Impact of environmental microplastic exposure on HepG2 cells: unraveling proliferation, mitochondrial dynamics and autophagy activation

Abstract

The rise of microplastic (MPs) pollution presents a pressing environmental issue, raising concerns about its potential health impacts on human populations. Given the critical role of the liver in detoxification and metabolism, understanding the effects of MPs on the human hepatoma cell line HepG2 cells is essential for comprehensively assessing the dangers associated with MPs pollution to human health. Until now, the assessment of the harmful impact of polyethylene (PE) and polyethylene terephthalate (PET) on HepG2 has been incomplete and lacks certain essential data points. In this particular setting, we examined parameters such as cell viability, oxidative stress, mtDNA integrity, mitochondrial membrane potential, and autophagy in HepG2 cells exposed for 72 h to PET and PE at a concentration of 10 µg/mL. Our data revealed that exposure of HepG2 to MPs causes an increase in cell viability accompanied by a heightened ROS and altered mitochondrial function, as revealed by decreased mtDNA integrity and membrane potential. In addition, results demonstrated that exposure to PET and PE activated autophagic events, as suggested by the increased levels of the specific markers LC3 and p62. This last point was further confirmed using bafilomycin, a specific blocker that hinders the merging of autophagosomes and lysosomes, thereby blocking autophagic degradation processes. Given the increasing evidence of food chain MPs contamination and its possible harmful effects, our data should be carefully considered.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12989-025-00632-x.

Introduction

The global plastic production from 1950 to 2050 is expected to rise from 9.2 billion tons in 2017 to 34 billion tons [1]. As a consequence, microplastics (MPs) pollution is now recognized as one of the four major global environmental issues, running in tandem with concerns such as ozone depletion, ocean acidification, and global climate change [2]. Therefore, there has been a growing focus from scientists and the public on MPs pollution in the last decade. The degradation of plastic caused by UV radiation leads to its fragmentation into minuscule particles known as MPs [3]. As per the European Food Safety Authority, MPs particles are typically characterized as small plastic pieces falling within a size range of 100 nanometers to 5 millimeters [4, 5]. It is crucial to understand that MPs enter the environment through both primary and secondary sources [6, 7]. Indeed, primary MPs are initially produced to be microscopic size, commonly for incorporation into cosmetic items like microbeads [8]. While, secondary MPs find their way into the environment when larger plastic items undergo fragmentation caused by exposure to UV light, wave action, abrasion, freezing, and wind [3, 9, 10]. It is crucial to emphasize that the issue of MPs pollution, a global concern, raises significant apprehensions regarding its impact on both the environment and human health [11]. This is particularly evident as MPs have been detected in bottled water [12, 13], sugar [14], mussels [15, 16], seafood like commercial fish [15, 17], also in salt [18], honey, beer and were ultimately found in human stool [19, 20]. In fact, MPs pose potential risks to human health through both physical and chemical pathways. Human exposure to MPs primarily occurs through the consumption of food containing these particles [21], breathing in MPs suspended in the air, and skin exposure to particles found in various items, textiles, or dust [3, 22]. The inhalation or ingestion of MPs has been linked to potential human pathologies, as indicated by various studies [23, 24]. Furthermore, MPs within the small micron range possess the capability to penetrate bodily barriers and cellular membranes, possibly inducing molecular impacts [25–27]. To date, considerable knowledge gaps remain potential hazards posed by MPs [4, 28]. Considering that liver plays a major role in xenobiotics detoxification, and for this, it represents one of the first tissue that are hit by contaminants, it is necessary to study the impact of polyethylene (PE; <2.6 µm) and polyethylene terephthalate (PET;<1 µm and< 2.6 µm) on the human hepatoma cell line (HepG2), focusing on the associated oxidative cellular response, cell viability, mitochondrial membrane potential (MMP), mitochondrial DNA (mtDNA) integrity, and autophagy. The outcome of this research may offer insights into the presence of PE and PET in HepG2 cells, contributing valuable information to comprehend their potential implications for human health.

Materials and methods

Cell culture

HepG2 Cells were obtained from the American Type Culture Collection (ATCC, HB-8065). Cell proliferation was examined in high glucose conditions; therefore, the cells were grown in DMEM (DMEM; Gibco, C11995500BT). Following this, the HepG2 cells were supplemented with 10% FBS, 1% L-glutamine, 100 µg/mL streptomycin and 100 U/mL penicillin. HepG2 were incubated in a humidified setting with 95% air and 5% CO2 at 37 °C.

