Amelioration of Acrolein‐Induced Cytotoxicity, Oxidative Stress, DNA Damage, Apoptosis and Mitochondrial Dysfunction in Human Pancreas Cells by Pycnogenol

ABSTRACT

Acrolein (AC) is a potential environmental contaminant that poses a serious threat to human health and the environment. Studies on human diseases that can result from AC have rapidly increased in recent years. Pycnogenol (PYC), a natural phenolic compound, has therapeutic and protective potential against some cancers. Currently, there is insufficient data on the potential effects of AC and PYC on pancreatic cells or on their interaction. Because toxicological effects of AC and interactions between AC and PYC in human pancreatic cells have not yet been investigated, we evaluated the toxicological profile of AC and possible protective effect of PYC on healthy human pancreatic cells (hTERT‐HPNE). Our results showed that AC (25, 50, and 100 µM) increased cytotoxicity and oxidative stress, induced apoptosis and DNA single‐ and double‐strand breaks, whereas it decreased mitochondrial membrane potential and antioxidant parameters in human pancreatic cells. Combined treatments with AC and PYC (100 µg/mL) improved these parameters compared to AC treatment alone. Therefore, our results suggest that PYC may have a protective effect against AC‐induced damage. Our results may also be particularly important for identifying the factors involved in the pathogenesis of mitochondrial diseases, which have rapidly increased in recent years, and for providing preventive solutions.

Acrolein induced oxidative stress, DNA damage, mitochondrial dysfunction, and apoptosis in human urothelial cells. Pycnogenol ameliorated acrolein‐induced cytotoxicity.

1. Introduction

Acrolein (AC) is a chemical compound classified as a highly reactive unsaturated aldehyde and environmental pollutant. It is found almost everywhere and poses a potentially serious threat to environmental health. Human exposure to this substance occurs through a variety of sources, including cigarette smoke, industrial emissions, overheated cooking oils, and combustion products [1, 2]. The excessive production of AC from these sources has always posed a global health risk [3].

On the cellular level, AC can induce DNA damage, reactive oxygen species (ROS) formation, oxidative stress, mitochondrial disruption, membrane damage, endoplasmic reticulum stress, and immune dysfunction [1, 4, 5, 6, 7, 8, 9, 10, 11]. AC disrupted the oxidant/antioxidant balance after decreasing antioxidant capacity and increasing oxidative injury markers [9]. AC increased the formation of ROS formation and oxidative stress by decreasing the antioxidant defenses and by depleting glutathione (GSH) levels in various cell types [3, 10]. AC induced DNA damage involving both single‐strand and double‐strand breaks (DSBs) as well as oxidative base modifications and DDR activation. It activated apoptosis, particularly intrinsic (mitochondrial) pathways linked to oxidative stress and genetic damage [3]. AC can stimulate oxidative stress by increasing thiobarbituric acid reactive substances and protein carbonyl levels and lowering GSH levels. It also induced mitochondrial dysfunction by decreasing the activity of oxidative phosphorylation (OXPHOS) complexes, tricarboxylic acid cycle (TCA) enzymes, and adenosine triphosphate (ATP) levels [2, 12]. Previous studies support the hypothesis that ROS formation and antioxidant depletion may be central mechanisms for AC‐induced genotoxicity.

AC in urban atmospheres poses a significant exposure risk for humans and leads to various health problems [6]. AC has been implicated in a range of pathological processes across multiple cell types and tissues [1]. It has been found that AC disrupts the function of DNA repair by modifying repair proteins, which increases cell sensitivity to mutagenesis associated with DNA damage [13]. Although AC has been identified as a risk factor for the development of lung, bladder, and colon cancer [13, 14], there are insufficient studies investigating the cytotoxic effects of AC on pancreatic cells. Therefore, the toxicological mechanism of AC‐induced toxicity in human pancreas cells is still not fully understood.

Natural phenolic compounds are phytochemical antioxidants that have been identified in various plant‐based products and foods. These compounds have demonstrated significant potential in offering protective effects against a wide range of diseases, including but not limited to cancer, diabetes, cardiovascular disease, and neurological disorders. Pycnogenol (PYC) is a phenolic compound obtained from the dried bark of the French maritime pine (Pinus pinaster). Due to its potent antioxidant activity, it is widely consumed as a dietary supplement. PYC is composed of a variety of phenolic components, including monomers such as taxifolin, epicatechin, and catechin; concentrated flavonoids, which are grouped as procyanidins and proanthocyanidins; and phenolic acids, including cinnamic acids and certain glycosides. It has been demonstrated that PYC exerts a protective effect against a range of pathologies, including inflammatory diseases, hypertension, diabetes, and obesity. PYC is accepted as a safe bioactive compound according to scientific safety and preclinical toxicology records. As demonstrated in numerous studies, PYC exhibits therapeutic and protective potential against cancer [15]. However, there is a lack of research on its efficacy against AC‐induced damage.

Toxicological effects of AC on human pancreatic cells, particularly in terms of cytotoxicity, genotoxicity, oxidative stress, and apoptosis, have not yet been fully elucidated. Furthermore, the investigation into pancreatic ductal models is comparatively lacking when compared to other cell types in this context. Therefore, AC‐induced toxicity and the potential protective effects of PYC were investigated in detail in human pancreatic ductal cells in the present study.

2. Materials and Methods

2.1. Chemicals

AC (5000 µg/mL in water) was purchased from Restek (Bad Homburg, Germany). Thiazolylblue tetrazolium bromide (MTT), 2´,7´‐dichlorofluorescein diacetate (DCFH‐DA), paraformaldehyde, ethylenediaminetetraacetic acid, Triton X‐100, hydrogen peroxide (H₂O₂), acridine orange (AO) and ethidium bromide (EB) were purchased from Sigma‐Aldrich; Merck KGaA (Darmstadt, Germany). Tissue culture flasks and plates were purchased from Corning. DMEM medium, trypsin/EDTA, fetal bovine serum, phosphate‐buffered saline and penicillin‐streptomycin were purchased from Gibco. Phospho‐histone H2A.X (Ser139) antibody was purchased from Abcam. Alexa Fluor 555 goat anti‐mouse and SYBR green I nucleic acid gel stain were purchased from Thermo Fisher Scientific. Comet slides, lysis solution and low melting agarose were purchased by Trevigen.

