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. 2026 Jul 20;16:28081. doi: 10.1038/s41598-026-62807-6

Enoxaparin induces apoptosis and autophagy, modulates inflammatory signaling, and reduces oxidative DNA damage in breast and liver cancer cells

Sedat Carkit 1,✉, Münevver Baran 2, Nazmiye Bitgen 3,4, Gozde Ozge Onder 4,5,✉, Damla Gündüz 6, Arzu Yay 4,5
PMCID: PMC13554178  PMID: 42477385

Abstract

Cancer progression involves intricate interactions between inflammatory signaling, programmed cell death mechanisms, and oxidative stress. Although enoxaparin is widely used for managing cancer-associated thrombosis, its direct cellular effects on tumor biology remain insufficiently characterized. This study aimed to evaluate the impact of enoxaparin on apoptosis, autophagy, inflammatory mediators, and oxidative DNA damage in breast (MDA-MB-231) and liver (HepG2) cancer cell lines. MDA-MB-231, HepG2, and non-cancerous HEK-293 cells were treated with varying concentrations (5, 10, 20, 40, and 80 mg/mL) of enoxaparin for 24 and 48 h. Cell viability was assessed using the MTT assay, while apoptosis was quantified by TUNEL analysis. Immunofluorescence staining was employed to evaluate the expression of NF-κB, IL-6, TNF-α, LC3, and p62. Oxidative DNA damage was determined by measuring extracellular 8-hydroxy-2′-deoxyguanosine (8-OHdG) levels using a competitive ELISA. Statistical analyses were conducted to compare the treated and control groups. Enoxaparin significantly reduced cell viability in MDA-MB-231 and HepG2 cells without inducing cytotoxicity in HEK-293 cells. Apoptosis was markedly increased in both cancer cell lines following treatment. Enoxaparin differentially modulated inflammatory signaling; NF-κB expression was significantly increased in MDA-MB-231 cells, accompanied by suppression of IL-6 and TNF-α, whereas no significant inflammatory changes were observed in HepG2 cells. Enoxaparin treatment was observed to increase LC3 and p62 expression in both MDA-MB-231 and HepG2 cells, triggering autophagy-related pathways. Moreover, enoxaparin significantly reduced extracellular 8-OHdG levels, suggesting a reduction in oxidative DNA damage. Enoxaparin exhibits multifaceted anticancer effects by promoting apoptosis and autophagy, selectively modulating inflammatory pathways, and reducing oxidative DNA damage in breast and liver cancer cells.

Keywords: 8-OHdG, Oxidative DNA damage, Apoptosis, Autophagy, Inflammatory signaling

Subject terms: Biochemistry, Cancer, Cell biology, Drug discovery, Molecular biology, Oncology

Introduction

Cancer is widely recognized as the second leading cause of mortality worldwide and remains a significant public health challenge. Although biomedical research has expanded rapidly and in great depth, the availability of anticancer therapies that significantly reduce the overall disease burden remains inadequate. This situation underscores the continuing need for treatment strategies that are both more effective and grounded in contemporary insights into cancer biology, supported by advanced technological approaches1,2. The core objective of anticancer therapy is to eliminate malignant cells as selectively as possible while minimizing injury to normal tissues. In practice, conventional chemotherapeutic agents do not sufficiently discriminate between malignant and healthy cells, resulting in limited selectivity that leads to unavoidable toxicity and side effects in noncancerous tissues3. In patients with solid tumors, metastatic dissemination accounts for more than two-thirds of cancer-related deaths, and metastases are reported to contribute to approximately 66.7% of all deaths4. Beyond their central role in hemostasis, platelets have been shown to engage with circulating tumor cells (CTCs), shielding them from hemodynamic shear forces, supporting immune evasion—particularly from natural killer (NK) cells—and enhancing metastatic colonization by increasing intravascular stalling5–7. Consistent with this concept, experimental platelet depletion significantly reduced metastasis in mouse models, underscoring the importance of platelet–tumor cell interactions during metastatic progression. At the same time, it is well established that cytotoxic anticancer agents can disrupt essential cellular functions, including membrane integrity, metabolic activity, biosynthetic pathways, ion homeostasis, and cell division, thereby contributing to a broad spectrum of adverse effects8.

Venous thromboembolism (VTE), which includes deep vein thrombosis and pulmonary embolism, is a common complication in patients with cancer; it is estimated that 20–30% of all VTE cases are associated with malignancy. Cancer patients exhibit a 3- to 9-fold increased risk of VTE compared to the general population9. Current guidelines recommend anticoagulant therapy for at least 3–6 months in patients with cancer-associated thrombosis (CAT), with extended treatment in selected high-risk groups10–13. However, anticoagulation carries an inherent risk of bleeding, particularly in oncology patients, further complicating disease management11,14,15. These observations highlight the complex interplay between thrombosis and cancer biology, both of which contribute to poor prognosis and increased mortality16–18.

Cancer progression is closely associated with dysregulation of key biological processes, including inflammatory signaling, programmed cell death pathways, and oxidative stress. Nuclear factor-κB (NF-κB) and pro-inflammatory cytokines such as TNF-α and IL-6 play central roles in tumor-associated inflammation and metastasis, while apoptosis and autophagy are critical determinants of cancer cell survival and therapeutic response19,20. In addition, oxidative DNA damage, commonly assessed by 8-hydroxy-2′-deoxyguanosine (8-OHdG), is a major contributor to genomic instability and tumor progression21.

Enoxaparin, a widely used low-molecular-weight heparin, is routinely administered in the management of cancer-associated thrombosis22. Beyond its anticoagulant properties, emerging evidence suggests that enoxaparin may exert additional biological effects, including modulation of tumor growth, inflammation, and cellular stress responses23. However, its direct impact on key molecular pathways involved in apoptosis, autophagy, and oxidative stress in cancer cells remains insufficiently characterized.

Based on these considerations, we hypothesized that enoxaparin may exert cell type–specific regulatory effects on critical molecular pathways associated with cancer progression. To the best of our knowledge, no study has comprehensively evaluated the simultaneous effects of enoxaparin on apoptosis, autophagy, inflammatory mediators, and oxidative DNA damage in both MDA-MB-231 breast cancer and HepG2 hepatocellular carcinoma cell lines. Therefore, the aim of this study was to investigate the modulatory effects of enoxaparin on apoptotic signaling, autophagic activity, inflammatory pathways, and oxidative DNA damage in these cancer models, thereby providing insight into its potential biological roles beyond anticoagulation.