HepG2 treatment with MPs

As reported by [29], we prepared MPs suspension and counting was based on the protocol described by [12, 21]. Briefly, Three types of microplastics (MPs) were utilized in this study: polyethylene (PE; <2.6 µm) and two fractions of polyethylene terephthalate (PET), designated as PET 1 (<1 µm) and PET 2.6 (<2.6 µm). The MPs were produced from uncolored water bottles, with polymer composition verified by Fourier-transform infrared (FTIR) spectroscopy (Thermo Scientific, MS, USA). Around 10 g of MPs material was cryogenically fractured in liquid nitrogen and mechanically milled using a swing mill to generate irregular particles. These were initially filtered through a 30 µm mesh, suspended in 100 ml of Milli-Q water, and subsequently fractionated by sequential filtration through cellulose nitrate membranes (Ø = 47 mm; pore sizes: 2.67 µm and 1 µm; Healthcare Life Sciences, UK) to obtain PET particles in two size distributions (<2.6 µm and <1 µm). PE MPs (<2.6 µm) were processed following the same method. The morphology and elemental composition of the MPs were characterized using scanning electron microscopy (SEM; Stereoscan 360, Cambridge Instruments, UK) coupled with energy-dispersive X-ray spectroscopy (SEM-EDX, Inca software). Particle concentrations were calculated and expressed in µg/L, based on polymer density, following the approach of Zuccarello et al [12]. Prior to cellular exposure, all MPs suspensions were sterilized by autoclaving for 40 minutes. HepG2 cells were then exposed to each MPs type (PET 1, PET 2.6, and PE) for a duration of 72 hours under standard culture conditions.

The MPs suspension in stock solution were as following:

• *PET (< 1 μm (PET 1)): 4.46 E⁺⁰⁴ particles/ml, mean radius 0.94 μm

• *PET (1–2.6 μm (PET2)): 3.57 E⁺⁰⁴ particles/ml, mean radius 1.12 μm

• *PE (1–2.6µm (PE2)): 2.90 E⁺⁰⁴ particles/ml, mean radius 1.18 μm

Cell Counting Kit-8 (CCK8) assay

Cell proliferation and viability were assessed using the CCK-8 colorimetric assay (Dojindo Molecular Technologies, CK04) according to the manufacturer’s instructions. Briefly, 1.500 cells were plated in 96- well plates and treated with microplastics (MPs) for 72 h. After treatment, 10 μL of CCK-8 solution was added to each well, and the plates were incubated for 24, 48, and 72 hours at 37 °C. The absorbance at 450 nm was measured using a microplate reader (Thermo Scientific, Varioskan Flash). To ensure reliable results, the experiment was performed with three biological replicates, where each biological replicate consisted of independent cell cultures. Each biological replicate included three technical replicates, with three wells per condition. The experiment was independently repeated eight times to confirm reproducibility.

Reactive oxygen species (ROS) detection

Intracellular ROS levels were assessed through both the thiobarbituric acid reactive substances (TBARS) and the 2,7-dichlorofluorescin diacetate (DCFH-DA) assays (Sigma Aldrich, 35845-1G).

TBARS levels assessment

The cell lysate was mixed with 0.5 mL of 20% acetic acid at pH 3.5 and 0.5 mL of a 0.78% aqueous solution of thiobarbituric acid (Sigma-Aldrich, T5500). Following an incubation period of 45 min at 95 °C, samples underwent centrifugation at 4000× g for 5 min. The supernatants were collected, and TBARS were measured via spectrophotometry at 532 nm, as described by Lama et al. [30]. Results were expressed as TBARS concentration per microgram (µM/µg) of extracted protein. Each measurement was performed three times.