2.2. Cell Culture and Cell Viability

hTERT‐HPNE cells, a human pancreatic duct cell line, were maintained in maintained in DMEM medium supplemented with 10%FBS and 1% PS. The cells of with passage number 5 (P5) were maintained in a humidified atmosphere of 5% CO₂ and 95% air at 37°C. Following the seeding of cells in a 96‐well plate at a density of 1 × 10⁴ cells/well, the cells were treated with different concentrations of AC (0, 1, 5, 10, 50, 100, and 500 µM) and/or PYC (0, 5, 10, 25, 50, 100, 250, 500, and 1000 µM) for 24, 48, and 72 h. Because all AC and PYC dilutions were performed using sterile distilled water, sterile distilled water was used as a solvent control. The MTT assay was performed as previously published [16, 17]. Following the determination of the IC50 values, the cells were treated with 25, 50, and 100 μM AC, 100 μM PYC and co‐treated with them for 24, 48, and 72 h for further experiments. A group containing only PYC treatment was included in all assays.

2.3. Alkaline Comet Assay

hTERT‐HPNE cells were seeded in six‐well plates at a density of 10 × 10³ cells/well and then treated with AC (25, 50, and 100 µM) and/or PYC (100 µg/mL) for 24, 48, and 72 h. Positive control cells were treated with 100 μM H₂O₂ for 30 min. Alkaline comet assay was performed according to our previously published study [17]. Comet images were captured using a fluorescence microscope (Leica DM2500, Leica Microsystems, Germany), and the images were analyzed using Comet Score 2.0 software (Tritek Corp, Sumerduck, VA, USA). After 100 cells were scored for each concentration, tail DNA (%), tail moment, and olive tail moment parameters were analyzed to assess DNA damage. Fluorescence comet images were imported into the software, and contrast and brightness were adjusted to clearly define comet heads and tails. After individual comets were manually selected, the software automatically detected tail parameters based on fluorescence intensity and applied background correction.

2.4. gH2AX Immunofluorescence Staining

To detect the formation of DNA double‐strand breaks, gH2AX nuclear foci were analyzed in hTERT‐HPNE cells treated with AC (25, 50, and 100 µM) and/or PYC (100 µg/mL) for 24, 48, and 72 h. 100 μM H₂O₂ was used as a positive control. Immunofluorescence staining was performed as previously published [17]. After slides were mounted, the nuclei of cells were examined under a Leica DM2500 fluorescent microscope at 40X magnification. All gH2AX foci per cell nucleus were counted in 50 cells in each treatment using the ImageJ software. The original color images were converted to 8‐bit grayscale, and then the threshold gray value was manually adjusted using the command image → adjust → threshold. Particle analysis was then performed on the images using analyze → analyze particles, with the options masks and display results enabled. The minimum particle size was optimized by starting from 0 pixels and increasing in increments of 1 pixel until the detected foci most closely matched the original focus image by visual inspection.

2.5. AO/Ethidium Bromide (AO/EB) Staining

hTERT‐HPNE cells were seeded in 24‐well plates at a density of 1 × 10⁵ cells/well and treated with AC (25, 50, and 100 µM) and/or PYC (100 µg/mL) for 24, 48, and 72 h. AO/EB staining was performed as previously published [17, 18]. Images were captured using a fluorescent microscope (Leica DM2500). The calculation of apoptosis rates was performed using the following formula: Apoptosis %= apoptotic cells/total cell count × 100. The images were analyzed, and the percentage of apoptosis was counted in 100 cells using ImageJ software. Fluorescence images were split into red/yellow and green channels (image → color → split channels) to analyze separately, and background fluorescence was minimized. Threshold values were adjusted to selectively identify fluorescent cells in each channel. Binary images were generated, and cells were counted using analyze → analyze particles, with appropriate size parameters set to exclude debris and the exclude on edges option enabled. The numbers of green and red/yellow fluorescent cells were recorded and expressed as cell counts or percentages of total cells.

2.6. JC‐1 Staining

The assessment of mitochondrial membrane potential (MMP) was conducted using a commercially available MMP Assay Kit with JC‐1 (Elabscience, E‐CK‐A301) according to the manufacturer's instructions. hTERT‐HPNE cells were seeded in 24‐well plates at a density of 1 × 10⁵ cells/well and treated with AC (25, 50, and 100 µM) and/or PYC (100 µg/ml) for 72 h. After treatment, the cells were treated with JC‐1 dye (5 µg/ml) for 30 min at 37°C and photographed under a fluorescence microscope (Leica DM2500). The transition from red to green fluorescence after JC‐1 staining reflects a decrease in MMP and apoptosis. 10 μM CCCP (carbonyl cyanide m‐chlorophenylhydrazone) was used as a positive control. We calculated the ratio of red to green fluorescence using ImageJ software. In healthy cells, the levels of MMP are elevated, and they exhibit red fluorescence. In the initial phase of apoptosis, there is a decline in MMP, JC1 exists in the mitochondrial matrix in the form of monomers, and cells produce green fluorescence. The transition from red to green with JC‐1 reflects a decrease in MMP and apoptosis. We calculated the ratio of red to green fluorescence using ImageJ software. Red and green fluorescence channels were separated using image → color → split channels. Background fluorescence was corrected in each channel using process → subtract background. Regions of interest corresponding to individual cells were manually defined and applied consistently to both channels. Mean fluorescence intensities for the red or green channels were quantified using analyze → measure. MMP was expressed as the ratio of red to green fluorescence intensity.

