# Benzo[a]pyrene induces an adaptive mitochondrial-dynamics response in endothelial cells, at human relevant concentration

Joan Guillouzouic, Camille Chauvin, Valentine Genet, Simon Martarello, Agnès Burel, Lydie Sparfel, Normand Podechard, Dominique Lagadic-Gossmann, Catherine Lavau, Eric Le Ferrec

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## To cite this version:

Joan Guillouzouic, Camille Chauvin, Valentine Genet, Simon Martarello, Agnès Burel, et al. Benzo[a]pyrene induces an adaptive mitochondrial-dynamics response in endothelial cells, at human relevant concentration. Environmental Pollution, 2026, 391, pp.127538. 10.1016/j.envpol.2025.127538. hal-05456356

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https://hal.science/hal-05456356v1

Submitted on 15 Jan 2026

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# Benzo[a]pyrene induces an adaptive mitochondrial-dynamics response in endothelial cells, at human relevant concentration☆

Joan Guillouzouic<sup>a</sup>, Camille Chauvin<sup>a</sup>, Valentine Genet<sup>a,●</sup>, Simon Martarello<sup>a</sup>, Agnes Burel<sup>b,†</sup>, Lydie Sparfel<sup>a</sup>, Normand Podechard<sup>a</sup>, Dominique Lagadic-Gossmann<sup>a,●</sup>, Catherine Lavau<sup>a</sup>, Eric Le Ferrec<sup>a,∗</sup>

## ARTICLE INFO

### Keywords:

Polycyclic aromatic hydrocarbons

Aryl hydrocarbon receptor

Endothelial cells

Mitochondria

Mitochondrial dynamics (fusion/fission)

Stress-induced mitochondrial hyperfusion

## ABSTRACT

Although polycyclic aromatic hydrocarbons (PAHs) are a public health concern in many countries, the cellular response at concentrations representative of human exposure remains poorly defined. At the cellular level, because of their function and composition, mitochondria are privileged targets of these contaminants. PAHs such as benzo[a]pyrene (B[a]P) can compromise mitochondrial functions. Using an integrated approach, we examined how low-dose B[a]P disrupts the three determinants of mitochondrial homeostasis (biogenesis, mitophagy, and dynamics) and triggers apoptosis. We assessed the cellular response to a low-dose of B[a]P (100 nM) in the human microvascular endothelial cell line HMEC-1. Endothelial cells are sensitive to PAHs and display marked reactivity to effectors of B[a]P toxicity, such as oxidative stress. We show that B[a]P doesn't modify mRNA expression of mitochondrial biogenesis markers but causes a blockage of autophagic flux. In parallel, we observe early and transient mitochondrial elongation accompanied by decreased expression of the fission factors DRP1 and MFF, and increased intracellular concentration of ATP. These events constitute cellular signatures of stress-induced mitochondrial hyperfusion (SIMH), a mechanism of resistance to apoptosis. B[a]P eventually causes apoptosis of HMEC-1 cells, which is mitochondria-dependent, caspase-independent, requires B[a]P bioactivation by CYP1 family enzymes, and is only detectable after 24 h of exposure when SIMH markers decline. Our results show that endothelial cells can mobilize SIMH to delay and limit B[a]P-induced apoptosis. We confirm that mitochondria, at human-relevant doses, are early sensitive targets of B[a]P and show that changes in mitochondrial dynamics constitute an early readout of cellular responses to B[a]P.

## 1. Introduction

It is now widely accepted that many environmental contaminants exert their toxicity by disrupting mitochondrial functions (K. Li et al.,

2023). This intracellular organelle is particularly vulnerable as the high phospholipid content of its membranes, the presence of calcium transporters, and its pH are factors that promote the accumulation of xenobiotics such as polycyclic aromatic hydrocarbons (PAHs) in

**Abbreviations:** AhR, Aryl Hydrocarbon Receptor; AIF, Apoptosis-Inducing Factor; ATP, Adenosine Triphosphate; A.U, Arbitrary Unit; α-NF, Alpha-Naphthoflavone; B[a]P, Benzo[a]Pyrene; CSA, Cyclosporine A; CYP1A1 / CYP1B1, Cytochrome P450 family 1 (subfamily A1 / B1); DMSO, Dimethyl Sulfoxide; DRP1, Dynamin-1-Like Protein (DNM1L); FCCP, Carbonyl Cyanide-4-(trifluoromethoxy)Phenylhydrazone; HMEC-1 cells, Human Microvascular Endothelial cells; HSC-70, Heat Shock Cognate 70 kDa protein; MFF, Mitochondrial Fission Factor; MFN1 / MFN2, Mitofusin 1 / 2; MTG / MTDR, MitoTracker™ Green / MitoTracker™ Deep Red; mPTP, mitochondrial Permeability Transition Pore; mtDNA / nDNA, mitochondrial DNA / nuclear DNA; NDP52, Nuclear Dot Protein 52 kDa (CALCOCO2); NRF2 (NFE2L2), Nuclear Factor Erythroid 2-Related Factor 2; Keap1, Kelch-like ECH-Associated Protein 1; PAHs, Polycyclic Aromatic Hydrocarbons; PARKIN, E3 Ubiquitin-Protein Ligase Parkin (PARKIN); PINK1, PTEN-Induced Kinase 1; pUb (S65), phospho-Ubiquitin (Serine 65); qPCR / RT-qPCR, quantitative PCR / Reverse-Transcription qPCR; ROS, Reactive Oxygen Species; SEM, Standard Error of the Mean; SIMH, Stress-Induced Mitochondrial Hyperfusion; SLP2, Stomatin-Like Protein 2; TEM, Transmission Electron Microscopy; TFAM, Mitochondrial Transcription Factor A; ZVAD-FMK, Cell-permeant pan-caspase inhibitor.