Materials and methods

Cell culture

The human breast cancer cell line MDA-MB-231, the human hepatocellular carcinoma cell line HepG2, and the human embryonic kidney cell line HEK-293 were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). All three cell lines (MDA-MB-231, HepG2, and HEK-293) were maintained in Dulbecco’s modified Eagle’s medium (DMEM; Gibco, Thermo Fisher Scientific, USA) supplemented with fetal bovine serum (10%, FBS, Sigma-Aldrich, USA), l-glutamine (1%, Gibco, USA), penicillin (100 units/ml, Gibco, USA), and streptomycin (100 mg/mL, Gibco, USA). Cells were incubated at 37 °C in a humidified atmosphere containing 5% CO2.

Cell viability assessment

For cell viability assessment, MDA-MB-231, HepG2, and HEK-293 cells were seeded into 96-well tissue culture plates at a density of 5 × 103 cells/mL 24 h before enoxaparin (Oksapar®, 6000 anti-Xa IU/0.6 mL, Koçak Farma, İstanbul, Türkiye) exposure. The cells were then treated with enoxaparin serially diluted in culture medium to final concentrations of 5, 10, 20, 40, and 80 mg/mL. For each concentration and cell type, four replicates were included, and incubation was performed for 24 and 48 h. Following the exposure period, 10 µL of MTT solution was added to each well, and the plates were incubated for 4 h at 37 °C. Subsequently, the culture medium was removed using a syringe. Formazan crystals were solubilized by adding 100 µl dimethyl sulfoxide to each well. Absorbance was measured at 570 nm using a plate reader24. Following absorbance measurements, cell viability was calculated as percentage relative to the corresponding time-matched control groups. The respective 24-h IC₅₀ concentrations determined from the MTT assay, were used in all subsequent TUNEL, immunofluorescence, and 8-OHdG experiments.

Analysis of apoptosis by TUNEL

To confirm DNA fragmentation associated with enoxaparin-induced apoptosis, a Terminal Deoxynucleotidyl Transferase-mediated dUTP Nick End Labeling (TUNEL) assay was conducted. MDA-MB-231 and HepG2 cells (5 × 104 cells/mL) were seeded on round coverslips in sterile 12-well culture plates. After 24 h of incubation, the cells were treated with enoxaparin for an additional 48 h. DNA strand breaks were detected by enzymatic end labelling using a commercial kit (Millipore S7110, Sigma Aldrich, US) in accordance with the manufacturer’s protocol, and the fluorescence signal was evaluated under a fluorescence microscope (Olympus BX74, Tokyo, Japan). TUNEL-positive cells were quantified from 10 randomly selected non-overlapping microscopic fields per sample using ImageJ software (ImageJ, Bethesda, USA)25.

Immunofluorescence assay

Immunofluorescence staining was performed to evaluate the expression profiles of nuclear factor-κB p65 (RelA) (NF-kB) (1:50, clone F-6, Cat. No. sc-8008, Santa Cruz Biotechnology, Texas, USA), Tumor necrosis factor-α (TNF-α) (1:100, Novus Biologicals, Littleton, USA), Interleukin 6 (IL-6) (1:100, Bioss, Massachusetts, USA), LC3 (1:250, Cell Signaling Technology, Danvers, USA), and p62 (1:200, Novus Biologicals, Littleton, USA). For protein expression quantification, ten images were randomly captured from microscopic fields at 400x magnification using an Olympus BX74 fluorescence microscope (Tokyo, Japan). Immunoreactivity intensity was quantified using ImageJ software (ImageJ, Bethesda, USA)26.

Oxidative DNA damage assay

Extracellular 8-hydroxy-2’-deoxyguanosine (8-OHdG) levels were determined to assess oxidative DNA damage. MDA-MB-231 and HepG2 cells were seeded at 1 × 106 cells/mL in 6-well plates and incubated for 24 h. Enoxaparin was then applied, and after 48 h of incubation, the cell culture supernatants were carefully collected to eliminate cellular debris. Extracellular 8-OHdG levels in the supernatant samples were measured using commercial kits based on the competitive chemiluminescence ELISA principle (Bioassay Technology Laboratory, Shanghai, China), strictly following the manufacturer’s instructions. Measurements were recorded at 450 nm, and analyte concentrations were calculated from the standard curve provided by the kit using the obtained absorbance values27.

Statistical analysis

All statistical analyses were performed using the Statistical Package for the Social Sciences (SPSS Statistics for Windows version 25.0; IBM Corp., Armonk, NY, USA). Quantitative data are expressed as mean ± standard deviation (SD). Before statistical comparisons, the normality of the data distribution was assessed using the Shapiro–Wilk test. For comparisons between two independent groups (control vs. enoxaparin-treated cells), the independent samples t-test was applied for normally distributed variables. In contrast, the Mann–Whitney U test was used for data that did not meet normality assumptions. For analyses involving multiple concentrations of enoxaparin, one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test was used for normally distributed datasets. Alternatively, the Kruskal–Wallis test followed by Dunn’s multiple comparison test was used for non-normally distributed data. Cell viability assays were performed in triplicate, and IC₅₀ values were calculated based on 48-hour dose–response curves using nonlinear regression analysis. Immunofluorescence intensity data (TUNEL, NF-κB, IL-6, TNF-α, LC3, p62, and 8-OHdG) were quantified from at least five randomly selected microscopic fields per sample and averaged before statistical evaluation. All statistical tests were two-tailed, and a p-value < 0.05 was considered statistically significant. The levels of statistical significance were denoted as follows: p < 0.05, p < 0.01, p < 0.001, and p < 0.0001.