2.7. Determination of ROS With DCFH‐DA

hTERT‐HPNE cells were seeded in 24‐well plates at the density of 1 × 10⁵ cells/well and treated with AC (25, 50, and 100 µM) and/or PYC (100 µg/mL) for 72 h. Following treatment, the cells were incubated with 2 µM DCFH‐DA, an indicator for the determination of ROS, at 37°C in the dark for 15 min [19]. After fluorescence images were captured using a Leica DM2500 fluorescence microscope (Leica Microsystems, Germany), fluorescence intensity was calculated using ImageJ software. Fluorescence images were converted to 8‐bit grayscale. Regions of interest corresponding to individual cells were manually defined. The measurement parameters were established, encompassing the area, mean, integrated density, mean gray value, and limited to threshold. Fluorescence intensities were quantified using analyze → measure. Background fluorescence was determined by selecting areas without cells and measuring them using the same threshold settings. Corrected total cell fluorescence was calculated using the formula: Corrected total cell fluorescence = Integrated density − (Cell area × Mean background fluorescence).

2.8. Glutation and Lipid Peroxidation Assays

GSH content in hTERT‐HPNE cells treated with AC (25, 50, and 100 µM) and/or PYC (100 µg/mL) for 72 h was assayed by a Human GSH, GSH ELISA Kit (BT Lab, EA0021Ge) and general malondialdehyde (MDA), a marker of lipid peroxidation, MDA ELISA Kit (BT Lab, E0017Ge) according to the manufacturer's instructions. The absorbance was measured at 450 nm using a microplate reader (BioTek ELx808). GSH concentrations (µg/mL) MDA concentrations (nmol/mL) in the samples were calculated from the standard curve generated using known standards, and then graphs were prepared from the GSH and MDA values.

2.9. Total Antioxidant Status (TAS) and Total Oxidant Status (TOS)

Following treatment of hTERT‐HPNE cells with AC (25, 50 and 100 µM) and/or PYC (100 µg/ml) for 72 h, total antioxidant status (TAS) and total oxidant status (TOS) were determined by using TAS Colorimetric Assay Kit (E‐BC‐K801‐M) and TOS Colorimetric Assay Kit (E‐BC‐K802‐M) (Elabscience) according to the manufacturer's instructions. The absorbance was measured at 660 nm for TAS and 590 nm for TOS using a microplate reader (BioTek ELx808). TAS values (mmol Trolox Equivalent/L) and TOS (μmol H₂O₂ Equiv./L) were calculated according to the Excel file on the kit's web pages, and then graphs were prepared from the resulting values.

2.10. Statistical Analysis

All experiments were repeated three independent replicate experiments. All data were expressed as the mean ± standard error (SE). The data were analyzed using one‐way ANOVA followed by post hoc test (Tukey's analysis) to compare significant differences between groups. p < 0.05 was considered statistically significant.

3. Results

3.1. Cytotoxicity and Cell Viability

The results of the MTT assay demonstrated that AC caused a concentration‐dependent decrease in cell viability for all treatment times. These decreases were found to be statistically significant at the higher AC concentrations (100 and 500 µM) for 24 and 72 h treatments when compared to the control group (p < 0.05). All tested AC concentrations significantly decreased the cell viability after 72 h treatment when compared to the control group (p < 0.05) (Figure 1). IC₅₀ values were 240, 214, and 151 µM for 24, 48, and 72 h, respectively. IC₅₀ is a practical and widely accepted metric in pharmacology and toxicology. IC₅₀ value is crucial in MTT‐based cytotoxicity evaluation because it can (a): provide a quantitative and standardized measure of toxicity, (b): summarize dose–response relationships, (c): enable comparison across compounds and conditions, and (d): guide experimental design and mechanistic studies. Sub‐IC₅₀ concentrations are used in mechanistic studies such as apoptosis and necrosis [20, 21, 22]. Based on the findings, the highest concentration was selected as 100 µM that below the 72‐h IC₅₀ value for further experiments. The concentrations of 50 µM (half of the highest concentration) and 25 µM (one‐fourth of the highest concentration) were selected as the medium and low concentrations, respectively. The concentration range was chosen to cover sub‐cytotoxic to moderately cytotoxic levels, allowing the evaluation of dose‐dependent effects on cellular redox status, DNA damage, and mitochondrial function.

Figure 1.

Effects of AC on cell viability of hTERT‐HPNE cells after 24, 48, and 72 h treatments. Data are expressed as mean ± SE for n = 3. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

Although some PYC concentrations decreased the cell viability values in a concentration‐independent manner, none of the values were statistically significant compared to the control after 24 and 48 h treatments. All PYC treatments except the lowest concentration (5 µg/mL) significantly decreased the cell viability compared to the control (p < 0.05) (Figure 2). While the cell viability values did not fall below 50% in 24 and 48 h PYC treatments in HPNE cells, the IC50 value was found to be 891 µg/mL in 72 h treatment. While the cell viability values were not below 50% after 24 and 48 h PYC treatments, 891 µM was found to be IC₅₀ value after 72 h treatment.

Figure 2.

Effects of PYC on cell viability of hTERT‐HPNE cells after 24, 48, and 72 h treatments. Data are expressed as mean ± SE for n = 3. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

The selection of PYC concentration for mechanistic assays was guided by both cytotoxicity data and literature evidence. Although the IC₅₀ in hTERT‐HPNE cells after 72 h of treatment was 891 µg/mL, a previous study indicated that PYC can induce DNA damage at concentrations above 100 µg/mL [23], which could confound the assessment of its protective mechanism against AC‐induced oxidative stress. Therefore, the concentration of 100 µg/mL for PYC was chosen for further experiments. We consider that 100 µg/mL represents a physiologically relevant and mechanistically appropriate dose that is high enough to elicit protective cellular responses without introducing direct genotoxicity. The PYC concentration was chosen to balance safety, efficacy, and interpretability, and future studies may explore dose‐response effects to refine optimal protective concentrations.