DCF-DA assay

The production of intracellular reactive oxygen species (ROS) was assessed using the fluorogenic probe 2′,7′- dichlorodihydrofluorescein diacetate (DCFH-DA; EuroClone S.p.A, ECB4053). Upon cellular uptake and deacetylation, DCFH-DA is oxidized by ROS into the highly fluorescent compound 2′,7′-dichlorofluorescein (DCF). HepG2 cells were incubated with 2 μM DCFH-DA in phosphate-buffered saline (PBS) containing 0.1% Pluronic F-127 (Sigma-Aldrich, 35845-1G) for 30 min at room temperature (RT) in the dark. After incubation, cells were washed twice with PBS to remove excess dye. Fluorescence intensity was immediately analyzed using a Guava® easyCyte™ flow cytometer (MilliporeSigma), and data were processed using the easyCyte™ software. For each condition, three independent biological replicates were performed, and for each biological replicate, three technical replicates were acquired and analyzed.

Assessing mtDNA integration: methodology and analysis

As reported by Kovalenko et al. [31], the integrity of mtDNA was evaluated. Briefly, mtDNA isolation was carried out utilizing the Mitochondrial DNA Isolation Kit (Abcam, ab65321/K280-50). Following this, 25 ng of DNA was employed for every PCR reaction. PCR assays were performed using a Veriti thermal cycler (Applied Biosystems, 9912), adhering to the manufacturer’s guidelines, and utilizing MegaFi™ Pro Fidelity DNA Polymerase (ABM, G886). To ensure precision and reproducibility, a minimum of three replicates were performed for each RT-PCR reaction. Qualitative assessment of DNA levels was performed with the Gel Doc UV System from Bio-Rad, while densitometric analysis of PCR product levels was conducted using the Molecular Analyst software, which is compatible with the Gel Doc 1000 system (Bio-Rad, CA, USA).

Assessing mitochondrial condition using JC1 dye

Mitochondrial condition is commonly assessed using JC-1 dye, which can permeate cell membranes. In our investigation, we employed a fluorescein-conjugated JC-1 kit (ThermoFisher, M34152) and measured fluorescence using a Guava easyCyte cytometer (ThermoFisher, Waltham, MA, USA) following the provided protocol. JC-1 dye displays distinct accumulation patterns within mitochondria based on their membrane potential. When in its monomeric state, JC-1 emits green fluorescence at approximately 529 nm. However, upon forming a red fluorescent JC-1 aggregates, a concentration-dependent process, the emission shifts to around 590 nm. Consequently, a decrease in the red-to-green fluorescence intensity ratio signifies a loss of MMP, offering valuable insights into mitochondrial health and function. Three biological replicates were conducted, each with three technical replicates, to ensure the reproducibility and precision of the results.

Cyto-ID autophagy detection kit

The assessment of autophagic vacuoles and monitoring of autophagic flux in living cells were conducted utilizing the Cyto-ID® Autophagy Detection Kit (Enzo Life Sciences, ENZ51031). This kit uses a cationic amphiphilic dye that selectively marks autophagic vacuoles. HepG2 were cultured on coverslips, treated with 100 µL of Microscopy Dual Detection Reagent adhering to the guidelines provided by the manufacturer, then HepG2 were incubated at 37 °C for a duration of 30 min. Subsequently, HepG2 were rinsed with 100 µL of 1X Assay Buffer, fixed with 4% Formaldehyde for 20 min, and subjected to three additional washes with 1X Assay Buffer. Wide-field fluorescence analysis was performed, and the percentage of Cyto-ID-positive cells was established in accordance with the guidelines provided by the manufacturer. To ensure the reliability and accuracy of the results, three biological replicates were conducted, each with three technical replicates.

Protein extraction

Following the removal of the culture medium, adherent cells were washed with PBS (EuroClone S.p.A, ECB4053). HepG2 were subsequently lysed using a specialized buffer ([1% Nonidet-P40, 0.1% sodium dodecyl sulfate, 0.5% sodium deoxycholate] in PBS) containing protease inhibitors (sodium orthovanadate, Sodium Fluoride, Phenylmethylsulfonyl fluoride, and Roche cocktail). This process involved scraping the cells, carefully resuspending them, then centrifugation at maximum speed in a cooling centrifuge (Centrifuge Eppendorf Italia, Milan, Italy) for 5 min. The supernatant was subjected to a Bradford assay to quantify the protein concentration. All reagents were sourced from Sigma-Aldrich.