<footnote><sup>a</sup> Univ Rennes, Inserm, EHESP, Irset (Institut de Recherche en Santé Environnement et Travail) - UMR_S 1085, F-35000, Rennes, France</footnote>

<footnote><sup>b</sup> Univ Rennes, Plate-forme MRic-TEM, Irset, Biosit - UMS 3480, F-35000, Rennes, France</footnote>

<footnote>* This paper has been recommended for acceptance by Wen Chen.</footnote>

<footnote>* Corresponding author. UMR Inserm U1085, IRSET, University of Rennes, Faculty of Pharmacy, 9 av. Pr. Léon Bernard, cedex, 35043, RENNES, France.</footnote>

<footnote>E-mail address: eric.leferrec@univ-rennes.fr (E. Le Ferrec).</footnote>
mitochondria (Pang et al., 2024a, 2024b). PAHs are among the environmental contaminants to which humans are continuously exposed, particularly through dietary intake (Agus et al., 2023; Bansal and Kim, 2015). It is established that human exposure to these environmental pollutants occurs daily and is associated with the onset and progression of ailments such as cancer and cardiovascular diseases (Choi et al., 2010). B[a]P, one of the most extensively studied PAH, is classified as a Group 1 human carcinogen by the IARC (IARC Working Group on the Evaluation of Carcinogenic Risks to Humans, 2010). B[a]P toxicity requires bioactivation by cytochrome P450 enzymes (CYP1 family). These enzymes, whose expression is regulated through activation of the cytosolic aryl hydrocarbon receptor (AhR), are found in the endoplasmic reticulum as well as within the mitochondria (Dong et al., 2009). During the biotransformation of B[a]P, reactive oxygen species (ROS) and toxic metabolites, such as B[a]P-7,8-dihydrodiol-9,10-epoxide (BPDE), are generated. These electrophilic intermediates can bind to nucleophilic sites within the cell, such as DNA. BPDE is therefore considered a genotoxic and mutagenic compound, primarily through the formation of adducts at the N² position of guanine (Piberger et al., 2018). This phenomenon is particularly evident in mitochondria, due to the presence of CYP1A1 and CYP1B1 in this organelle (Backer and Weinstein, 1980; Bhagwat et al., 1999), as well as a mitochondrial DNA (mtDNA) repair system that is less efficient than that of nuclear DNA (nDNA). As a result, mtDNA is considered to be over 40 times more sensitive to adduct formation than nDNA (Backer and Weinstein, 1980).

Mitochondrial homeostasis results from the balance between mitochondrial biogenesis, mitochondrial dynamics (fusion, fission), and mitophagy (Pang et al., 2024a, 2024b). This balance can be modified by the pathophysiological state of the host cell. While mitochondrial fission can isolate daughter mitochondria destined for mitophagy, mitochondrial fusion can, on the other hand, preserve mitochondria by diluting damaged mitochondrial components within the mitochondrial network. The latter is observed during stress-induced mitochondrial hyperfusion (SIMH). SIMH is a transient state used by cells to maintain ATP production and enhance their resistance to various stresses (e.g., oxygen deprivation, UV irradiation, or inhibition of protein synthesis, notably during exposure to xenobiotics) (Tondera et al., 2009). By counteracting autophagy (Gomes et al., 2011; Seager et al., 2020), SIMH is more specifically described as an adaptive response induced by moderate levels of mitochondrial damage and aimed at protecting mitochondria. Therefore, SIMH can be considered as a mechanism by which the cell enhances its functional robustness and resistance against cell death (Tondera et al., 2009). Mitochondrial hyperfusion in SIMH can result from an increase in fusion (notably via OPA1 [optic atrophy 1] and mitofusins) (Tondera et al., 2009) or from a decrease in fission (notably due to the reduction in DRP1) (van der Bliek, 2009). These morphological changes are accompanied by an increase in mitochondrial ATP production.

Under cellular stress conditions, alteration of the different determinants of mitochondrial homeostasis can lead to a maladaptive response. This phenomenon is particularly observed following exposure to environmental contaminants such as PAHs (Reddam et al., 2022; Schnegelberger et al., 2021), as the determinants of mitochondrial homeostasis are all potential targets of the PAHs/AhR pathway. Indeed, AhR has been associated with an increase in mitochondrial biogenesis as part of a survival response in human melanocytes exposed to oxidative damage (Wang et al., 2019). It has also been demonstrated that, in the presence of B[a]P, cells can initiate an adaptive mitochondrial response involving mitophagy processes to resist apoptosis (Das et al., 2017). Recent work from our team has shown that B[a]P induces mitochondrial elongation in hepatic epithelial F258 cells (Hardonnière et al., 2017). This result suggests that B[a]P is capable of modulating mitochondrial dynamics. These modifications of mitochondrial homeostasis determinants induced by B[a]P may be a defense strategy implemented by cells in response to toxicant exposure.

Although some studies have examined the effect of exposure to low

concentrations of B[a]P, these remain the exception (Hardonnière et al., 2017). Many studies employ high B[a]P concentrations, in the micromolar range (from 1 to 20 µM). While highly informative, such concentrations are not representative of human exposure. Although the reported levels vary, as these are strongly associated with lifestyle (e.g., smoker vs. non-smoker), the average serum concentrations of B[a]P in humans are in the nanomolar range (Neal et al., 2008; Ramesh et al., 2015; Song et al., 2013).

This study evaluates the effects of human-relevant concentrations of B[a]P on the three principal determinants of mitochondrial homeostasis (biogenesis, dynamics [fusion/fission]), and mitophagy) in a human endothelial model (HMEC-1 cell line). We selected an endothelial model because these cells are highly responsive to B[a]P (Shi et al., 2021) and display marked reactivity to B[a]P toxicity effectors such as oxidative stress (Anasooya Shaji et al., 2019). Moreover, their position at the interface between the bloodstream, the vector of pollutants, and the liver, the primary site of detoxification, renders them particularly susceptible to dietary toxicants such as PAHs.

We show that, in HMEC-1 endothelial cells, exposure to low concentrations of B[a]P does not induce significant changes in mitochondrial biogenesis and is associated with impairment of autophagic flux (accumulation of autolysosomes). In contrast, such exposure elicits a transient response characterized by pronounced mitochondrial elongation (decreased circularity index and increased perimeter), increased ATP production, and reduced expression of DRP1 and MFF, two key regulators of mitochondrial fission. The combination of mitochondrial elongation, elevated ATP output, and downregulation of fission factors is consistent with SIMH and in line with NRF2 activation as an initiating event. Our data suggest that this adaptive SIMH may confer a degree of resistance to B[a]P-induced apoptosis.