Results

Effect of enoxaparin on cell viability

Cell viability values were normalized to the corresponding time-matched control groups and expressed as percentage viability. Enoxaparin treatment resulted in concentration- and time-dependent alterations in cell viability in both MDA-MB-231 and HepG2 cells. In MDA-MB-231 cells, cell viability remained relatively stable at lower concentrations, whereas significant reductions were observed at 40 mg/mL (p < 0.01) and 80 mg/mL (p < 0.001), particularly after 48 h exposure. Similarly, HepG2 cells exhibited a marked decrease in viability at higher concentrations of enoxaparin, with significant reductions detected at 40 mg/mL (p < 0.01) and 80 mg/mL (p < 0.001), with a more pronounced effect observed following 48 h treatment compared to 24 h. The IC₅₀ values determined from the 24-hour dose–response curves were 44.5 mg/mL for MDA-MB-231 cells and 40 mg/mL for HepG2 cells, and these concentrations were used in the subsequent experiments. In contrast, HEK-293 cells did not show a statistically significant reduction in viability across most concentrations and time points. Minor fluctuations observed at lower concentrations may reflect normal biological variability in proliferative non-cancerous HEK-293 cells rather than treatment-related cytotoxicity. Overall, these findings suggest that enoxaparin selectively decreases cell viability in cancer cell lines while largely preserving the viability of non-cancerous cells (Fig. 1).

Fig. 1.

Fig. 1

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Impact of enoxaparin on cell viability in (A) MDA-MB-231, (B) HepG2, and (C) HEK-293 cell lines after 24 and 48 h of treatment, as assessed by the MTT assay. Cell viability values were normalized to the corresponding time-matched control groups and expressed as percentage viability. Data are presented as mean ± SD. Statistical analysis was performed using two-way ANOVA. Statistical analyses were performed separately for the 24 h and 48 h groups. **p < 0.01, ***p < 0.001.

Effect of enoxaparin on apoptosis

The effect of enoxaparin on apoptosis in breast and liver cancer cell lines was evaluated using the TUNEL assay (Fig. 2). Enoxaparin treatment significantly increased apoptosis in both the MDA-MB-231 and HepG2 cell lines. In MDA-MB-231 cells, the median TUNEL positivity increased from 0.27 in the control group to 0.50 following enoxaparin treatment. Similarly, in HepG2 cells, the median apoptotic index rose from 0.27 in the control group to 0.57 after treatment. These increases were statistically highly significant in both cell lines (p < 0.0001) (Table 1; Fig. 3). Accordingly, comparisons between the control and enoxaparin-treated groups demonstrated statistically significant differences, indicating the pro-apoptotic effect of enoxaparin in both cell types.

Fig. 2.

Fig. 2

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TUNEL staining of MDA-MB-231 and HepG2 cells after 48 h enoxaparin treatment. Apoptotic cells appear as green fluorescence (×40).

Table 1.

Statistical analysis results of TUNEL of MDA-MB-231 and HepG2 cancer cell line at 48 h.

Control Enoxaparin p
MDA-MB-231 0.27(0.27–0.29) 0.50(0.45–0.58) < 0.0001
HEPG2 0.27(0.26–0.27) 0.57(0.53–0.64) < 0.0001
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Data are expressed as median (quarter1-quarter3). p: refers to the significance of the difference between the groups. p value of < 0.05 was used for significance.

Fig. 3.

Fig. 3

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Effects of enoxaparin on the intensity of TUNEL immunoreactivity in MDA-MB-231 and HepG2 cells. Cells were treated with enoxaparin for 48 h. Fluorescence intensity was quantified using Image J software. ***p < 0.0001.

Effect of enoxaparin on expression of NF-κB, TNF-α and IL-6

We investigated the impact of enoxaparin on the expression of NF-κB, TNF-α, and IL-6 in MDA-MB-231 and HepG2 cells. Protein expression levels were quantified using immunofluorescence analysis. Alterations in NF-κB, TNF-α, and IL-6 immunoreactivity were observed following 48-h enoxaparin treatment (Figs. 4, 5 and 6). In MDA-MB-231 cells, enoxaparin treatment resulted in a marked and statistically significant increase in NF-κB expression, rising from 0.36 ± 0.02 in the control group to 2.19 ± 0.58 in the treated group (p < 0.0001). In contrast, no significant change in NF-κB expression was observed in HepG2 cells, where levels remained comparable between control (0.42 ± 0.07) and enoxaparin-treated groups (0.43 ± 0.04) (p = 0.74) (Table 2; Fig. 7).

Fig. 4.

Fig. 4

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Representative immunofluorescence images of NF-κB expression in MDA-MB-231 and HepG2 cells after 48 h enoxaparin treatment. NF-κB is visualized as green fluorescence (×40).

Fig. 5.

Fig. 5

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Representative immunofluorescence images of TNF-α expression in MDA-MB-231 and HepG2 cells after 48 h enoxaparin treatment. TNF-α is visualized as green fluorescence (×40).

Fig. 6.

Fig. 6

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IL-6 expressions of the MDA-MB-231 and HepG2 cell lines treated with 48 h enoxaparin. Green immunofluorescence is reflective of IL-6 expressions under the fluorescent microscope (×40).

Table 2.

Statistical analysis results of NF-kB, TNF-α, and IL-6 expressions of MDA-MB-231 and HepG2 cancer cell line at 48 h.

Control Enoxaparin p
Nf-KB MDA-MB-231 0.36 ± 0.02 2.19 ± 0.58 < 0.0001
HEPG2 0.42 ± 0.07 0.43 ± 0.04 0.74
TNF-α MDA-MB-231 0.74 ± 0.23 0.45 ± 0.07 0.003
HEPG2 0.48 ± 0.08 0.46 ± 0.07 0.55
IL-6 MDA-MB-231 2.5 ± 0.70 1.49 ± 0.33 0.001
HEPG2 0.35 ± 0.05 0.34 ± 0.03 0.41
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Data are expressed as mean±standard deviation. p: refers to the significance of the difference between the groups. p value of < 0.05 was used for significance.

Fig. 7.

Fig. 7

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Immunoreactivity intensity of NF-κB, TNF-α, and IL-6 in MDA-MB-231 and HepG2 cells treated with enoxaparin for 48 h. Data are presented as mean ± SD (n = 10). Statistical analysis was performed using an unpaired t-test. **p < 0.01, ***p < 0.0001, ns: not significant.