3.2. Comet Assay

Imaging and quantification of DNA damage after the alkaline comet assay were demonstrated in Figures 3A,B, respectively. PYC treatments did not induce DNA tail (%), tail moment, and olive tail moment. Concentration‐dependent increases were observed in DNA tail (%) after AC treatments. These increases were significant at all AC and AC + PYC treatments after 48 and 72 h treatments compared to the control group (p < 0.05). AC + PYC treatments for 48 and 72 h significantly reduced the DNA tail values compared to the AC treatments alone (p < 0.05). AC and AC + PYC significantly increased tail moment in a concentration‐dependent manner after 48 h and 72 h treatments compared to the control group (p < 0.05). AC + PYC treatments after 72 h significantly reduced the tail moment compared to the AC treatments alone (p < 0.05). However, AC + PYC treatments could not normalize the damage to the same extent as the control group. AC and/or AC + PYC significantly increased olive tail moment at 50 and 100 µM AC after 48 h treatment and at all AC concentrations after 72 h treatment when compared to the control group (p < 0.05). AC + PYC treatments after 72 h treatment significantly reduced the tail moment compared to the AC treatments alone (p < 0.05) (Figure 3B).

Figure 3.

(A) Representative comet images in hTERT‐HPNE cells treated with AC and/or PYC after 72 h. (a) Control, (b) 100 µM PYC, (c) H₂O₂ (100 µM) as positive control, (d) 25 µM AC, (e) 50 µM AC, (f) 100 µM AC, (g) 25 µM AC + 100 µM PYC, (h) 50 µM AC + 100 µM PYC, (i) 100 µM AC + 100 µM. Magnification = ×200; scale bar = 100 μm. (B) DNA damage measured as comet DNA tail (%), tail moment, and olive tail moment in hTERT‐HPNE treated with AC and/or PYC treated with FLUP for 24, 48, and 72 h at indicated concentrations. Data are expressed as mean ± SE for n = 3. The results are represented as mean ± Standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

3.3. gH2AX Foci Formation

Figure 4A shows gH2AX foci (red) in nuclei of hTERT‐HPNE cells treated with AC and/or PYC. All AC treatments significantly increased gH2AX foci in a concentration‐dependent manner compared to the control group (p < 0.05). PYC treatments did not induce the formation of gH2AX foci. AC + PYC treatments also caused significant increases in the mean gH2AX foci number compared to the control group (p < 0.05) except at 25 µM AC + 100 µg/mL PYC after 72 h treatment. Significant decreases in foci formation have been observed at 50 µM AC + 100 µg/mL PYC for 24 h, at all AC + PYC treatments for 48 h, and at the higher concentrations (50 µM AC + 100 µg/mL PYC and 100 µM AC + 100 µg/mL PYC) for 72 h when compared to AC treatments alone (p < 0.05) (Figure 4B). These results may indicate that PYC can prevent or reduce the DNA damage caused by AC.

Figure 4.

(A) Representative fluorescence images of gH2AX foci in hTERT‐HPNE cells after immunofluorescence staining at 400X magnification. First column, DAPI staining; second column, gH2AX foci staining; third column, merged image. (B) Mean gH2AX foci number in hTERT‐HPNE cells treated with AC and/or PYC for 24, 48 and 72 h at indicated concentrations. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

3.4. AO/EB Staining

The effects of AC and/or PYC on apoptosis after AO/EB staining were shown in Figure 5A. PYC did not induce apoptosis in hTERT‐HPNE cells. Although all AC treatments significantly increased the percentage of apoptotic cells after 72 h treatment, significant increases in apoptosis rates were found at the higher AC concentrations (50 and 100 µM AC) after 24 and 48 h treatments when compared to the control group (p < 0.05). Although decreased apoptosis rates were detected in all AC + PYC treatments, significantly increased apoptosis rates were also found at 100 µM AC + 100 µg/mL PYC after all treatment times when compared to the control group (p < 0.05). All AC + PYC treatments after 72 h treatment significantly reduced the percentage of apoptotic cells when compared to AC treatments alone (p < 0.05) (Figure 5B). Based on the result, we can hypothesize that long‐term PYC exposure may reduce the rate of AC‐induced apoptosis.

Figure 5.

(A) Fluorescent micrographs of AO/EB‐stained hTERT‐HPNE cells after 72 h treatment (A: control, B: 25 µM AC, C: 50 µM AC and D: 100 µM AC, E: 100 µM PYC, F: 25 µM AC + 100 µM PYC, G: 50 µM AC + 100 µM PYC, H: 100 µM AC + 100 µM PYC). Magnification = ×200; scale bar = 100 μm. (B) The percentage of apoptosis in hTERT‐HPNE cells treated with AC and/or PYC for 24, 48, and 72 h. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

3.5. MMP

The effects of AC and/or PYC on MMP after JC‐1 staining were shown in Figure 6A. In normal cells, the dye forms red fluorescent aggregates in the mitochondria under conditions of high membrane potential. Conversely, under conditions of mitochondrial membrane depolarization, the dye forms green fluorescent monomers in the apoptotic cells. Significant decreases in MMP were found at 50 and 100 µM AC after 24 and 72 h treatments, and at all AC concentrations after 72 h treatment when compared to the control group (p < 0.05). Significant decreases in red/green fluorescence intensity ratio were detected at 50 µM AC + 100 µg/mL PYC after 24 and 72 h treatments, and at 100 µM AC + 100 µg/mL PYC for all treatment times compared to the control group (p < 0.05). PYC did not decrease the MMP. Although AC + PYC treatments increased the ratio of red/green fluorescence intensity, no significant increase in the MMP was found when compared to the AC treatment alone (p < 0.05) (Figure 6B).