## 2. Materials and methods

### 2.1. Chemicals and reagents

B[a]P, carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP), cyclosporine (CSA), dimethyl sulfoxide (DMSO), Hoechst 33342, epithelial growth factor (EGF), α-naphthoflavone (α-NF), and cell-permeant pan-caspase inhibitor (ZVAD-FKM) were purchased from Sigma-Aldrich (USA). MCDB131 was purchased from US Biological Life Sciences (USA). Sodium bicarbonate, L-glutamine, penicillin/streptomycin, gentamycin, and phosphate-buffered saline (PBS) were purchased from Gibco (USA). Fetal bovine serum (FBS) was purchased from Dutscher (France). Hydrocortisone was purchased from Upjohn (USA). Bovine serum albumin fraction V (BSA), PhosSTOP™ cOmplete Mini and Protease Inhibitor Cocktail™, brusatol, and sulforaphane were purchased from Merck (Germany). MitoTracker™ Green FM (MTG), MitoTracker™ DeepRed FM (MTDR), LysoTracker™ DND-99 Red, High-Capacity cDNA Reverse Transcription Kit™, Master Mix PCR Power SYBR™ Green, SuperSignal ELISA Femto Maximum Sensitivity Substrate™, and Pierce™ BCA Protein Assay Kit were purchased from Thermo Fisher Scientific (USA). Bio-Rad Protein Assay Dye Reagent™ was purchased from Bio-Rad (USA). TOM20 (D8T4N), OPA1 (D6U6N), DRP1 (D6C7), MFF (E5W4M), Mitofusin-1 (D6E2S) and Mitofusin-2 (D1E9) antibodies are part of the Mitochondrial Dynamics Antibody Sampler Kit II; NDP52 (D1E4A), Optineurin (D2L8S), PARKIN (Prk8), PINK1 (D8G3), BNIP3L/NIX (D4R4B), LC3B (D11) and Phospho-Ubiquitin Ser65 (E2J6T) antibodies are from the Mitophagy Antibody Sampler Kit; COX IV (3E11) and AIF (D39D2) antibodies were also purchased from Cell Signaling (USA). HSC70 (SC-8432), β-Actin (SC-7298), and NRF2 (A-10) antibodies were purchased from Santa Cruz Biotechnology (Germany).

### 2.2. Cell culture

The HMEC-1 cell line used in this study was obtained from the
Centers for Disease Control and Prevention (Atlanta, USA). Cells were cultured in Endothelial Basal Medium MCDB131, adjusted to pH 7.6 with sodium bicarbonate, and supplemented with 10 % heat-inactivated FBS (56 °C, 30 min), L-glutamine (10 mM), penicillin/streptomycin (100 U/mL), gentamicin (500 µg/mL), EGF (10 ng/mL), and hydrocortisone (1 µg/mL). Cells were maintained at 37 °C in a humidified incubator with 5 % CO₂ in 150 cm² culture flasks. At 90 % confluence, cells were incubated overnight in FBS-free medium. Subsequently, cells were exposed to B[a]P, prepared as a stock solution in DMSO. The final vehicle concentration (DMSO) did not exceed 0.05 % (v/v), and control cultures received the same amount of DMSO. In experiments involving chemical inhibitors or activators, these were applied 1 h before and maintained during the exposure to B[a]P.

## 2.3. Apoptosis

Cells were cultured in 6-well plates for apoptosis assays. Following treatment, both the culture medium and cells trypsinized with 0.05 % trypsin were collected and centrifuged (1500 rpm, 5 min, 4 °C). The cell pellet was resuspended and incubated in 200 µL of Hoechst 33342 solution (50 µg/mL) at 37 °C for 20 min to stain DNA. Cells were then examined for apoptotic features (chromatin condensation and nuclear fragmentation) using a fluorescence microscope (Axio Scope A1, Zeiss, Germany). A minimum of 300 cells per condition was counted in randomly selected fields. Results are expressed as the percentage of apoptotic cells, normalized to the control condition (DMSO).

## 2.4. Mitochondria isolation

Mitochondria from HMEC-1 cells were isolated following the procedure described by Zhou et al., (2023). All steps were performed at 4 °C. Cells were harvested by scraping and centrifugation in PBS (5 min, 1500 rpm). The cell pellet was resuspended in isolation buffer (20 mM HEPES, pH 7.5, 1.5 mM MgCl₂·6H₂O, 10 mM KCl, 1 mM EDTA, 1 mM EGTA, 0.25 M sucrose) supplemented with a protease inhibitor cocktail (Roche cOMPlete™ Mini, 04693124001). After Dounce homogenization and a 20 min incubation on ice, the supernatant was collected by centrifugation (1620×g, 10 min) and clarified by a second centrifugation under the same conditions (1620×g, 10 min). The resulting supernatant was centrifuged at 15,700×g for 20 min to pellet mitochondria. The mitochondrial pellet was resuspended in 50 µL of isolation buffer and stored at -80 °C.

## 2.5. Protein quantification and western blot

Cells were lysed in RIPA buffer supplemented with phosphatase and protease inhibitors, and lysates were stored at -20 °C until protein quantification. Protein concentration was determined using either the Bradford assay with Protein Assay Dye Reagent Concentrate (Bio-Rad, USA; 1:5 dilution in ultrapure water) or the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, USA). BSA served as the standard. Absorbance was measured using a CLARIOstar® Plus microplate reader (BMG LABTECH, Germany) at 595 nm for the Bradford assay and at 562 nm for the BCA assay.

Thirty µg of protein were mixed with Laemmli buffer, denatured at 95 °C for 10 min, and loaded onto 8–12 % SDS-PAGE polyacrylamide gels (acrylamide/bis-acrylamide). Electrophoresis was performed in running buffer (50 mM Tris-HCl, 400 mM glycine, 0.1 % SDS, pH 8.3). Proteins were then transferred to a nitrocellulose membrane (Millipore, France) overnight at 30 V and 4 °C using transfer buffer (25 mM Tris-HCl, 192 mM glycine, 20 % ethanol). After transfer, the membrane was blocked with 4 % TBS/BSA. Primary and secondary antibody incubations (respectively 2 h and 1 h at RT) were performed in TBS/BSA. Between each step, membranes were washed several times with 1 × Tris-buffered saline containing 0.1 % Tween-20 (TBS-T). Immunolabeled proteins were visualized using the SuperSignal™ ELISA Femto

Maximum Sensitivity Substrate (Thermo Fisher Scientific, USA). Chemiluminescence signals were detected using a LAS3000 imaging system (Bio-Rad, USA), and image analysis was performed with Image Lab 6.0 software (Bio-Rad, USA). HSC70 and β-Actin were used as loading controls.