Regarding TNF-α expression, enoxaparin significantly reduced its levels in MDA-MB-231 cells, decreasing from 0.74 ± 0.23 in the control group to 0.45 ± 0.07 following treatment (p = 0.003). However, no statistically significant difference was observed in HepG2 cells, with TNF-α levels remaining similar between control (0.48 ± 0.08) and treated groups (0.46 ± 0.07) (p = 0.55) (Table 2; Fig. 7).

Similarly, IL-6 expression was significantly decreased in MDA-MB-231 cells after enoxaparin exposure, declining from 2.50 ± 0.70 in the control group to 1.49 ± 0.33 in the treated group (p = 0.001). In contrast, HepG2 cells showed no significant change in IL-6 expression, with values of 0.35 ± 0.05 in the control group and 0.34 ± 0.03 in the treated group (p = 0.41) (Table 2; Fig. 7). Overall, these findings indicate that enoxaparin selectively modulates inflammatory signaling in MDA-MB-231 cells, while exerting minimal effects on these markers in HepG2 cells.

Effect of enoxaparin on autophagy

Autophagy-related protein expression was analyzed by assessing LC3 and p62 levels in both MDA-MB-231 and HepG2 cells following enoxaparin treatment. Figures 8 and 9 illustrate LC3 and p62 expression profiles, respectively. In MDA-MB-231 cells, LC3 expression increased significantly from a median value of 1.01 in the control group to 1.51 following enoxaparin treatment (p = 0.006). Similarly, in HepG2 cells, LC3 levels showed a marked and statistically significant elevation, rising from 0.47 in the control group to 0.91 in the treated group (p < 0.0001) (Table 3; Fig. 10).

Fig. 8.

Fig. 8

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Representative immunofluorescence images of LC3 expression in MDA-MB-231 and HepG2 cells after 48 h enoxaparin treatment. LC3 is visualized as green fluorescence (×40).

Fig. 9.

Fig. 9

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p62 expressions of the MDA-MB-231 and HepG2 cell lines treated with enoxaparin. Red immunofluorescence is reflective of p62 expressions under the fluorescent microscope (×40).

Table 3.

Statistical analysis results of LC3 and p62 expressions of MDA-MB-231 and HepG2 cancer cell line at 48 h.

Control Enoxaparin p
LC3 MDA-MB-231 1.01(0.84–1.08) 1.51(1.06–1.82) 0.006
HEPG2 0.47(0.42–0.62) 0.91(0.71–1.68) < 0.0001
P62 MDA-MB-231 2.73(2.14–3.78) 3.60(3.46–4.72) 0.024
HEPG2 0.58(0.46–0.68) 0.93(0.76–1.57) 0.0002
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Data are expressed as median (quarter1-quarter3). p: refers to the significance of the difference between the groups. p value of < 0.05 was used for significance.

Fig. 10.

Fig. 10

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Immunoreactivity intensity of LC3, and p62 in MDA-MB-231 and HepG2 cells treated with enoxaparin for 48 h. Data are presented as median (Q1–Q3) (n = 10). Statistical analysis was performed using the Mann–Whitney U test. *p < 0.05, **p < 0.01, ***p < 0.0001.

A comparable pattern was observed for p62 expression. In MDA-MB-231 cells, p62 levels significantly increased from 2.73 in the control group to 3.60 following enoxaparin treatment (p = 0.024). Likewise, HepG2 cells exhibited a significant rise in p62 expression, from 0.58 in the control group to 0.93 after treatment (p = 0.0002) (Table 3; Fig. 10). Overall, these findings indicate that enoxaparin significantly enhances the expression of autophagy-related markers LC3 and p62 in both breast and liver cancer cell lines.

Effect of enoxaparin on 8-OHdG

Table 4 summarizes the statistical analysis of oxidative DNA damage in MDA-MB-231 and HepG2 cells. In MDA-MB-231 cells, 8-OHdG levels decreased significantly from 262.4 ± 2.73 in the control group to 108.0 ± 0.12 following enoxaparin treatment (p < 0.0001). Similarly, in HepG2 cells, a pronounced reduction was observed, with 8-OHdG levels declining from 84.51 ± 1.22 in the control group to 2.27 ± 0.18 in the treated group (p < 0.0001) (Fig. 11). Overall, these findings demonstrate that enoxaparin significantly reduces oxidative DNA damage in both breast and liver cancer cell lines.

Table 4.

Statistical analysis results of 8-OHdG (ng/mL) in MDA-MB-231 and HepG2 cancer cell line at 48 h.

Control Enoxaparin p
MDA-MB-231 262.4 ± 2.73 108.0 ± 0.12 < 0.0001
HEPG2 84.51 ± 1.22 2.27 ± 0.18 < 0.0001
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Since the data showed normal distribution, data are expressed as mean ± standard deviation. p: refers to the significance of the difference between the groups. p value of < 0.05 was used for significance.

Fig. 11.

Fig. 11

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Bar graph showing mean 8-hydroxy-2′-deoxyguanosine (8-OHdG) levels in MDA-MB-231 and HepG2 cells following enoxaparin treatment. Group comparisons were performed using an independent samples t-test. Data are presented as mean ± SD, ***p < 0.0001.

Discussion

This study aimed to elucidate the interaction between enoxaparin and MDA-MB-231 and HepG2 cancer cells and to determine how this interaction influences key pathways involved in tumor progression. In the present study, enoxaparin did not exert cytotoxic effects on healthy cells but significantly promoted cell death in both MDA-MB-231 and HepG2 cancer cells through pro-inflammatory mediators, apoptosis and autophagy-related mechanisms. These findings highlight a selective anticancer effect, suggesting that enoxaparin may preferentially target malignant cells while sparing non-cancerous cells.

Low-molecular-weight heparins (LMWHs) have been extensively used in the management of cancer-associated thrombosis and have also been associated with potential antitumor effects. Previous studies have demonstrated that enoxaparin can inhibit tumor cell proliferation, migration, and metastasis in various experimental models28–31. However, the clinical relevance of these findings remains uncertain, as many studies involve patients receiving concurrent chemotherapy32,33. In this context, our results provide additional mechanistic insight by demonstrating that enoxaparin directly modulates multiple cellular pathways in cancer cells, independent of its anticoagulant function, consistent with emerging evidence that anticancer strategies may act not only through direct cytotoxicity but also by modulating protein stability and stress-related cellular pathways, including protein aggregation and degradation mechanisms34,35. The concentrations used in the present study were higher than clinically achievable plasma levels of enoxaparin. However, in vitro studies often require higher concentrations to overcome the absence of systemic metabolism, protein binding, and tissue distribution and to achieve detectable cellular responses under controlled experimental conditions. In the present study, treatment concentrations were selected based on preliminary dose–response analyses and measurable biological effects in cancer cells, while no significant cytotoxicity was observed in non-cancerous HEK-293 cells36,37.