Figure 6.

(A) Fluorescent micrographs of JC‐1 staining representing the mitochondrial membrane depolarization in hTERT‐HPNE cells after 72 h treatment (A: control, B: 100 µM PYC, C: CCCP as positive control, D: 25 µM AC, E: 50 µM AC, F: 100 µM AC, G: 25 µM AC + 100 µM PYC, H: 50 µM AC + 100 µM PYC, I: 100 µM AC + 100 µM PYC). Magnification = ×200; scale bar = 100 μm. (B) Effects of AC on red/green ratio after JC‐1 staining in hTERT‐HPNE cells treated with AC and/or PYC for 24, 48, and 72 h. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

3.6. Oxidative Stress and Antioxidant Markers

As shown in Figure 7A, PYC did not decrease GSH levels. Although AC decreased GSH levels in a concentration‐dependent manner in hTERT‐HPNE cells, significant decreases were determined at 50 µM and 100 µM after 24 and 48 h treatments, and at all AC concentrations after 72 h treatment when compared to the control (p < 0.05). Although AC + PYC treatments increased AC‐induced GSH levels, there were significantly higher GSH levels at 100 µM AC + 100 µg/mL PYC treatment for 48 and 72 h treatments when compared to the control (p < 0.05). Co‐treatments of PYC with AC significantly increased AC‐decreased GSH levels at higher concentrations (50 µM AC + 100 µg/mL PYC after all treatment times and 100 µM AC + 100 µg/mL PYC after 72 h treatment) when compared to AC treatments alone (p < 0.05).

Figure 7.

GSH (µg/mL) (A) and MDA (nmol/mL) (B) levels in hTERT‐HPNE cells treated with AC and/or PYC for 24, 48, and 72 h treatments. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

Significantly increased MDA levels in a concentration dependent manner were found at 50 µM AC after 72 h treatment, and at 100 µM AC for all treatment times when compared to the control cells (p < 0.05). PYC did not increase MDA levels. Although AC + PYC treatments decreased AC‐induced MDA levels, there was a significant increase in MDA levels after 100 µM AC + 100 µg/mL PYC treatment for 72 h when compared to the control (p < 0.05). Co‐treatments of PYC with AC significantly decreased MDA levels after 24 and 72 h treatments when compared to AC treatments alone (p < 0.05) (Figure 7B).

PYC treatment did not alter TOS or TAS levels (Figure 8). AC significantly increased TOS values at all treatments except for the lowest concentration after 24 h treatment compared to the control group (p < 0.05). Although AC + PYC co‐treatments reduced AC‐induced TOS levels, significantly increased TOS levels were found at 50 µM AC + 100 µg/mL PYC for 72 h and at 100 µM AC + 100 µg/mL PYC for all treatment times when compared to the control group (p < 0.05). However, all AC + PYC treatments significantly reduced the TOS levels when compared to AC treatments alone (p < 0.05) (Figure 8A). Significantly decreased TAS values were found after all AC treatments for 48 and 72 h when compared to the control group (p < 0.05). Co‐treatments of PYC with AC increased TAS levels after all treatments. The increased TAS value at 50 µM AC + 100 µg/mL PYC for 72 h was significant when compared to AC treatments alone (p < 0.05) (Figure 8B).

Figure 8.

TOS (A) and TAS (B) levels in hTERT‐HPNE cells treated with AC and/or PYC for 24, 48, and 72 h treatments. The results are represented as mean ± standard error for three independent experiments. Asterisk (*) indicates a significant difference in comparison with negative control group within the same experimental period (p < 0.05), using one‐way ANOVA followed by Tukey's multiple comparison test (n = 3).

4. Discussion

AC exposure caused cytotoxicity in retinal epithelial cells [24]. It reduced cell viability in a time‐ and dose‐dependent manner in human middle ear epithelial cells [25]. AC (20, 40, and 80 μM) induced dose‐dependent alterations in cell morphology, reduced cell viability, and decreased the number of adherent cells in human corneal stromal fibroblasts [9]. Reduced cell viability numbers were found in human embryonic kidney cells (HEK 293AD) treated with AC (20−100 µM) in a dose‐dependent manner [26]. A previous study has shown that AC decreased the cell viability of HUVEC cells [10]. AC increased cytotoxicity and reduced the cell viability of human pancreatic cells in the present study. The decreases in cell viability after AC treatment observed in our study, as well as in previous studies, provide evidence that AC can induce toxicity in human cells.

4.1. Oxidative Stress and ROS Production

Several recent studies showed that AC exposure increased the intracellular ROS levels and oxidative stress in multiple human cell types. AC increased oxidative stress in retinal epithelial cells [24]. It increased ROS levels and decreased GSH levels in HepG2 cells [27]. The increase in the endogenous GSH antioxidant index in lung fibroblasts has been demonstrated to be an effective way against AC‐induced oxidative stress [28]. AC showed toxicity by inducing neuronal injury, oxidative and ER stress, ROS formation, and lipid peroxidation in cochlear nucleus neurons [29]. Significant increases in mitochondrial lipid peroxidation and protein carbonyl levels were observed in liver and brain tissues of rats treated with AC [30]. AC (20, 40, and 80 μM) induced oxidative stress by increasing ROS levels and reducing antioxidant defenses in primary human corneal fibroblasts [9]. Human bronchial epithelial cells (BEAS‐2B) exposed to AC exhibited significant ROS formation accompanied by depleted GSH levels, indicating acute oxidative stress before cell death [3]. Human umbilical vein endothelial cells (HUVECs) showed increased ROS levels and oxidative stress markers after AC treatments. ROS elevation also activated stress signaling pathways such as p38 MAPK and JNK [10]. AC induced oxidative stress in human proximal tubule cells (HK‐2) [31]. Our results are in agreement with the previous findings because of significant increases in ROS levels in human pancreatic cells.