## 2.6. Flow cytometry

Following treatment, cells were rinsed with pre-warmed MCDB131 medium without FBS. They were then incubated at 37 °C for 20 min in MCDB131 medium supplemented with either MTDR and MTG (final concentration: 100 nM each), or LysoTracker™ DND-99 (final concentration: 100 nM). Hoechst33342 (final concentration: 5 µg/mL) was added to all staining conditions. Following incubation, cells were rinsed with pre-warmed PBS, trypsinized, and centrifuged (1500 rpm, 2 min, RT). The resulting cell pellets were resuspended in 1 mL of PBS, kept on ice, protected from light, and analyzed by flow cytometry within 1 h. Data acquisition was performed using a BD LSR Fortessa flow cytometer (BD FACSCanto™ II software, BD Biosciences), with the allophycocyanin-A (APC-A) filter for MTDR, the fluorescein isothiocyanate-A (FITC-A) filter for MTG, and the phycoerythrin-A (PE-A) filter for LysoTracker™. Between 15,000 and 20,000 cells were recorded per condition. Unstained cells were analyzed in parallel under identical conditions to determine baseline autofluorescence. The gating strategy was designed to exclude more than 99 % of autofluorescent signals. Below this threshold, fluorescence was not considered indicative of mitochondrial presence, mitochondrial polarization, or lysosomal content.

## 2.7. ATP measurement

ATP production levels were measured using the CellTiter-Glo® Luminescent Cell Viability Assay (Promega, USA). Briefly, cells were seeded at a density of 30,000 cells per well in a 96-well plate. After treatment, cells were washed with pre-warmed PBS. After removal of PBS, 100 µL of CellTiter-Glo reagent was added directly to each well. Plates were incubated for 30 min at RT to allow cell lysis and luminescence stabilization. Luminescence was measured at 560 nm using the EnSpire™ Multimode Plate Reader (PerkinElmer, USA). Results were normalized to total protein content per well (in µg), as determined by the Bradford assay.

## 2.8. Transmission electron microscopy

Cells were rinsed with sodium cacodylate buffer (0.15 M, pH 7.2) and fixed with 2.5 % glutaraldehyde for 1 h at RT, followed by post-fixation in 1.5 % osmium tetroxide for 1 h. Samples were then dehydrated in a graded ethanol series (50 %, 70 %, 90 %, and 100 % v/v) and progressively infiltrated with increasing concentrations of ethanol/ epoxy resin mixtures (30 %, 50 %, and 70 %; 3 h per step). Cells were incubated overnight in pure epoxy resin before final embedding in 2,4,6-tris(dimethylaminomethyl)phenol-30-epoxy resin (3 h at RT, followed by polymerization at 60 °C for 24 h). Ultrathin sections (80 nm) were cut using an ultramicrotome (UC6, Leica, Germany), collected on copper grids (Agar Scientific, UK), and post-stained with 2 % uranyl acetate (30 min) and 3 % lead citrate (20 min). Grids were examined at 120 kV using a JEOL 1400 transmission electron microscope (Jeol Ltd., Japan) equipped with a Gatan Orius SC1000 camera (Gatan, USA). Mitochondrial morphology was assessed using ImageJ software by measuring perimeter, surface area, and circularity index ($4\pi$-(surface area/perimeter²)), as described previously (Lam et al., 2021; Picard et al., 2013). We performed three rounds of imaging per condition, analyzed more than 50 images per condition, and measured the circularity and perimeter of over 1500 mitochondria per condition.
## 2.9. Analysis of mRNA expression

The study of mRNA expression levels for genes of interest was carried out as previously described (Le Goff et al., 2019). Briefly, total RNA was extracted using TRIzol® reagent (Invitrogen, France), and reverse transcription was performed using the High-Capacity cDNA Reverse Transcription Kit™ (Applied Biosystems, USA). Quantitative PCR (qPCR) was conducted using gene-specific primers (Eurogentec, Belgium). The primers used are listed in the supplementary data table (S1). Amplification data were analyzed using the ABI Prism SDS software with the comparative cycle threshold (ΔΔCt) method. Relative mRNA expression levels were calculated by normalizing target gene Ct values to those of endogenous 18S rRNA.

## 2.10. Statistical analysis

Data were analyzed using GraphPad Prism v8.0.1 (USA) and are presented as mean ± standard error of the mean (SEM), based on at least three independent experiments. Statistical analyses were performed using either one-way analysis of variance (ANOVA) with Dunnett's correction or Student's t-test.

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Effect of B[a]P on mRNA levels of CYP1A1, CYP1B1, and mitochondrial biogenesis marker genes. a) CYP1A1 and CYP1B1 mRNA relative fold induction rates were measured in HMEC-1 cells treated with B[a]P (10, 50, 100, 1000, 10000 nM) or vehicle control (DMSO). b) mRNA relative fold induction rates of mitochondrial genes (ND1, ND2, ND6, COX1, COX3, CYB, and ATP6) and nuclear genes (TFAM, NRF1 and GABP1) were measured in HMEC-1 cells treated with 100 nM of B[a]P or DMSO for 3 h or 24 h mRNA expression of the genes of interest was assessed by RT-qPCR. Results represent the mean (±SEM), normalized to the corresponding control (DMSO), which was arbitrarily set at a value of 1 (horizontal dashed line in b), from at least three independent experiments. \*, *p* < 0.05; \*,\*, *p* < 0.01; \*,\*,\*, *p* < 0.001, compared with the corresponding DMSO-treated control.</figcaption></figure>
## 3. Results

### 3.1. Effect of B[a]P on the expression of molecular markers of mitochondrial biogenesis

Our first objective was to select a B[a]P concentration that elicits a cellular response while remaining within human-relevant exposure levels. B[a]P-induced toxicity depends on its bioactivation by enzymes such as CYP1A1 and CYP1B1, whose expression is regulated by AhR (Uppstad et al., 2010). To determine a relevant concentration of B[a]P exposure, we first conducted a dose-response study (at 24 h exposure) of CYP1 induction. As expected, we observed that the two transcriptional targets of the AhR receptor, i.e., CYP1A1 and CYP1B1, are induced by B [a]P. mRNA induction is detectable at concentrations above 10 nM, with a significant induction obtained above 50 nM (Fig. 1a). The half-maximal effective concentrations (EC50) are 170 nM for CYP1A1 and 64 nM for CYP1B1. Taking this into account, along with our previous work, we chose to use 100 nM as our working concentration for B [a]P.