Apoptosis is a critical determinant of tumor progression and therapeutic response38. In the present study, enoxaparin significantly increased apoptosis in both MDA-MB-231 and HepG2 cells, as evidenced by elevated TUNEL positivity. This consistent pro-apoptotic effect across two distinct cancer cell types suggests that enoxaparin may activate conserved cell death pathways that are not restricted to a specific tissue origin. These findings are consistent with previous reports showing variable apoptosis-related effects depending on cellular context and concentration36,39. Moreover, the heterogeneous composition of enoxaparin, including pro- and antiproliferative oligosaccharides, may contribute to these differential responses39.

Inflammation is closely linked to both thrombosis and cancer progression40. IL-6 and TNF-α are key mediators of inflammatory signaling and immune regulation41,42, while NF-κB represents a central pathway controlling inflammatory responses and tumor progression43. Previous studies have shown that platelet–tumor interactions can enhance tumor invasion through NF-κB activation44,45. Under basal conditions, NF-κB is sequestered in the cytoplasm by IκB proteins and translocates to the nucleus upon activation to regulate gene expression46,47 In the present study, enoxaparin exerted cell type–specific effects on inflammatory mediators. Notably, NF-κB expression was significantly increased in MDA-MB-231 cells, whereas TNF-α and IL-6 levels were significantly reduced. This pattern suggests that enoxaparin may regulate inflammatory signaling through NF-κB–independent mechanisms or through post-transcriptional regulation of cytokine expression. In contrast, the absence of significant changes in HepG2 cells supports the notion that enoxaparin’s immunomodulatory effects are dependent on tumor-specific signaling contexts. These findings emphasize the complexity of the thrombo-inflammatory axis and its modulation by LMWHs48–50. Also, in MDA-MB-231 cells, the observed increase in cytosolic NF-κB expression compared to the control suggests a robust induction of the pathway, though the protein remains primarily localized in the cytoplasm at this time point. In contrast, the marked nuclear localization in HepG2 cells indicates a more rapid or efficient translocation, potentially reflecting earlier activation kinetics in this cell line. These results highlight that Enoxaparin modulates NF-κB signaling through cell-type-specific kinetic profiles.

Autophagy is a fundamental cellular process involved in maintaining homeostasis and regulating tumor progression51–54. Depending on the cellular context, autophagy may either promote tumor survival or contribute to programmed cell death55,56. It has also been implicated in tumor growth, metastasis, stemness, and chemoresistance57–64. In the present study, enoxaparin significantly increased LC3 and p62 expression in both MDA-MB-231 and HepG2 cells, indicating activation of autophagy-related pathways. It has been reported that p62 levels can increase during prolonged stress despite active autophagy, due to its transcriptional upregulation via the Nrf2 pathway65. Therefore, the simultaneous increase in LC3 and p62 in our 48-hour samples may reflect a high rate of autophagosome formation that temporarily exceeds the degradative capacity, or a compensatory transcriptional increase in p62 expression. This consistent response suggests that enoxaparin induces a conserved cellular stress mechanism across different tumor types.

Oxidative stress plays a crucial role in carcinogenesis by inducing DNA damage and genomic instability66–68. 8-hydroxy-2′-deoxyguanosine (8-OHdG) is a well-established biomarker of oxidative DNA damage and has been associated with cancer prognosis and progression68–70. Elevated levels of 8-OHdG have been reported in various tumor types, including breast, bladder, kidney, and other cancers71–74. In the present study, enoxaparin significantly reduced 8-OHdG levels in both MDA-MB-231 and HepG2 cells, indicating a protective effect against oxidative DNA damage. This finding suggests that enoxaparin may contribute to the regulation of redox homeostasis. The simultaneous modulation of oxidative stress, apoptosis, and autophagy further supports the concept that enoxaparin induces a coordinated cellular response involving multiple interconnected pathways.

Limitations

This study has several limitations that should be acknowledged. First, the enoxaparin concentrations used exceeded clinically achievable plasma levels; although this is common in in vitro pharmacological studies, it limits direct extrapolation to clinical dosing. Second, the analysis was restricted to two cancer cell lines (MDA-MB-231 and HepG2) without validation in additional cell lines or in vivo tumor models, which limits the generalizability of the findings. Third, the mechanistic pathways identified here are based on correlative expression changes rather than direct causal manipulation, such as pathway inhibition or gene silencing; future studies incorporating such approaches are needed to establish causality. Finally, functional endpoints relevant to tumor progression, including migration, invasion, and in vivo tumor growth, were not assessed in the present study.

Conclusion

In conclusion, the present study provides in vitro evidence that enoxaparin produces multifaceted anticancer effects in MDA-MB-231 and HepG2 cells through coordinated modulation of apoptosis, autophagy, inflammatory signaling, and oxidative DNA damage. Taken together, these findings indicate that enoxaparin may have biological relevance beyond its role in thromboprophylaxis. Although the data were generated under in vitro conditions, they provide a mechanistic rationale for further investigation of enoxaparin’s potential relevance to oncologic treatment strategies. Future studies, including in vivo and clinical investigations, are needed to clarify whether these cellular effects translate into improved therapeutic responses, reduced metastatic potential, and better clinical outcomes in cancer patients.

Author contributions

S.C. and N.B. conceived of and designed the study. Material preparation, data collection, and analyses were performed by S.C., N.B., M.B., G. O., and D.G. The first draft of the manuscript was written by S.C., N.B., G. O., D.G., and A.Y. critically reviewed and edited the manuscript. A.Y. supervised the study and provided critical revisions. All authors have read and approved the final manuscript.

Data availability

Data supporting the findings of this study are available from the corresponding author upon reasonable request.