It has been demonstrated that, in addition to suppressing antioxidant capacity, AC enhanced ROS production by activating pro‐oxidant enzymes, promoting lipid oxidation and disrupting mitochondrial respiration. AC‐induced oxidative stress subsequently triggers pathological processes such as inflammation, endoplasmic reticulum stress, apoptosis, and autophagy, thereby amplifying cellular injury [11]. All findings suggest that oxidative stress plays a central role in mediating AC‐induced toxicity and that increases in ROS levels and disturbance of redox homeostasis are primary mechanisms by which AC exerts toxicity in human cells.

4.2. Antioxidant/Oxidant Balance: GSH, MDA, TAS/TOS

AC induced a variety of stress pathways in human hepatocytes, including oxidative stress, decreasing of GSH, ER stress, and mitochondrial dysfunction, which collectively contribute to AC toxicity [32]. AC caused oxidative damage by reducing cellular GSH and triggering the generation of intracellular ROS [1, 33]. AC decreased GSH levels in human neuroblastoma SH‐SY5Y cells [28], primary human corneal stromal fibroblasts cells [9], and in mouse pancreatic β‐cell MIN6 cells [34]. Treatment of rats with AC was found to result in a significant increase in MDA levels in the vestibulocochlear nerve tissue, while the GSH level was reduced in comparison to that obtained in the healthy groups. These results demonstrated that the oxidant‐antioxidant balance in the AC‐treated group shifted towards oxidants [35]. AC can react quickly with biological nucleophilic targets, such as GSH, because of its high electrophilicity [36]. The reaction of AC with GSH can decrease the intracellular GSH and form glutathionyl propionaldehyde, which then undergoes an enzymic catalytic reaction to generate ROS [3, 36]. AC decreased GSH levels and increased ROS levels and cell death markers in BEAS‐2B and HUVEC cells [3, 10].

The primary source of AC exposure is attributed to the lipid oxidation of polyunsaturated fatty acids during the processing and storage of foods, as well as from environmental sources such as cigarette smoke. It exerts harmful effects in various cell types and tissues primarily by inducing oxidative stress. It acts both as a product and as a propagator of lipid peroxidation, further amplifying oxidative damage [11]. Although direct measurements of TAS and TOS and indirect markers like lipid peroxidation products such as MDA are rare in human cell culture models, some studies reported that GSH depletion after AC exposure, indicating consumption of intracellular antioxidants. AC induces oxidative stress primarily by depleting GSH and forming adducts with antioxidants and redox‐regulating enzymes, thereby impairing endogenous antioxidant defenses [11, 37]. Rapidly depleting GSH by direct binding and increasing ROS can make the cells more vulnerable to oxidative damage. Increases in ROS can cause the weakening of the cell's antioxidant defenses by driving lipid peroxidation and producing markers like MDA that indicate membrane damage. In vivo changes in TAS and TOS also showed that AC shifts the redox balance toward a pro‐oxidant state, leading to oxidative stress and cellular injury [35]. AC also decreased GSH and TAS levels and increased MDA, TOS and ROS levels in our study. All these patterns reflect a shift toward a pro‐oxidant state where antioxidant capacity is overwhelmed by ROS generation.

4.3. DNA Damage: Single and DSBs

When the ROS level is excessive, it can damage cells and biomolecules, such as DNA [38]. AC can react readily with cellular nucleophilic components such as proteins and DNA because of its high electrophilicity [11, 32]. DNA adducts and DNA‐protein crosslinks are formed by AC itself, which hinders replication and repair processes, thereby exacerbating genomic instability [1]. Molecular docking results also showed that AC can bind to DNA in a spontaneous process [3].

The presence of excessive oxidative stress has been proven to directly result in DNA damage within human cells. Some studies provide evidence that AC can cause nuclear and mitochondrial DNA damage. AC‐induced ROS can cause a range of genotoxic lesions, including single‐strand breaks (SSBs) and DSBs, as well as oxidative base modifications such as 8‐oxo‐7,8‐dihydro‐2′‐deoxyguanosine (8‐OHdG). It has been stated that AC‐induced DNA damage can be related to oxidative stress because it induced 8‐oxodGuo formation, DNA migration, DNA strand breaks and DNA‐protein crosslinks in HepG2 cells [27]. AC can induce DNA damage in primary human small airway epithelial cells by increasing the percentage of tail DNA in the comet assay [5]. AC increased oxidative DNA lesions and strand breaks in BEAS‐2B cells by increasing the levels of 8‐OHdG and DNA damage parameters such as the percentage of tail DNA and olive tail moment measured by comet assay. Additionally, increased γ‐H2AX levels were found in BEAS‐2B cells. AC exposure activated DDR, including ATM/ATR/Chk signaling, cell cycle checkpoints (e.g., G2/M arrest), and DNA lesions to downstream effects [3]. AC also increased DNA damage response in HUVEC [10]. Our results showed that AC can increase DNA single‐ and DSBs by increasing comet parameters and the formation of gH2AX foci in human pancreas cells. All data showed that AC can induce both single‐ and DSBs along with oxidative nucleotide modifications in human cells.

AC has been linked to an increase in DNA damage, which could potentially lead to colon cancer [14]. AC has been found to contribute to bladder carcinogenesis by inducing DNA damage and inhibiting repair. It has also been shown to induce cisplatin resistance and tumour progression in both non‐muscle‐invasive and muscle‐invasive bladder cancer cells [13]. However, there are very limited studies investigating the cytotoxic effect of AC on pancreatic cells, despite it being identified as a risk factor for lung, bladder, and colon cancer. Our results demonstrate that AC can induce DNA strand breaks in human pancreatic cells, which play a significant role in cancer development. In agreement with previous studies using different human cell types, our results demonstrated for the first time that AC can trigger DNA damage in human pancreatic cells by increasing DNA strand breaks compared to a control group.