To evaluate the influence of B[a]P on mitochondrial biogenesis, we used a qPCR-based approach to quantify its effect on the expression of mitochondrial and nuclear encoded markers of mitochondrial biogenesis (Fig. 1b). We tested two exposure times (24 h and 3 h). The early 3 h time point was included because our previous work showed that HMEC-

1 cells are able to respond rapidly to B[a]P (Mayati et al., 2017). The mitochondrial genes analyzed were: ND1, ND2, and ND6 (genes encoding proteins of Complex I of the respiratory chain), CYB (encoding a subunit of Complex III), ATP6 (encoding the F0 subunit 6 of ATP synthase, Complex V), and COX1 and COX3 (encoding subunits of cytochrome c oxidase, Complex IV). The nuclear genes studied included TFAM (encoding mitochondrial transcription factor A), GABP1 (GA binding protein transcription factor subunit α), and NRF1 (encoding nuclear respiratory factor 1). The results (Fig. 1b) show no significant change in the mRNA levels of these genes, regardless of the duration of exposure to B[a]P, except for ND2 and CYB. Indeed, exposure to B[a]P led to a significant decrease in their expression, at 3 h for ND2 and at 24 h for CYB. These results indicate that, under our experimental conditions, exposure to B[a]P does not cause upregulation of mitochondrial biogenesis-associated transcripts.

### 3.2. Effect of B[a]P on expression of protein markers of mitophagy

Mitophagy is a specialized form of autophagy that plays a critical role in maintaining cellular health and survival by selectively targeting and eliminating defective mitochondria. To investigate the influence of B[a]P on mitophagy, we first examined intracellular autophagic vesicles in cells exposed or not to B[a]P (for 3 h or 24 h) using transmission electron microscopy (TEM). TEM images (Fig. 2a) revealed an

<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Effect of B[a]P on autophagic flux. a,b) Representative transmission electron microscopy images of HMEC-1 cells. Cells were analyzed a) after exposure, or not (DMSO), for 3 or 24 h to 100 nM of B[a]P, or b) after 24 h exposure, or not (DMSO), to 100 µM of leupeptin (used as a positive control for autophagic flux inhibition). White arrows point to autolysosomes; N indicates the nucleus, M indicates mitochondria. The image on the right (zoom) shows a magnified view of the region outlined by the black rectangle in the corresponding left panel (scale bar = 1 µm). c) Effect of B[a]P on lysosomal mass. Cells were exposed, or not (DMSO), to 100 nM of B[a]P for 3 or 24 h, and lysosomal content was quantified using LysoTracker™ staining, assessed by flow cytometry. Data represent mean fluorescence intensity (±SEM) in arbitrary units (A.U.), normalized to the control (DMSO), which was arbitrarily set to 1 (indicated by the horizontal dashed line). Results are based on three independent experiments. *, p < 0.05 compared to the DMSO control.</figcaption></figure>
accumulation of electron-dense vesicular structures, particularly visible after 24 h of B[a]P exposure. Their heterogeneous appearance and content are similar to those of electron-dense autolysosomes that accumulate in HMEC-1 cells exposed to leupeptin (a lysosomal protease inhibitor, 100 µM, 24 h), used here as a positive control (Fig. 2b) (Haspel et al., 2011; Klionsky et al., 2021). B[a]P thus appears to impair the establishment of a complete autophagic process, resulting in the accumulation of autolysosomes (Button et al., 2017). Autophagy is a degradative process dependent on lysosomal activity. Some studies indicate that autophagy may be associated with an increase in lysosomal mass (Indira et al., 2018). To assess the mass of the acidic compartment in HMEC-1 cells exposed or not to B[a]P, we performed flow cytometry to assay LysoTracker™ staining. LysoTracker™ fluorescence intensity typically increases during autophagy (Chikte et al., 2014; Warnes, 2015). Cells were exposed to 100 nM of B[a]P for 3 and 24 h to evaluate the influence of exposure time. After 3 h, no variation in the signal was observed. In contrast, after 24 h, a significant decrease in the signal was

detected (Fig. 2c). These observations are consistent with a report that B[a]P impairs lysosomal function, particularly by increasing lysosomal membrane permeability (Anandakumar et al., 2009), and with our previous work demonstrating that a low-dose of B[a]P affects lysosomal function notably by alkalinization (Gorria et al., 2008).

These results confirm that, in the presence of B[a]P, autophagic flux is altered, an effect particularly evident after 24 h of exposure. To further substantiate this, we assessed by Western blot the impact of B[a]P (100 nM, 24 h) on canonical autophagy markers (p62, LC3B) and on mitophagy-related proteins (NDP52, PINK1, PARKIN, Optineurin, BNIP3L/NIX, Ser65-phosphorylated ubiquitin (pUb (S65)) a marker of mitophagy activation (Iorio et al., 2021; Wauer et al., 2015). The results presented in Fig. 3 indicate that B[a]P does not significantly affect the expression of the analyzed mitophagy markers. Furthermore, B[a]P does not alter pUb (S65) levels, whereas FCCP (a mitochondrial uncoupler that dissipates ΔΨm, stabilizes PINK1, and drives ubiquitin Ser65 phosphorylation), used as a positive control, induces its marked increase

<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Effect of B[a]P on the expression of autophagy and mitophagy markers. Following exposure of HMEC-1 cells to DMSO (Control) or to 100 nM of B[a]P for 24 h, cell lysates were analyzed by Western blotting to assess the levels of autophagy markers (p62, LC3B, NDP52) and mitophagy markers (PINK1, PARKIN, Optineurin, BNIP3L/NIX, and pUb (S65)). Densitometric analysis was performed for each blot. Values (arbitrary units, A.U.) represent the mean densitometric ratios (protein of interest/HSC-70, used as a loading control) from at least three independent experiments (±SEM), normalized to the control condition (DMSO), which was arbitrarily set to 1. Abbreviations: BNIP3L/NIX, BCL2/adenovirus E1B 19-kDa-interacting protein 3-like; HSC-70, heat shock cognate 70 kDa protein; LC3B, light chain 3B; NDP52, nuclear dot protein 52 kDa; PINK1, PTEN-induced kinase 1; p62, ubiquitin-binding protein 62; PARKIN, Parkinson protein 2 (E3 ubiquitin ligase); pUb (S65), phospho-ubiquitin (Ser65). FCCP (carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone) was used as a positive control for pUb (S65) induction.</figcaption></figure>
(Gao et al., 2020). These results, consistent with the literature (Paing et al., 2024), confirm that, under our experimental conditions, cells fail to establish a complete autophagic process, and therefore mitophagy, in response to B[a]P-induced stress. It is nevertheless noteworthy that among the proteins analyzed, the expression of NDP52, described as a redox sensor and transcriptionally regulated by the NRF2/Keap1 pathway (Kataura et al., 2023), is increased, albeit without reaching statistical significance, in the presence of B[a]P. This is consistent with the known ability of B[a]P bioactivation to induce oxidative stress and activate the NRF2/Keap1 pathway.