Declarations

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Contributor Information

Sedat Carkit, Email: sedatcarkit@erciyes.edu.tr, Email: opdrsedatcarkit@gmail.com.

Gozde Ozge Onder, Email: gozdeozgekorkmaz@hotmail.com.

References

  • 1.Kelkar, S. S. & Reineke, T. M. Theranostics: Combining imaging and therapy. Bioconjug. Chem.22 (10), 1879–1903 (2011). [DOI] [PubMed] [Google Scholar]
  • 2.Purkait, K., Chatterjee, S., Karmakar, S. & Mukherjee, A. Alteration of steric hindrance modulates glutathione resistance and cytotoxicity of three structurally related Ru II-p-cymene complexes. Dalton Trans.45 (20), 8541–8555 (2016). [DOI] [PubMed] [Google Scholar]
  • 3.Chandra, A., Singh, K., Singh, S., Sivakumar, S. & Patra, A. K. A luminescent europium (III)–platinum (II) heterometallic complex as a theranostic agent: A proof-of-concept study. Dalton Trans.45 (2), 494–497 (2016). [DOI] [PubMed] [Google Scholar]
  • 4.Dillekås, H., Rogers, M. S. & Straume, O. Are 90% of deaths from cancer caused by metastases? Cancer Med.8 (12), 5574–5576 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Labelle, M. & Hynes, R. O. The initial hours of metastasis: The importance of cooperative host–tumor cell interactions during hematogenous dissemination. Cancer Discov.2 (12), 1091–1099 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Morris, K., Schnoor, B. & Papa, A-L. Platelet cancer cell interplay as a new therapeutic target. Biochim. Biophys. Acta (BBA)-Rev. Cancer1877 (5), 188770 (2022). [DOI] [PubMed] [Google Scholar]
  • 7.Schlesinger, M. Role of platelets and platelet receptors in cancer metastasis. J. Hematol. Oncol.11 (1), 125 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Seibert, H. et al. Acute toxicity testing in vitro and the classification and labelling of chemicals: The report and recommendations of ECVAM Workshop 16. Altern. Lab. Anim.24 (4), 499–510 (1996). [Google Scholar]
  • 9.Timp, J. F., Braekkan, S. K., Versteeg, H. H. & Cannegieter, S. C. Epidemiology of cancer-associated venous thrombosis. Blood J. Am. Soc. Hematol.122 (10), 1712–1723 (2013). [DOI] [PubMed] [Google Scholar]
  • 10.Falanga, A. et al. Venous thromboembolism in cancer patients: ESMO Clinical Practice Guideline. Ann. Oncol.34 (5), 452–467 (2023). [DOI] [PubMed] [Google Scholar]
  • 11.Konstantinides, S. V. et al. 2019 ESC Guidelines for the diagnosis and management of acute pulmonary embolism developed in collaboration with the European Respiratory Society (ERS) The Task Force for the diagnosis and management of acute pulmonary embolism of the European Society of Cardiology (ESC). Eur. Heart J.41 (4), 543–603 (2020). [DOI] [PubMed] [Google Scholar]
  • 12.Lobo, J. L. et al. WITHDRAWN: Multidisciplinary consensus for the management of pulmonary thromboembolism. In Archivos de Bronconeumología (English Edition) (2021).
  • 13.Ortel, T. L. et al. American Society of Hematology 2020 guidelines for management of venous thromboembolism: Treatment of deep vein thrombosis and pulmonary embolism. Blood Adv.4 (19), 4693–4738 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Farge, D. et al. 2022 international clinical practice guidelines for the treatment and prophylaxis of venous thromboembolism in patients with cancer, including patients with COVID-19. Lancet Oncol.23 (7), e334–e47 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Khorana, A. A. et al. Cancer-associated venous thromboembolism. Nat. Rev. Dis. Primers8 (1), 11 (2022). [DOI] [PubMed] [Google Scholar]
  • 16.Ording, A. G. et al. Increasing incidence and declining mortality after cancer-associated venous thromboembolism: A nationwide cohort study. Am. J. Med.134 (7), 868–876 (2021). e5. [DOI] [PubMed] [Google Scholar]
  • 17.Overvad, T. F. et al. Validation of the Khorana score for predicting venous thromboembolism in 40 218 patients with cancer initiating chemotherapy. Blood Adv.6 (10), 2967–2976 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Weitz, J. I. et al. Cancer associated thrombosis in everyday practice: Perspectives from GARFIELD-VTE. J. Thromb. Thromb. 50 (2), 267–277 (2020). [DOI] [PubMed] [Google Scholar]
  • 19.Moneva-Sakelarieva, M. et al. The role of the transcription factor NF-kB in the pathogenesis of inflammation and carcinogenesis. Modulation capabilities. Pharmacia72, 1–13 (2025). [Google Scholar]
  • 20.Sethi, G., Shanmugam, M. K., Ramachandran, L., Kumar, A. P. & Tergaonkar, V. Multifaceted link between cancer and inflammation. Biosci. Rep.32 (1), 1–15 (2012). [DOI] [PubMed] [Google Scholar]
  • 21.Valavanidis, A., Vlachogianni, T. & Fiotakis, C. 8-hydroxy-2′-deoxyguanosine (8-OHdG): A critical biomarker of oxidative stress and carcinogenesis. J. Environ. Sci. Health Part. C27 (2), 120–139 (2009). [DOI] [PubMed] [Google Scholar]
  • 22.Elbadawi, A., Shnoda, M., Mahmoud, K. & Elgendy, I. Y. Efficacy and safety of direct oral anticoagulants vs. low molecular weight heparin for cancer-related venous thromboembolism: A meta-analysis of randomized trials. Eur. Heart J. Cardiovasc. Pharmacother.7 (5), 380–388 (2021). [DOI] [PubMed] [Google Scholar]
  • 23.Russo, V. et al. Anti-inflammatory and anticancer effects of anticoagulant therapy in patients with malignancy. Life13 (9), 1888 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Verso, M. et al. Recurrent venous thromboembolism and major bleeding in patients with localised, locally advanced or metastatic cancer: An analysis of the Caravaggio study. Eur. J. Cancer165, 136–145 (2022). [DOI] [PubMed] [Google Scholar]
  • 25.Baran, M., Bitgen, N., Onder, G. O., Gurbuz, P. & Yay, A. Cell proliferation, apoptosis and inflammation response of Melissa officinalis and Thymus vulgaris in SW480 colon cancer cells. S. Afr. J. Bot.162, 282–295 (2023). [Google Scholar]
