphorylated ATM, ATR, and their respective substrates, including phosphorylated CHK1, CHK2, and BRCA1.

When a reduction in DNA-PKcs leads to decreased efficiency of NHEJ, co-administration of treatments in hepatocellular carcinoma cells may compensate by enhancing alternative repair pathways, such as HR. HR depends on the activation of ATM and ATR, which subsequently triggers increased phosphorylation of downstream proteins, including CHK1, CHK2, and BRCA1 [38]. Moreover, if NHEJ is impaired, unresolved double-strand breaks accumulate, leading to persistent DNA damage signaling. This continuous signaling activates ATM (in response to DSBs) and ATR (potentially through replication fork stalling or increased single-stranded DNA regions). Persistent activation of ATM and ATR typically indicates unresolved DNA damage or ongoing replication stress [39]. These kinases sustain the activation of checkpoint proteins such as CHK1 and CHK2, enforcing cell cycle arrest to allow for repair processes. If the damage is excessive or irreparable, sustained activation of these pathways causes prolonged cell cycle arrest, particularly at the G1/S or G2/M checkpoint, preventing propagation of damaged DNA [40]. When repair mechanisms fail, ATM and ATR signaling can trigger apoptotic pathways [ 41, 42] . This involves the activation of pro-apoptotic proteins, such as P53 (transcriptionally regulated by ATM), potentially leading to Mdm2 degradation and increased accumulation of p53 [43]. The role of ATM/ATR activation in the combined treatment of L-Arg and 5-FU was further verified via the use of a DNA-PKcs inhibitor (AZD-7648) and an ATM/ATR inhibitor (CGK733). AZD-7648 enhances the suppression of the NHEJ repair pathway induced by combined treatment with L-Arg and 5-FU. This inhibition shifts repair dynamics toward HR, a high-fidelity repair mechanism reliant on the activation of the ATM and ATR pathways. HR involves extensive signaling and precise processing of DNA damage, marked by the upregulation of critical proteins such as p-ATM, p-ATR, and γ-H2AX, which function as DSB markers to facilitate the assembly of HR repair complexes. CGK733, which has minimal effects on the NHEJ pathway, suppresses the activation of HR repair pathways induced by L-Arg and 5-FU, thereby reducing DNA damage and apoptosis. This underscores the intricate balance between the NHEJ and HR pathways in shaping cellular responses to DNA damage.

The PI3K/AKT pathway plays a pivotal role in regulating apoptosis in carcinoma cells and inhibiting DNA damage repair through both the NHEJ and HR pathways [44]. The two isoforms of AKT, AKT1 and AKT3, interact with the key NHEJ protein DNA-PKcs, promoting tumor cell proliferation and enhancing DNA repair following ionizing radiation-induced DNA damage [ 45, 46] . This study demonstrated that combination treatment decreases both phosphorylated PI3K and AKT expression, suggesting that this approach may inhibit the PI3K/AKT pathway, thereby suppressing DNA damage repair. Moreover, the application of LY294002, a PI3K inhibitor, effectively blocks downstream signaling events, making it an indispensable tool for investigating cancer, apoptosis, and other biological phenomena. The results demonstrated that LY294002 can further reduce the level of DNA-PKcs while increasing p-ATM and p-ATR expression when LY294002 is combined with these two drugs, indicating that the efficacy of combined treatment on the DNA damage repair process is closely associated with the suppression of the PI3K/AKT signaling pathway. When the PI3K/AKT pathway is inhibited, cells exhibit increased sensitivity to DNA damage due to reduced tolerance and survival capacity, leading to overactivation of ATM and ATR as compensatory mechanisms [47]. The observed suppression of the AKT pathway impairs DNA-PKcs-mediated DNA repair and promotes the activation of the ATM and ATR pathways, resulting in the formation of γ-H2AX, a marker of DNA damage, and ultimately the induction of apoptosis. Abnormal PI3K/AKT signaling triggers multiple molecular mechanisms that increase ROS levels either directly through mitochondrial bioenergetic regulation and NADPH oxidase (NOX) activation, or indirectly via ROS generation of ROS as metabolic byproducts [48]. The combination of L-Arg and 5-FU was found to increase ROS levels, disrupting cellular redox homeostasis and inducing oxidative stress. When NAC, a ROS inhibitor, was added to the cells, it alleviated AKT and DNA-PKcs suppression, as well as ATM and ATR activation, and increased the expression of cleaved caspase-3 and γ-H2AX caused by combined treatment with L-Arg and 5-FU. These results suggest that elevated ROS levels play a critical role in promoting DNA damage during combination therapy, highlighting the importance of ROS in mediating the synergistic effects of drugs. The NO scavenger carboxy-PTIO was found to reduce the increase in ROS induced by the combination of L-Arg and 5-FU. These findings underscore the role of NO in mediating ROS production during combination therapy, further supporting the involvement of the iNOS/NO pathway in the synergistic effects of L-Arg and 5-FU. The present results indicate that, upon the two cell lines were transfected with iNOS shRNA, the levels of DNA-PKcs and Bcl-2 were increased, whereas those of p-ATM, p-ATR, p-p53, and cleaved caspase-3 were significantly decreased. These results emphasize the therapeutic potential of targeting iNOS/NO-mediated ROS elevation in hepatocellular carcinoma cells and provide insight into the complex interplay between DNA repair suppression and apoptosis induction. In the primary HCC model in rats, the results demonstrated that co-administration reduced the levels of DNA-PKcs, p-AKT, and Bcl-2 while increasing the cleavage of caspase-9, caspase-3, and PARP, as well as the levels of iNOS, p-p53, p-ATM, and p-ATR.

In summary, the combination of L-Arg and 5-FU increases ROS levels through the iNOS/NO pathway, thereby inhibiting AKT signaling. This inhibition suppresses DNA-PKcs-mediated DNA repair mechanisms and subsequently activates the ATM and ATR pathways. As a result, this cascade induces sustained DNA damage, leading to G2/M phase cell cycle arrest and promoting apoptosis in hepatocellular carcinoma cells.

COMPETING INTERESTS

The authors declare that they have no conflict of interest.

Funding Statement

This work was supported by the grant from the Scientific Research Fund of Henan University of Science and Technology (No. 4020/13480023).