### 3.3. Effect of B[a]P on mitochondrial dynamics

In response to metabolic or environmental stressors, particularly at moderate levels, cells frequently mobilize factors involved in mitochondrial dynamics (fusion/fission balance). The activation of regulatory fission or fusion factors is accompanied by morphological changes in mitochondria, resulting in fragmentation or elongation, respectively. We therefore decided to study the influence of B[a]P on mitochondrial morphology and on the expression of mitochondrial fusion and fission regulators.

#### 3.3.1. B[a]P induces a morphological change of mitochondria in HMEC-1 cells

We used TEM to investigate the effect of exposure to B[a]P (100 nM) for 3 h (Fig. 4a) or 24 h (Fig. 4b) on mitochondrial morphology. Compared with the control condition, cells exposed to B[a]P exhibited elongated mitochondria. This morphological change was particularly pronounced in cells exposed to B[a]P for 3 h. Following the protocol described by Lam et al., (2021), we determined the mitochondrial perimeter and circularity index. Consistent with the images, we observed a significant decrease in the mitochondrial circularity index

(indicating a loss of circularity) and a significant increase in the mitochondrial perimeter (indicative of increased size) after 3 h of exposure to B[a]P (Fig. 4a, right panel). These changes were no longer significant after 24 h of exposure to B[a]P (Fig. 4b, right panel).

#### 3.3.2. B[a]P decreases levels of DRP1 and MFF in HMEC-1 cells

We studied the influence of B[a]P on mitochondrial dynamics proteins that regulate mitochondrial morphology and integrity, including fusion proteins (OPA1, MFN1, MFN2) and fission proteins (SLP2, DRP1, MFF). Protein level analysis by Western blot, regardless of the treatment duration (3 h, Fig. 5a, or 24 h, Fig. 5b), failed to reveal any influence of B[a]P on the expression of fusion proteins. However, the proteins involved in fission appeared to be more sensitive to B[a]P exposure. Among the fission proteins studied, B[a]P exposure led to a significant decrease in the expression of MFF and DRP1. This decrease was observed at 3 h of exposure (Fig. 5a) and persisted at 24 h (Fig. 5b).

#### 3.3.3. B[a]P enhances intracellular ATP concentration in HMEC-1 cells

Exposure to low levels of toxic agents prompts cells to fuse their mitochondria into a network that functions to protect cells against further insults (Schnegelberger et al., 2021). This phenomenon can result from the activation of mitochondrial fusion factors or, as in our case, the inhibition of fission factors. Elongated mitochondria are also associated with an increase in ATP production (Tondera et al., 2009; Strauss et al., 2008). We therefore explored, using a biochemical approach, the intracellular ATP concentration in HMEC-1 cells in the presence of increasing doses of B[a]P (0, 10, 50, or 100 nM). After 3 h of exposure, doses of 50 or 100 nM of B[a]P induced a significant increase in the intracellular ATP concentration (Fig. 6), which is in line with SIMH.

As SIMH (mitochondrial elongation, decreased expression of the fission factors, and increased intracellular concentration of ATP) is

<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Effect of B[a]P on mitochondrial morphology in HMEC-1 cells. a,b) Representative transmission electron microscopy images of HMEC-1 cells. Cells were analyzed after exposure, or not (DMSO), to 100 nM of B[a]P for a) 3 h or b) 24 h. Images (a, b) on the left panel (zoom) show a magnified view of the region outlined by the black rectangle in the corresponding left panel (scale bar = 2 µm). Violin plots, on the left panel, show analysis of mitochondrial perimeter and circularity index after a) 3 h or b) 24 h of exposure to B[a]P. Inserts represent mean values (±SEM) from three independent experiments. ***, *p* < 0.001.</figcaption></figure>
<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

(caption on next column)

**Fig. 5.** Effect of B[a]P on the expression of mitochondrial dynamics markers. Following exposure, of HMEC-1 cells to DMSO (control) or 100 nM B [a]P for a) 3 h or b) 24 h, cell lysates were analyzed by Western blotting to assess the expression of mitochondrial dynamics markers, including fusion factors [OPA1 (optic atrophy 1), MFN1, MFN2 (mitofusines 1 and 2] and fission factors [DRP1 (dynamin-related protein 1), MFF (mitochondrial fission factor), SLP2 (stomatin-like protein 2)]. Densitometric analysis was performed for each blot. Values (arbitrary units, A.U.) represent the mean densitometric ratios (protein of interest/HSC-70, loading control) from at least three independent experiments (±SEM), normalized to the control (DMSO), which was arbitrarily set to 1. *, p < 0.05 compared to the DMSO control.

<figure><img src="image_12.png" /></figure>

considered a mitochondrial protective mechanism employed by cells under moderate-stress conditions (Brown et al., 2025; Khacho et al., 2014), we therefore decided to test the hypothesis that the SIMH observed in HMEC-1 cells exposed to low doses of B[a]P represents a resistance mechanism implemented by the cells, aiming at least to delay apoptosis, if not to resist it altogether.

### 3.4. B[a]P induces mitochondria-dependent but caspase-independent apoptosis

To characterize apoptosis in HMEC-1 cells exposed to B[a]P, we used fluorescence microscopy and Hoechst staining to assess the proportion of cells exhibiting condensed and fragmented nuclei as a function of dose and exposure time. As shown in Fig. 7a, analysis at 24 h of exposure reveals a dose-related increase of apoptosis with a significant effect seen at 100 nM. Compared to a 24 h exposure, exposure to 100 nM B[a]P for 3 or 6 h did not induce apoptosis (Fig. 7b); interestingly, these earlier time points coincide with the period during which SIMH markers are particularly prominent.