  • 26.Bıtgen, N., Onder, G. O., Baran, M. & Yay, A. Cytotoxicity screening of Thymus vulgaris L. in breast cancer: In vitro study. Toxicol. Res.12 (4), 584–590 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Morita, M., Yano, S., Yamaguchi, T. & Sugimoto, T. Advanced glycation end products-induced reactive oxygen species generation is partly through NF-kappa B activation in human aortic endothelial cells. J. Diabetes Complic.27 (1), 11–15 (2013). [DOI] [PubMed] [Google Scholar]
  • 28.Balzarotti, M. et al. In vitro study of low molecular weight heparin effect on cell growth and cell invasion in primary cell cultures of high-grade gliomas. Oncol. Res.16 (5), 245–250 (2006). [DOI] [PubMed] [Google Scholar]
  • 29.Ellensen, V. S., Abrahamsen, I., Lorens, J. & Jonung, T. Effects of enoxaparin and dalteparin on proliferation and migration of patient-derived vascular smooth muscle cells. Vasa43 (2), 124–131 (2014). [DOI] [PubMed] [Google Scholar]
  • 30.Ichikawa, J. et al. Thrombin induces osteosarcoma growth, a function inhibited by low molecular weight heparin in vitro and in vivo: Procoagulant nature of osteosarcoma. Cancer118 (9), 2494–2506 (2012). [DOI] [PubMed] [Google Scholar]
  • 31.Mousa, S. A., Linhardt, R., Francis, J. L. & Amirkhosravi, A. Anti-metastatic effect of a non-anticoagulant low-molecular-weight heparin versus the standard low-molecular-weight heparin, enoxaparin. Thromb. Haemost.96 (12), 816–821 (2006). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Niu, Q. et al. Low molecular weight heparin ablates lung cancer cisplatin-resistance by inducing proteasome-mediated ABCG2 protein degradation. PLoS One. 7 (7), e41035 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Pan, Y. et al. Enoxaparin sensitizes human non-small-cell lung carcinomas to gefitinib by inhibiting DOCK1 expression, vimentin phosphorylation, and Akt activation. Mol. Pharmacol.87 (3), 378–390 (2015). [DOI] [PubMed] [Google Scholar]
  • 34.Yang, C. et al. Heat stress targets and degrades BCR:: ABL1 oncoproteins to overcome drug-resistance in Philadelphia chromosome-positive acute lymphoblastic leukemia: Acute lymphoblastic leukemia. Leukemia39 (9), 2140–2151 (2025). [DOI] [PubMed] [Google Scholar]
  • 35.Yu, P-H., Zhu, C-Y., Kang, Y-Y., Naranmandura, H. & Yang, C. Mutation in the unrearranged PML allele confers resistance to arsenic trioxide in acute promyelocytic leukemia. Research8, 0696 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Abu Arab, W., Kotb, R., Sirois, M. & Rousseau, E. Concentration-and time-dependent effects of enoxaparin on human adenocarcinomic epithelial cell line A549 proliferation in vitro. Can. J. Physiol. Pharmacol.89 (10), 705–711 (2011). [DOI] [PubMed] [Google Scholar]
  • 37.Young, E., Cosmi, B., Weitz, J. & Hirsh, J. Comparison of the non-specific binding of unfractionated heparin and low molecular weight heparin (Enoxaparin) to plasma proteins. Thromb. Haemost.70 (10), 625–630 (1993). [PubMed] [Google Scholar]
  • 38.Nikoletopoulou, V., Markaki, M., Palikaras, K. & Tavernarakis, N. Crosstalk between apoptosis, necrosis and autophagy. Biochim. Biophys. Acta (BBA)-Mol. Cell. Res.1833 (12), 3448–3459 (2013). [DOI] [PubMed] [Google Scholar]
  • 39.Lean, Q. Y., Patel, R. P., Stewart, N., Sohal, S. S. & Gueven, N. Identification of pro-and anti-proliferative oligosaccharides of heparins. Integr. Biol. 6 (1), 90–99 (2014). [DOI] [PubMed] [Google Scholar]
  • 40.Rawish, E., Nording, H., Münte, T. & Langer, H. F. Platelets as mediators of neuroinflammation and thrombosis. Front. Immunol.11, 548631 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Holst, R-M. et al. Prediction of spontaneous preterm delivery in women with preterm labor: Analysis of multiple proteins in amniotic and cervical fluids. Obstet. Gynecol.114 (2 Part 1), 268–277 (2009). [DOI] [PubMed] [Google Scholar]
  • 42.Sorokin, Y. et al. Maternal serum interleukin-6, C-reactive protein, and matrix metalloproteinase-9 concentrations as risk factors for preterm birth < 32 weeks and adverse neonatal outcomes. Am. J. Perinatol.27 (08), 631–640 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Jung, T. W., Park, H. S., Choi, G. H., Kim, D. & Lee, T. β-Aminoisobutyric acid attenuates LPS-induced inflammation and insulin resistance in adipocytes through AMPK-mediated pathway. J. Biomed. Sci.25 (1), 27 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Liao, K. et al. The role of platelets in the regulation of tumor growth and metastasis: The mechanisms and targeted therapy. MedComm4 (5), e350 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Xia, Y., Shen, S. & Verma, I. M. NF-κB, an active player in human cancers. Cancer Immunol. Res.2 (9), 823–830 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Oeckinghaus, A. & Ghosh, S. The NF-κB family of transcription factors and its regulation. Cold Spring Harb. Perspect. Biol.1 (4), a000034 (2009). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Liu, T., Zhang, L., Joo, D. & Sun, S-C. NF-κB signaling in inflammation. Signal. Transduct. Target. therapy. 2 (1), 1–9 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Chen, X. et al. Nepetin inhibits IL-1β induced inflammation via NF-κB and MAPKs signaling pathways in ARPE-19 cells. Biomed. Pharmacother.101, 87–93 (2018). [DOI] [PubMed] [Google Scholar]
  • 49.Franks, Z. et al. Methicillin-resistant Staphylococcus aureus-induced thrombo-inflammatory response is reduced with timely antibiotic administration. Thromb. Haemost.109 (04), 684–695 (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Tavares, V., Pinto, R., Assis, J., Pereira, D. & Medeiros, R. Venous thromboembolism GWAS reported genetic makeup and the hallmarks of cancer: Linkage to ovarian tumour behaviour. Biochim. Biophys. Acta (BBA)-Rev. Cancer ;1873(1), 188331 (2020). [DOI] [PubMed]