Western blot

A quantity of 40 µg of proteins was subjected to electrophoresis on 9% and 15% SDS polyacrylamide gels, followed by transfer onto polyvinylidene difluoride membranes (Amersham Pharmacia Biotech, GE10600023) at 280 mA for 2.5 h at 4 °C. Prior to protein extraction, cells were incubated for 2 h with 100 nM Bafilomycin A1 (an inhibitor of lysosomal degradation).A blocking solution (5% milk [Sigma-Aldrich, M7409] in Tris-buffered saline) with 0.25% Tween X20 (Sigma Aldrich Corp, P9416) was applied to the filters for a duration of 3 h before the incubation with the primary antibodies (SM Table 1): anti-LC3, antiβ-actin, anti- p62 antibodies were left overnight at 4 °C. Following three washes with T-TBS, the filters were then incubated with the suitable secondary antibody (SM Table 1) in the identical blocking solution. Following this, the filters were subjected to three more washes in T-TBS, and the ECL-western blot analysis detection system from Amersham Pharmacia Biotech was employed to reveal the immunocomplexes. The intensity of the β-actin band was utilized for the normalization of protein levels. Quantification of band density was achieved through the utilization of ImageJ software.

Immunofluorescence assay

Cells were initially seeded on a coverslip in six-well plates and subsequently treated. Following treatment with MPs, HepG2 were treated with a 4% formaldehyde solution for 15 min at ambient temperature to fix them. Permeabilization was carried out using PBS (pH 7.4) supplemented with 0.3% Triton-X-100 (Sigma-Aldrich, T8787) for 10 min. To block non-specific binding sites, a solution consisting of PBS with 0.1% Triton and normal goat serum (Sigma-Aldrich, NS02L) (diluted 1:5) was applied. Later, HepG2 cells were exposed to primary antibodies (SM Table 1) overnight at 4 °C. After three PBS washes, the suitable secondary antibody (SM Table 1), diluted in the blocking solution, was added for 1 h at room temperature. Nuclear staining was achieved using a DAPI (ABCAM, ab104139), and micrographs were taken using a fluorescence microscope (Leica, Germany). The mean for pixel intensity was quantified using ImageJ software. To ensure the robustness and reproducibility of the results, three independent biological replicates were performed, and 5 to 10 random fields were analyzed per replicate.

Statistical analysis

Statistical analysis was conducted using GraphPad Prism 5.01 software (GraphPad, CA, USA). The data were presented as mean ± standard error (SEM). To compare statistical differences, one-way analysis of variance (ANOVA) was performed, followed by Tukey’s test. A p-value below 0.05 was considered to indicate a statistically significant distinction.

Results

MPs increase cell proliferation of HepG2 cells

HepG2 cell cultures were subjected to two distinct types of MPs, PET with particle sizes less than 1 µm and 2.6 µm, as well as PE with particle sizes less than 2.6 µm. The exposure duration ranged from 24 to 72 h, with MPs concentrations varying from 10 µg/ml to 70 µg/ml. The obtained data revealed concentration-dependent responses, yielding Minimum Active Concentration (MAC values) of 50.49 µg/ml for PET (2.6 µm) (Fig. 1B), 40.48 µg/ml for PET 1 µm (Fig. 1A) and 40.07 µg/ml for PE (Fig. 1C). Subsequent assessment of cell viability using the CCK-8 assay confirmed a significant increase in the proliferation of HepG2 cells upon exposure to both PET1 (Fig. 1D), PET 2.6 µm (Fig. 1E) and PE (Fig. 1F). These findings underscore the noteworthy impact of MPs in enhancing the proliferative activity of HepG2 cells. Following the CCK-8 assay, all subsequent experiments were conducted using a concentration of 10 µg/mL for each MPs type, unless otherwise specified.

Fig. 1.

MPs (PET 1 µm, PET 2,6 µm and PE 2,6 µm) exposure increase cell viability of HepG2 cell line. A Standard curves and IC50 values of PET 1 µm for HepG2 cell line. B Standard curves and IC50 values of PET 2.6 µm for HepG2 cell line. C Standard curves and IC50 values of PE 2.6 µm for HepG2 cell line. Calculation of IC50 values in Graph Pad Prism. Fraction of alive cells [%] is provided on the vertical axis and the log (concentration) on the horizontal axis. The IC50 is the concentration at which the curve passes through the 50% inhibition level. D, E and F Effect of MPs on cell viability. HepG2 cells were treated with different concentrations of PET and PE. CCK8 assay was performed to measure cell viability