As the toxicity of B[a]P depends on its bioactivation through the AhR/CYP1 pathway, we next used α-NF, which acts both as an AhR antagonist and a CYP1 activity inhibitor (Merchant et al., 1993), to test its involvement. As expected (Fig. 7c), α-NF prevented the increase in apoptosis induced by B[a]P, demonstrating that apoptosis is indeed dependent on the bioactivation of B[a]P. We then used a cell-permeant pan-caspase inhibitor (ZVAD-FKM) and an inhibitor (CSA) of the mitochondrial permeability transition pore (mPTP) opening, a critical event in certain types of apoptotic cell death. Fig. 7c shows that only CSA inhibits the apoptosis induced by B[a]P. These results indicate that, in our assay, apoptosis caused by B[a]P is caspase-independent but mitochondria-dependent. To complement this work, we explored the role of apoptosis-inducing factor (AIF). This pro-apoptotic factor can translocate from the mitochondria through mPTP opening, triggering caspase-independent programmed cell death (Candé et al., 2002; Galuzzi et al., 2008). Using Western blot, we evaluated the presence of AIF in the mitochondria after exposure of HMEC-1 cells to B[a]P (100 nM, 24 h). We observed a significant decrease in mitochondrial AIF, consistent with our previous observations (Fig. 7d). Because mPTP opening can cause mitochondrial depolarization (Yalamanchili et al., 2022), we evaluated mitochondrial polarity by flow cytometry using two different mitochondrial staining dyes (MTG and MTDR) (Fig. 7e). B [a]P (100 nM) did not cause any significant change in mitochondrial polarity in HMEC-1 cells exposed for 3 h. In contrast, a significant decrease in mitochondrial polarity was detected at 24 h. Similar results at 24 h were obtained using another mitochondrial staining dye, 3, 3'-Di-n-hexyloxacarbocyanine iodide (data not shown).

These results indicate that mitochondrial perturbation plays an important role in apoptotic cell death in HMEC-1 cells exposed to a low dose of B[a]P. In the presence of B[a]P, mitochondrial depolarization and apoptosis are significant at 24 h, coinciding with a reduction in SIMH markers, notably mitochondrial hyperfusion. This observation supports the hypothesis that HMEC-1 cells engage SIMH as a means of protection from B[a]P toxicity.
<figure><img src="image_13.png" /><figcaption>Fig. 6. Effect of B[a]P on the intracellular level of ATP. ATP content in HMEC-1 cells was measured by spectrofluorimetry after exposure to 10, 50, or 100 nM B[a]P for 3 h. Data represent mean luminescence intensity (±SEM) in arbitrary units, normalized to the control (DMSO), which was arbitrarily set to 1 (indicated by the horizontal dashed line). Results represent the mean (±SEM) of at least three independent experiments. *, *p* < 0.05; **, *p* < 0.01, compared to the DMSO control.</figcaption></figure>

## 3.5. NRF2 involvement in the mitochondrial response to B[a]P exposure

### 3.5.1. B[a]P increases NRF2 levels in HMEC-1 cells

Because SIMH-associated mitochondrial hyperfusion has been linked to Keap1-NRF2 activation via DRP1 downregulation (Das and Chakrabarti, 2020; Sabouny et al., 2017) and B[a]P activates this pathway through oxidative stress (Nguyen et al., 2010; He et al., 2020), we hypothesized an NRF2-dependent link between B[a]P exposure, SIMH, and apoptosis resistance in HMEC-1 cells. We further predicted that if the three elements (NRF2 activation, SIMH, protection against apoptosis) are indeed interconnected, then an activator of NRF2 (sulforaphane) should protect cells from the apoptosis observed at 24 h of exposure to B[a]P, while an NRF2 inhibitor (brusatol) should accelerate the apoptotic process and trigger its onset at earlier time points.

NRF2 activation was evaluated by using immunoblotting to assay its accumulation in B[a]P-treated HMEC-1 cells. In accordance with the literature (Washimkar et al., 2023), we found an increase in NRF2 following exposure to B[a]P. Interestingly, such an increase is only observed at 3 h of exposure to B[a]P, but not at 24 h (Fig. 8a). This suggests that NRF2 activation, the absence of apoptosis, and the activation of SIMH are concomitant events at 3 h.

We therefore measured the rate of apoptosis at 3 h of exposure to B[a]P, in the presence or absence of sulforaphane (5 µM), an NRF2 activator, as well as at 24 h of exposure, with or without brusatol (80 nM), an NRF2 inhibitor. As expected, the presence of sulforaphane led to a complete disappearance of apoptosis at 24 h (Fig. 8b). In contrast, brusatol not only induced apoptosis on its own as early as 3 h of exposure but, as expected, exacerbated apoptosis-related cell death in the presence of B[a]P (Fig. 8c). The same result was obtained using trigonelline, an inhibitor of NRF2 nuclear translocation (data not shown).

## 4. Discussion

In this work, we examined the cellular response to B[a]P concentrations relevant to human exposure (Bukowska et al., 2022), with respect to mitochondrial homeostasis and its three pillars: biogenesis, (auto)mitophagy, and mitochondrial dynamics. Overall, our results demonstrate that in the presence of low doses of B[a]P, endothelial HMEC-1 cells appear to rapidly implement SIMH as a strategy to avoid apoptosis. It is well established that bioactivation of B[a]P, associated with the activation of the AhR pathway and the induction of CYP family 1 enzymes, leads to toxicity resulting in apoptosis mainly triggered by oxidative stress and/or the production of electrophilic metabolites (Bukowska and Duchnowicz, 2022). For this study, we chose a concentration of 100 nM B[a]P, which we found to be sufficient to trigger activation of HAP effectors and which is comparable to levels found in human serum (Walker et al., 2016).

In this study, we demonstrate that apoptosis, dependent on bioactivation, is associated with mitochondrial distress. The involvement of bioactivation is evidenced by the ability of α-NF, an AhR antagonist and CYP1 inhibitor, to suppress B[a]P-induced apoptosis. The mitochondrial origin of apoptosis is confirmed by 1) the decrease in mitochondrial AIF levels in the presence of B[a]P, known to be associated with opening of the mPTP channel and to trigger caspase-independent apoptosis (Candé et al., 2002), and 2) the ability of CSA, an mPTP channel inhibitor, to block this apoptosis. AIF and opening of the mPTP channel are associated with the cellular response to chemical stress involving the production of ROS (Kent et al., 2021; Urbano et al., 2005). These phenomena have been described at low to moderate ROS levels (Wiraswati et al., 2016; Endlicher et al., 2023). In our experimental model, oxidative stress likely contributes to the cellular response (Verma et al., 2017). Nevertheless, we were unable to detect ROS production in our model (using the ROS probe CM-H2DCFDA (2,5 µM), H₂O₂ (100 µM) as positive control and spectrofluorometry, in cells exposed to B[a]P for 3 h) (Oparka et al., 2016; Souki et al., 2023), possibly because the levels of ROS generated by low concentrations of B[a]P may be below the detection limit of our assays. Consistent with this hypothesis, Verma et al., (2017) reported, in RT4 bladder cancer cells, a modest 3 % increase in ROS after a 24 h exposure to 500 nM B[a]P, a concentration fivefold higher than the one we used. The involvement of oxidative stress nevertheless remains highly likely, given that B[a]P toxicity is linked to the production of ROS and that AIF and NRF2, two major actors of mitochondrial physiology and implicated in this work, are widely recognized as responsive to oxidative stress.