4.4. Mitochondrial Dysfunction and Membrane Potential

An increasing number of studies are focusing on the importance of mitochondria in explaining disease occurrence and therapeutic targets [39]. Some studies have demonstrated that mitochondria are a key target of AC‐induced cytotoxicity because they can disrupt mitochondrial homeostasis, leading to energy depletion and structural impairment. Isolated mitochondria from rat livers showed that AC has toxic effects on mitochondrial respiratory enzymes, leading to mitochondrial dysfunction and hepatotoxicity [40]. AC (20, 40, and 80 μM) significantly decreased the MMP levels and destroyed MMP, which are the earliest phenomena in the mitochondrial apoptosis [3, 41]. Exposure to AC has been reported to cause oxidative mitochondrial damage in retinal pigment epithelial cells [42]. AC caused mitochondrial dysfunction in human hepatocytes [32]. AC caused mtDNA damage, reduced mtDNA copy number, decreased MMP, lowered ATP production, and impaired mitochondrial respiration in human lung cell lines (A549 epithelial cells and MRC‐5 fibroblasts). These changes compromise bioenergetics and promote intrinsic apoptotic signaling. It has been documented that AC‐induced mitochondrial fission and DNA damage can trigger apoptosis. All alterations impaired respiration, ultimately activating caspase‐dependent apoptosis [6]. AC reduced MMP and mitochondrial ATP production in cochlear nucleus neurons [29]. There were significant decreases in mitochondrial GSH levels and in the levels of antioxidant enzymes, electron transport chain enzymes (such as complexes I and IV), TCA enzymes, and ATP levels [2, 12]. Mitochondrial apoptosis pathway involved in apoptosis, MMP and ATP content of BEAS‐2B cells decreased in BEAS‐2B cells [3]. It reduced MMP and ATP content, indicating compromised bioenergetics in HUVEC cells [10]. These mitochondrial effects coincided with increased ROS production, suggesting that mitochondrial dysfunction both contributes to and is exacerbated by oxidative stress. Consistent with these findings, our results also support the hypothesis that AC can reduce MMP and lead to mitochondrial dysfunction.

4.5. Apoptosis and Cell Death Pathways

Previous studies have reported that there is a link between AC exposure and activation of programmed cell death. AC is capable of readily traversing cell membranes and tissues due to its solubility in water and alcohol. Consequently, elevated AC concentrations, resulting from lipid peroxidation, can disseminate from the initial cell of origin, potentially causing damage or death to adjacent cells [32]. AC increased the apoptosis rate approximately threefold in smooth muscle cells [25]. It has been reported that AC (25–40 µmol/L) induced ER stress and cell death in rat pancreatic acinar cells (AR42J) in a dose‐dependent manner [43]. It has been reported that AC induced apoptosis in cochlear nucleus neurons [28]. High apoptosis rate was confirmed by TUNEL and caspase assays alongside ROS increase and GSH depletion in human corneal fibroblasts [9]. After AC treatment, the expression of Bax/Bcl‐2 and cleaved caspase‐3 increased, the JNK signaling pathway was activated, and the apoptosis rate increased in BEAS‐2B cells [3]. AC‐treated HUVEC cells exhibited upregulated Bax/Bcl‐2 and cleaved caspase‐3, confirming involvement of the mitochondrial apoptotic pathway [10]. AC‐induced cell death was found in human proximal tubule cells (HK‐2), resulting in kidney disease [31]. Some studies have shown that AC induced inflammation and mitochondria‐dependent apoptosis as a major pathway of cytotoxicity [11]. Previous apoptosis results after AC treatment were also associated with oxidative stress and DNA damage signaling [10]. Similarly, in this study, AC induced both oxidative stress and apoptosis in human pancreatic cells.

4.6. Summary of AC Toxicity

The concentration‐dependent depletion of GSH levels suggests that AC induces oxidative stress in hTERT‐HPNE cells. This reduction indicates increased consumption of intracellular antioxidants in response to elevated ROS levels, ultimately reflecting an imbalance in cellular redox homeostasis. An increase in GSH consumption means that GSH synthesis cannot keep up, resulting in a drop in GSH levels. In other words, a decrease in GSH indicates that the cell's antioxidant defense system is being depleted. This indicates that the redox balance is disrupted, making DNA, proteins, and lipids vulnerable to oxidative damage [44, 45]. Elevated oxidative stress markers (high TOS and MDA, low GSH and TAS) are associated with increased oxidative DNA damage. Increased TOS together with decreased TAS are used as indicators of oxidative stress in biological systems, and this imbalance is often associated with pathological processes involving apoptosis and cellular damage [46]. GSH depletion has been mechanistically linked to the initiation of apoptotic signaling because it enhances ROS accumulation and mitochondrial oxidative damage. Cells with low GSH are more susceptible to MMP loss and activation of mitochondrial apoptotic signaling. Previous studies showed that early disruption of GSH redox status caused alterations in mitochondrial membrane permeability and release of apoptogenic factors like cytochrome c, leading to caspase activation and apoptosis [47, 48]. Elevated MDA reflects increased lipid peroxidation caused by ROS, which implicates oxidative damage to cellular membranes, including the mitochondrial membrane. This damage can compromise mitochondrial integrity, contributing to loss of MMP and the downstream apoptotic cascade [49]. An increase in TOS and a decrease in TAS indicate a net shift toward oxidative conditions, meaning that ROS generation overwhelms antioxidant defense. This imbalance is a hallmark of oxidative stress that promotes DNA damage, mitochondrial dysfunction, and apoptosis [50]. Loss of MMPl is a key early event in intrinsic apoptosis and is highly sensitive to oxidative stress. When antioxidants such as GSH are depleted and oxidant burden increases (TOS, MDA), mitochondria produce more ROS and undergo depolarization, which releases pro‐apoptotic factors into the cytosol [45].