MPs contribute to heightened oxidative stress, resulting in an elevated generation of ROS

ROS accumulation in HepG2 cells treated with MPs was assessed using both the lipid peroxidation test and the DCFH-DA dye. As demonstrated in (Fig. 2), when cells were exposed to MPs at 10 µg/mL for 72 h, there was a significant increase in malondialdehyde formation (Fig. 2A), which is a biomarker for lipid peroxidation, compared to control cells (p<0.05, p<0.01). Furthermore, the fluorescence levels of DCFH-DA in HepG2 cells treated with MPs (p<0.05, p<0.01) exhibited a significant elevation (Fig. 2B), as quantified by the Guava® easyCyte™ flow cytometer. These findings suggest that MPs induce an overproduction of ROS.

Fig. 2.

Influence of MPs (PET 1 µm, PET 2,6 µm and PE 2,6 µm) exposure on ROS generation. A Histogram showing TBARS levels of HepG2 cells exposed to MPs. B Histogram showing the percentage of DCF-DA positive cells. All values are expressed as means ± standard deviation (*p < 0.05, ** p < 0.01). Statistical significance was evaluated by ANOVA (at least p < 0.05) followed by Tukey test for multigroup comparison. All the experiments were performed in triplicate

MPs impact on mtDNA integrity and MMP

We assessed DNA integrity through PCR, detecting both long and short amplification of mtDNA. Then the DNA integration ratio (short amplification/long amplification) was measured. Our results (Fig. 3A) demonstrated a reduction in mtDNA integrity in HepG2 cells treated with PET 1 (p<0,01), PET 2.6 (p<0,001) and PE (p<0,05) compared to control. These findings underscore the comprehensive impact of MPs on cellular physiology, encompassing mitochondrial dysfunction, and compromised mtDNA integrity.

Fig. 3.

MPs exposure altered mitochondrial function and mtDNA copy number. A Histogram of Mitochondrial DNA integrity B Histogram showing the percentage of dysfunctional mitochondria C The plot of JC1 from flow cytometry on mitochondrial membrane potential. All Data are expressed as means ± SD (*p < 0.05, ** p < 0.01, ***p < 0.001). Data are representative of three independent measurements

Our study delved into the detrimental effects of microplastics (MPs) on cellular dynamics, particularly focusing on mitochondrial function. To assess the MMP in HepG2 cells exposed to PET (1 µm, 2.6 µm) and PE (2.6 µm), JC-1, a cationic probe, was utilized. JC-1 aggregates in mitochondria due to the electronegative environment inside, emitting a red fluorescence under normal conditions. However, in conditions of mitochondrial membrane depolarization (low MMP), JC-1 exists in a monomeric state, causing it to emit green fluorescence (Fig. 3C). Dot plot analysis showed a notable change in green fluorescence intensity after 72 h of MPs exposure compared to untreated HepG2 cells. Specifically, the dot plot analysis indicated a noteworthy increase in the green fluorescence of JC-1. Therefore, an increase in green fluorescence indicates a shift towards lower membrane potential, often associated with a notable impairment in mitochondrial function. This impairment was further supported by the rise in the percentage of dysfunctional mitochondria in cells treated with MPs, particularly PET 1, PET 2.6 (p<0,001), and PE (p<0,01) (Fig. 3B).

Effect of PET and PE on autophagy

Intrigued by the cytotoxic effects of MPs on cellular dynamics, our study delved into the potential involvement of autophagy as a key player in mediating these effects. We employed the Cyto-ID® Autophagy Detection Kit to assess autophagic responses in cells exposed to MPs. Our goal was to unravel the intricate interplay between MPs and the cellular autophagic machinery. Western blot analyses of autophagic markers LC3 and p62 were conducted under conditions both with and without bafilomycin, an inhibitor of autophagosome- lysosome fusion. Immunofluorescence further enriched our understanding, revealing spatial and temporal nuances in the distribution of LC3 and p62.

Increased autophagic flux in HepG2-treated with MPs was verified using the Cyto-ID® Autophagy Detection assay. This assay utilizes a cationic amphiphilic tracer dye, which readily infiltrates cells and facilitates the marking of vacuoles linked to the autophagy pathway. As depicted in the figure (Fig. 4A, B), there was a notable rise in the percentage of autophagic vacuoles in HepG2 cells treated with MPs PET 1 (p<0,001), PET 2.6 (p<0,05) and PE (p<0,01) in comparison to the control group.