  • 51.Amaravadi, R. K., Kimmelman, A. C. & Debnath, J. Targeting autophagy in cancer: Recent advances and future directions. Cancer Discov.9 (9), 1167–1181 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Lu, G. et al. Autophagy in health and disease: From molecular mechanisms to therapeutic target. MedComm3 (3), e150 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Rybstein, M. D., Bravo-San Pedro, J. M., Kroemer, G. & Galluzzi, L. The autophagic network and cancer. Nat. Cell Biol.20 (3), 243–251 (2018). [DOI] [PubMed] [Google Scholar]
  • 54.Tyutyunyk-Massey, L. & Gewirtz, D. A. (eds) Roles of Autophagy in Breast Cancer Treatment: Target, Bystander or Benefactor. Seminars in Cancer Biology (Elsevier, 2020). [DOI] [PubMed]
  • 55.Fan, X. et al. Daphnetin triggers ROS-induced cell death and induces cytoprotective autophagy by modulating the AMPK/Akt/mTOR pathway in ovarian cancer. Phytomedicine82, 153465 (2021). [DOI] [PubMed] [Google Scholar]
  • 56.Markov, A. V. et al. Novel derivatives of deoxycholic acid bearing linear aliphatic diamine and aminoalcohol moieties and their cyclic analogs at the C3 position: Synthesis and evaluation of their in vitro antitumor potential. Molecules24 (14), 2644 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 57.Entrialgo-Cadierno, R. et al. The phospholipid transporter PITPNC1 links KRAS to MYC to prevent autophagy in lung and pancreatic cancer. Mol. Cancer. 22 (1), 86 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 58.Fang, L. et al. Circular CPM promotes chemoresistance of gastric cancer via activating PRKAA2-mediated autophagy. Clin. Transl. Med.12 (1), e708 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59.Luo, Y. et al. Long noncoding RNA (lncRNA) EIF3J-DT induces chemoresistance of gastric cancer via autophagy activation. Autophagy17 (12), 4083–4101 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60.Park, J. W. et al. Autophagy inhibits cancer stemness in triple-negative breast cancer via miR‐181a‐mediated regulation of ATG5 and/or ATG2B. Mol. Oncol.16 (9), 1857–1875 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 61.Ren, K. et al. A neutrophil-mediated carrier regulates tumor stemness by inhibiting autophagy to prevent postoperative triple-negative breast cancer recurrence and metastasis. Acta Biomater.145, 185–199 (2022). [DOI] [PubMed] [Google Scholar]
  • 62.Ryu, K-J. et al. Chaperone-mediated autophagy modulates snail protein stability: Implications for breast cancer metastasis. Mol. Cancer. 23 (1), 227 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 63.Xing, Y. et al. Autophagy inhibition mediated by MCOLN1/TRPML1 suppresses cancer metastasis via regulating a ROS-driven TP53/p53 pathway. Autophagy18 (8), 1932–1954 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 64.Ye, M. et al. FABP5 suppresses colorectal cancer progression via mTOR-mediated autophagy by decreasing FASN expression. Int. J. Biol. Sci.19 (10), 3115 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 65.Sahani, M. H., Itakura, E. & Mizushima, N. Expression of the autophagy substrate SQSTM1/p62 is restored during prolonged starvation depending on transcriptional upregulation and autophagy-derived amino acids. Autophagy10 (3), 431–441 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 66.Klaunig, J. E. & Kamendulis, L. M. The role of oxidative stress in carcinogenesis. Annu. Rev. Pharmacol. Toxicol.44 (1), 239–267 (2004). [DOI] [PubMed] [Google Scholar]
  • 67.Skinner, A. M. & Turker, M. S. Oxidative mutagenesis, mismatch repair, and aging. Sci. Aging Knowl. Environ.2005 (9), re3–re (2005). [DOI] [PubMed] [Google Scholar]
  • 68.Yun, C. W. & Lee, S. H. The roles of autophagy in cancer. Int. J. Mol. Sci.19 (11), 3466 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 69.Lee, J. D., Cai, Q., Shu, X. O. & Nechuta, S. J. The role of biomarkers of oxidative stress in breast cancer risk and prognosis: A systematic review of the epidemiologic literature. J. Women’s Health. 26 (5), 467–482 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 70.Shukla, S. et al. Oxidative stress and antioxidant status in high-risk prostate cancer subjects. Diagnostics10 (3), 126 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 71.Hamidavi, A. et al. Lesion (8-hydroxydeoxyguanosine), 8-oxoguanine DNA glycosylase 1, and antioxidant status in Iranian patients diagnosed with high-grade oral squamous cell carcinoma. Oral Maxillofac. Pathol. J.8(2) (2017).
  • 72.Wada, T. et al. Mitochondrial DNA mutations and 8-hydroxy-2′-deoxyguanosine content in Japanese patients with urinary bladder and renal cancers. Anticancer Res.26 (5A), 3403–3408 (2006). [PubMed] [Google Scholar]
  • 73.Burger, M. S., Torino, J. L. & Swaminathan, S. DNA damage in human transitional cell carcinoma cells after exposure to the proximate metabolite of the bladder carcinogen 4-aminobiphenyl. Environ. Mol. Mutagen.38 (1), 1–11 (2001). [DOI] [PubMed] [Google Scholar]
  • 74.Matsui, A. et al. Increased formation of oxidative DNA damage, 8-hydroxy-2′-deoxyguanosine, in human breast cancer tissue and its relationship to GSTP1 and COMT genotypes. Cancer Lett.151 (1), 87–95 (2000). [DOI] [PubMed] [Google Scholar]

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Data Availability Statement

Data supporting the findings of this study are available from the corresponding author upon reasonable request.

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