The apoptosis findings are consistent with and strongly supported by the DNA damage results obtained from the comet assay and gH2AX foci formation. Our DNA damage results demonstrated increases in DNA strand breaks in AC‐treated cells, indicating genotoxic stress. Such DNA damage is a well‐established trigger of apoptotic pathways, particularly through activation of DNA damage response mechanisms. In line with this, the observed increase in apoptotic cell populations suggests that the extent of DNA damage reached a threshold sufficient to initiate programmed cell death. This correlation indicates that apoptosis is likely a downstream consequence of the induced DNA damage rather than an independent event. Moreover, the dose‐dependent increase observed in comet assay parameters and apoptotic markers further supports a mechanistic link between DNA damage and apoptosis [51, 52]. Together, these findings suggest that the cytotoxic effect of the treatment is mediated, at least in part, through DNA damage‐induced apoptosis.

4.7. PYC

PYC is a defined mixture of polyphenols, primarily procyanidins, catechin, taxifolin, and a minor quantity of phenolic acids. It has exhibited several biological activities, especially antioxidant properties in vitro and a variety of biomodulating activities in vivo [53]. The polyphenolic constituents of PYC can act as potent free‐radical scavengers, directly neutralize ROS such as superoxide and hydroxyl radicals, and thereby reduce oxidative damage to cellular macromolecules, including DNA. PYC has been shown to enhance endogenous antioxidant defenses by increasing intracellular GSH levels and upregulating the activity of antioxidant enzymes such as SOD and catalase, promoting a more robust redox homeostasis capable of counteracting excessive ROS production [11, 54].

The antioxidant effects of PYC have been linked to the upregulation of nuclear factor erythroid 2–related factor 2 (Nrf2) gene expression, indicating the involvement of both antioxidant and anti‐inflammatory mechanisms [55]. PYC activates Nrf2‐dependent transcription by strengthening antioxidant defenses, including GSH and superoxide dismutase (SOD), while concurrently inhibiting ROS‐sensitive NF‐κB signaling pathways, thereby alleviating oxidative stress and related pro‐inflammatory responses [56]. Furthermore, it has been shown to significantly reduce oxidative stress and modulate the overexpression of Nrf2 and aryl hydrocarbon receptor (AHR), both of which are critical regulators of oxidative stress and detoxification processes [57].

Beyond the direct induction of apoptosis, PYC modulates key molecular pathways involved in cancer cell survival and migration. In MCF‐7 breast cancer cells, PYC exhibits pronounced pro‐apoptotic and anti‐migratory effects, as evidenced by decreased cell viability, chromatin condensation, and the downregulation of genes such as B‐cell lymphoma 2 (Bcl‐2), vascular endothelial growth factor (VEGF)/fibroblast growth factor (FGF), and p53. These findings indicate that the modulation of VEGF‐related signaling pathways contributes significantly to the anticancer activity of PYC [58]. Moreover, PYC demonstrates anticancer potential through multiple complementary mechanisms, including the suppression of Bcl‐2 expression [59], stabilization of the pro‐apoptotic Bak protein [60], and the enhancement of both caspase‐independent [61] and caspase‐3–dependent apoptotic pathways [60].

Decreased AC‐induced ROS production, NOX activation and GSH depletion were reported in human neuroblastoma cells (SH‐SY5Y) treated with PYC (50 μg/mL) for 1 h followed by AC (5–25 μM) for 24 h [62]. The protective effects of PYC against AC‐induced toxicity in SH‐SY5Y cells have been demonstrated through the scavenging of free radicals and the inhibition of the assembly of NADPH oxidase subunits. PYC can decrease lipid peroxidation and inhibit ROS‐induced DNA strand breaks, indicating a direct antigenotoxic effect. Moreover, in vivo studies demonstrated that PYC ameliorated DNA damage and restored antioxidant enzyme activities in tissues subjected to oxidative insults. These combined mechanisms—radical scavenging, enhancement of antioxidant capacity, and inhibition of oxidative DNA damage—support the ability of PYC to attenuate oxidative stress and reduce genotoxicity in biological systems [54]. All results support our findings in which PYC decreased the AC‐induced DNA damage, ROS formation, oxidative stress, MMP and apoptosis in human pancreas cells.

5. Conclusions

In summary, our results indicate that AC can disrupt the oxidant–antioxidant balance by increasing ROS, TOS and other oxidative biomarkers, while reducing GSH and TAS levels. This imbalance can trigger DNA damage and mitochondrial dysfunction, eventually activating apoptotic mechanisms in human pancreas cells. This multifaceted toxicity emphasized the importance of AC as a mediator of cellular injury and disease pathogenesis, with implications for the susceptibility of pancreatic cells, where similar oxidative and genotoxic mechanisms are expected to play a key role. The findings can also provide evidence that AC may contribute to the development of pancreatic carcinomas. However, further epidemiological and mechanistic human‐based studies are required to clarify the relationship between AC exposure and pancreatic carcinoma risk.

Based on our findings, it can also be said that PYC has antioxidative effects, prevents oxidative stress, DNA and mitochondrial damage, and apoptosis in AC‐induced human pancreas cells. Therefore, PYC may be helpful for the prevention of AC‐induced toxicity or AC‐induced pancreatic disease in humans.

Author Contributions

Zülal Atlı Şekeroğlu: project administration, resources, supervision, funding acquisition; writing – original draft, writing – review and editing. Vedat Şekeroğlu: conceptualization, writing – original draft, writing – review and editing, software. Şükrüye Kamanlı: Investigation, validation, methodology. Seval Kontaş Yedier: formal analysis, software, data curation, visualization. Birsen Aydın: writing – review and editing, software, methodology, visualization, conceptualization. Ceren Börçek Kasurka: investigation, methodology, formal analysis, supervision; conceptualization.

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.