Fig. 4.

Autophagic flux analysis through Cyto-D assay, western blotting and immunofluorescence of control and MPs (PET 1 µm, PET 2,6 µm and PE 2,6 µm) treated HepG2. A Representative intracellular Flow Cytometric Cyto-ID B Histogram showing the percentage of Cyto-D positive cells. C WB analysis shows the expression of LC3-I (16 kDa), LC3-II (14 kDa) and p62 (62 kDa) in HepG2 exposed to MPs. D The histogram shows the LC3-II level. Data were normalized with beta-Actin (44 kDa). E Histogram showing the relative level of p62. Proteins levels were quantified using the ImageJ program. Data were normalized with beta- Actin and reported as OD ratio. All the values are expressed as means ± SEM. Asterisks indicate a significant difference from the respective control (*: p < 0.05), (** p < 0.01) and (***: p < 0.001) after one-way ANOVA using Tukey’s Post Hoc test. F LC3 (green), phalloidin (Red) immunolocalization HepG2 cells treated with MPs. Cell nuclei were stained with DAPI (blue). The images were captured at × 20 magnification. Scale bars represent 20 µm. G the histogram shows the quantification of LC3 fluorescence signal intensity. Data were normalized with the signal of phalloidin using ImageJ. All values are expressed as means ± standard deviation. PET 1, PET 2,6 and PE 2,6 Vs CTRL: (*: p < 0.05, ** p < 0.01). H Representative micrographs of p62 immunostaining (green), and phalloidin (red) on HepG2 cultures treated with MPs. Cell nuclei were stained with DAPI (blue). I The graph shows the quantification of p62 fluorescence signal intensity Using Tukey’s Post Hoc test. Asterisks indicate a significant difference from the respective control (*: p < 0.05,**: p < 0.01) after one-way ANOVA using Tukey’s Post Hoc test

To ascertain if autophagy is triggered by MPs toxicity, LC3 and p62, autophagy markers, were analyzed (Fig. 4C). As depicted in (Fig. 4E), the level of p62 was notably decreased in HepG2 cells exposed to environmental MPs-PET and MPs-PE (p<0,001) compared to the control group, confirming that MPs induced the autophagic degradation process. While the level of LC3II was higher in HepG2 treated with PET 1 µm (p<0,001), PET 2.6 µm (p<0,05) and PE (p<0,05) compared to control (Fig. 4D), suggested an increase in autophagic activity.

We carried out quantitative immunofluorescence autophagic markers (LC3II and p62), and a significant variation was noted across all tested conditions, after HepG2 cells treatment with MPs from the control group. As shown in the (Fig. 4F, G) immunofluorescence of LC3II, showed a notable rise in HepG2 treated with PET (p<0,05) and PE (p<0,01) with the enhancement of fluorescence intensity while p62 decreased significantly (Fig. 4H, I), which suggests an augmentation in autophagic flow. Both LC3II and p62 were influenced by MPs treatment compared to control.

Finally, to better confirm the activation of the autophagic flux, we quantified the expression of LC3-II and p62 in the presence of bafilomycin A1 (Baf A1), a specific inhibitor of type HC-ATPase vacuolar, prevents autophagy at a later stage by blocking the fusion between the autophagosomes and lysosomes was used in this study (Fig. 5). We noted that the treatment with Baf A1 produced an increased protein levels of both LC3-II and p62 (Fig. 5B, C), these results suggest an activation of autophagy, as the accumulation of LC3-II and p62 indicates increased autophagosome formation and cargo recruitment, respectively. However, the blockade in the fusion between autophagosomes and lysosomes at a later stage may indicate impaired autophagic flux. This underscores the complex interaction between microplastics and the cellular autophagic machinery.

Fig. 5.

Co-treatment with bafilomycin A1 A WB analysis showing the expression of LC3-I (16 kDa), LC3-II (14 kDa) and p62 (62 kDa) in HepG2 exposed to MPs in the presence of Bafilomycin A1. Prior to protein extraction, cells were incubated for 2 h with 100 nM Bafilomycin A1 (an inhibitor of lysosomal degradation). B Histogram shows the LC3-II level. Data were normalized with beta-Actin (44 kDa). C Histogram showing the relative level of p62. Protein levels were quantified using the ImageJ program. Data were normalized with beta-Actin and reported as OD ratio. All the values are expressed as means ± SEM. Asterisks indicate a significant difference from the respective control (*: p < 0.05), (** p < 0.01) and (***: p < 0.001) after one-way ANOVA using Tukey’s Post Hoc test

Discussion

The accumulation of plastic waste has evolved into a significant environmental issue. MPs which are debris resulting from the degradation of plastic, have emerged as a novel form of pollution. Indeed, MPs and nanoplastics are gaining significant notice owing to their extensive presence and dispersion in natural environments, along with their adverse effects on ecosystems [32] and potential health hazards for humans [3]. Human exposure to MPs is unavoidable, and the liver, being a crucial organ for detoxification, is particularly susceptible to the bioaccumulation and potential toxicity of MPs [33]. Alternatively, our comprehension of the potential negative health effects of microparticles of plastics remains limited. Moreover, the HepG2 cell line serves as a prevalent tool for assessing the hepatotoxic effects of external substances.

In alternative terms, HepG2 cells were utilized to study variances in hepatocyte toxicity and to assess the potential harmful effects of PET and PE, in terms of viability and oxidative stress. Indeed, after 72 h of exposure to MPs, results demonstrated that HepG2 cells are more sensitive to PE because the Minimum Active Concentration (MAC values) was 40.07 µg/ml which was lower than that of PET. The CCK-8 assay demonstrated that this exposure significantly disturbed cell proliferation. In fact, after three days of treatment, a significant increase in the proliferation rate for both types of MPs was observed. Our findings did not align with prior research, which showed that MP-PET decreased proliferation in mesenchymal stromal cells (MSCs) [29], and that MPs-polystyrene (PS) led to a notable decrease in cell proliferation in Human Lung Cells [34], as well as in human hepatocellular liver cells (HepG2) and human embryonic kidney cells (HEK 293) [35]. This discrepancy may be due to differences in the type of microplastics used. In our study, the MPs were extracted from real-life PET bottles and were characterized by smaller particle sizes (submicron to low micrometric range), irregular shapes, and possibly the presence of surface oxidation or residual additives. These physicochemical properties can significantly influence bioavailability, cellular uptake, and the activation of adaptive cellular responses, which may contribute to the observed divergence in cytotoxicity and proliferation outcomes.To ascertain the factors contributing to this rise, we carried out the intracellular level of ROS using DCF-DA and TBARS assay. In a similar vein, our findings underscored that exposing HepG2 cells to PET and PE resulted in an accumulation of ROS across all treatment conditions. This observation aligns with numerous studies indicating that the exposure of living cells to MPs can trigger an excessive generation of free radicals [36, 37]. Other studies highlighted that the treatment of human cells such as T98G and HeLa with polyethylene and polystryrene caused a high level of ROS [38]. It is also important to note that Poma and colleagues [39] found that 100 nm PS NPs prompt the generation of ROS and cause genotoxic stress and DNA damage. In the face of stressful conditions, autophagy, a natural cellular process of self-degradation, becomes crucial for sustaining normal homeostasis through the breakdown of proteins and the turnover of cell organelles [40]. This cellular process can be categorized into two main types: microautophagy, characterized by direct encasement of cytoplasmic material by the lysosomal membrane for degradation, and macroautophagy, which entails the formation of autophagosomes in the cytosol to engulf cargo, fuse with lysosomes, and release their contents for degradation [41]. The process of selectively degrading mitochondria through autophagy is termed “mitophagy.” Mitophagy serves as a regulatory mechanism that helps maintain the integrity of mitochondrial networks when ROS are generated during stressful conditions, aiming to prevent the buildup of mutations in mtDNA. It is well- established that the excessive generation of mitochondrial ROS (mROS) is acknowledged as one key factor driving mitochondrial damage. In simpler terms, when there is an overproduction of mROS, it leads to harm to the mitochondria [42, 43]. In this context, we carried out the mitochondrial mtDNA integrity analysis and the MMP assay using JC-1. Our results demonstrated a reduction in mitochondrial mtDNA integrity and a significant increase of dysfunctional mitochondria in HepG2 cells treated with PET and PE. Certainly, our findings align with several other studies that have also demonstrated significant mitochondrial damage induced by PS-NPs. This damage is characterized by alterations in mitochondrial morphology, mtDNA, reduction in MMP, and the induction of mitochondrial dysfunction [44, 45]. Additionally, Rottenberg and colleagues [46] have shown that the release of excess mROS within cells disrupts the integrity of the mitochondrial membrane. This disruption impacts MMP, contributing to increased mitochondrial damage.

While MPs may eventually be transported to the lysosome, they are not readily digested. Instead, their accumulation in endosomes or late lysosomes disrupts the degradative functions of these organelles. This disruption significantly impacts the essential process of cell membrane renewal through macroautophagy [47]. We evaluated the autophagic process using Cyto-ID and protein levels of LC3-II and p62 by immunofluorescence and western blot. Our results indicated a surge in cyto-ID, indicating an elevation in autophagic flux. LC3-II and p62 play pivotal roles in regulating the autophagic process [48]. Specifically, elevated LC3 level is known to trigger autophagy activation [49], whereas reduced p62 expression facilitates autophagosome formation and enhances LC3 expression, consequently boosting autophagic flux [50, 51].

Furthermore, our data showcased an upregulation of LC3II and a decline in p62, validated through both western blot and immunofluorescence analyses. Taken together, these results provide evidence of an induction of autophagy. Our results correlate with those of Lim et al. [47], who demonstrated the occurrence of autophagy in lung cells in response to oxidative stress provoked by exposure to PS. It is worth noting that compromised autophagic clearance could initiate positive feedback mechanisms ultimately leading to autophagic cell death. Conversely, microplastics/nanoplastics can also trigger autophagy. Additionally, nanoparticles have been shown to regulate autophagy [52]. It is also elucidated PS-MPs elicited autophagic cell death in bronchial epithelial cells [53], and in human umbilical vein endothelial cells [54].

Therefore, elevated levels of autophagy can indicate either an augmentation in autophagy itself or a hindrance in lysosomal processing downstream of these autophagosomes, or both scenarios simultaneously. Baf A1, a selective inhibitor of vacuolar type H^+-ATPase wich hinders autophagy at a later stage by impeding the fusion between autophagosomes and lysosomes was used. We evaluated the expression levels of LC3-II isoforms and p62 under conditions with and without Baf A1. Our data disclosed an increase in both LC3 and p62 levels in the presence of Baf A1, indicating that the autophagic flux is active. Indeed, the elevation in LC3 levels indicates enhanced autophagosome formation, suggesting that the presence of MPs stimulates the initiation of autophagy. Concurrently, the accumulation of p62 suggests that autophagic flux is impaired. Despite the induction of autophagosome formation, the degradation of cargo (including p62 itself) within autolysosomes is hindered, likely due to the presence of MPs. These data highlighted the complex effects of MPs on the autophagic process and underscored the importance of considering both initiation and flux dynamics when assessing autophagy modulation in the presence of MPs.

Conclusion

In conclusion, our study demonstrated that HepG2 cells exposed to PET and PE increase cell proliferation, which subsequently induces cytotoxic effects by triggering oxidative stress and disrupting mitochondrial function. This interference extends to various intracellular biological pathways, such as compromising mtDNA integrity and triggering autophagy processes. Given these findings, it appears imperative to conduct further research in this area. Specifically, there is a potential for HepG2 cells to exhibit increased tumorigenicity. Consequently, additional investigations are warranted to enhance our understanding of MP contamination, human ingestion patterns, and the associated health ramifications.

Authors’ contributions

HN, NA, and MV provided substantial contributions to conception and design, data acquisition, and data analysis and interpretation; HN, NA, MV, SM, MB, GOC, MF: Writing—original draft, Writing—review & editing, and critically revising it for important intellectual content; GOC, MF, MB: provided substantial contributions to data analysis and exposures methodology; GOC, MF, GB, UG, IM, SM, MB, they provided final approval of the version to be published; NH, NA, MV, IL, DA, GOC, MF, GB, UG, SM and MB are agreement to be accountable for all aspects of the work in ensuring that questions related to the accuracy and integrity of the work are appropriately investigated and resolved.

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

Not applicable.

Competing interests

The authors declare no competing interests.