Consistent with the literature, which describes mitochondria as organelles particularly sensitive to the toxicity of PAHs (Meyer et al., 2013), mitochondrial dysfunction appears to be a key element of the cellular response to B[a]P in our model (Wang et al., 2018). To maintain mitochondrial homeostasis, cells rely on three mechanisms: mitochondrial biogenesis, mitophagy, and mitochondrial dynamics, which allow morphological and functional adaptation of the organelle. In our experimental model, mitochondrial biogenesis does not appear to be activated, and mitophagy appears to be impaired at late stages. Indeed, we detected no induction in the mRNA levels of genes associated with mitochondrial biogenesis. In parallel, the accumulation of autolysosomes with reduced LysoTracker™ staining (indicative of non-productive autophagy) together with the lack of significant changes in mitophagy markers (NDP52, PINK1, PARKIN, Optineurin, BNIP3L/-NIX) suggests that mitophagy does not proceed to completion under these conditions. This phenomenon has already been reported in other cell models, such as hepatocytes, also in response to low concentrations of B[a]P (Y. Li et al., 2023), and has been confirmed *in vivo* in mice, where B[a]P exposure leads to decreased protein levels of PINK1 and PARKIN (Li et al., 2022). In our model, accumulation of autolysosome-like structures only appears after 24 h of exposure, suggesting a potential impairment of the late stages of autophagy, which appears to be consistent with the onset of apoptosis. This observation is
<figure><img src="image_14.png" /><figcaption>**Fig. 7.** B[a]P-induced, limited, and caspase-independent apoptosis and mitochondrial dysfunction at 24 h. Apoptosis was assessed in HMEC-1 cells exposed, or not (DMSO), to: a) increasing concentrations of B[a]P (0, 50, 100, 500, and 1000 nM, 24 h), b) 100 nM B[a]P for 3, 6, or 24 h, and c) 100 nM of B[a]P (24 h) in the presence or absence of CSA (5 µM), Z-VAD (10 µM), or α-NF (10 µM). Apoptosis induction was estimated by microscopic observation of condensed and/or fragmented chromatin after Hoechst staining. Data are expressed as means (±SEM) from five independent assays, normalized to the control (DMSO), which was arbitrarily set to 100% (horizontal dashed line in b, c). d) Western blot analysis of mitochondrial AIF after exposure to 100 nM of B[a]P for 24 h. COX IV was used as a mitochondrial loading control. e) Mitochondrial membrane potential assessed by flow cytometry, expressed as a percentage of the control (DMSO) after exposure to B[a]P (100 nM) for 3 or 24 h. Data are presented as mean ± SEM from at least three independent experiments. Statistical analyses were performed using one-way ANOVA for panels a, b, and e, and Student's t-test for panels c and d. Significant differences compared to the corresponding control (DMSO) are indicated as follows: \* or \#, *p* < 0.05; \*\* or \*\*\*, *p* < 0.01; \*\*\* or \*\*\*\*, *p* < 0.001. Abbreviations: α-NF, α-naphthoflavone; AIF, apoptosis-inducing factor; COX IV, cytochrome c oxidase subunit IV; CSA, cyclosporin A; DRP1, dynamin-related protein 1.</figcaption></figure>

consistent with the literature, indicating that inhibition of autophagic flux promotes apoptosis (Boya et al., 2005).

In the presence of B[a]P, we observed a decrease in the expression levels of DRP1 and MFF, two key proteins of mitochondrial fission. This decrease, detected as early as 3 h of exposure, is accompanied by mitochondrial elongation (decrease in circularity index and increase in perimeter), revealing an activation of mitochondrial dynamics. The ability of B[a]P to modulate this process has already been reported, but at much higher concentrations (5–10 µM) (Paing et al., 2024). Using B[a]P at concentrations closer to those encountered in humans, our results confirm that alterations in mitochondrial dynamics, and more specifically mitochondrial elongation, could constitute a relevant early cellular event in low-dose exposure to B[a]P.

We also observed an increase in ATP synthesis, a phenomenon typically associated with mitochondrial elongation. Thus, in the presence of low doses of B[a]P, HMEC-1 cells exhibit typical features of SIMH: decreased expression of fission proteins, mitochondrial elongation, and increased ATP synthesis. SIMH has been described in response to various stressors, particularly when these stressors are of low intensity, as in our case, or involve oxidative stress. SIMH is a transient survival response (Brown et al., 2025; Shutt and McBride, 2013; Zahedi

et al., 2019). Our results confirm this transient nature: the decreased expression of DRP1 and MFF, the mitochondrial elongation, and the increased ATP synthesis were primarily observed at early time points in HMEC-1 cells exposed to B[a]P, before the detection of apoptosis. These observations support the hypothesis that SIMH represents an attempt to delay cell death by apoptosis.

Among all the mitophagy markers, only NDP52, described as a redox sensor (Kataura et al., 2023), showed a tendency toward increased expression in the presence of B[a]P, although this change was not significant. It is noteworthy that NDP52 expression is known to be regulated by the transcription factor NRF2, a key component of the cellular antioxidant response (Jo et al., 2014). NRF2 directly regulates NDP52 transcription through antioxidant response elements located in its promoter (Pajares et al., 2016). NRF2 also regulates the expression of certain mitochondrial dynamics actors such as DRP1 (Sabouny et al., 2017). Moreover, NRF2 activators, such as resveratrol, reduce DRP1 expression and induce mitochondrial elongation (Ren et al., 2017). The induction of NDP52, the decrease in DRP1, and the mitochondrial morphological changes we observed in the presence of B[a]P thus support the activation of the NRF2 pathway. In line with this, we detected an NRF2 accumulation in HMEC-1 cells exposed to B[a]P for 3 h, a
<figure><img src="image_15.png" /><figcaption>**Fig. 8.** B[a]P-induced NRF2 activation modulates apoptosis in HMEC-1 cells. a) Cellular NRF2 levels were assessed by Western blotting of HMEC-1 lysates after exposure, or not (DMSO), to 100 nM B[a]P for 3 h or 24 h. The blot shown is representative of three independent experiments. b,c) Apoptosis was quantified in HMEC-1 cells exposed, or not (DMSO), to 100 nM B[a]P for b) 24 h in the presence or absence of the NRF2 activator sulforaphane (5 µM), or c) 3 h in the presence or absence of the NRF2 inhibitor brusatol (80 nM). Apoptosis was scored by microscopic detection of condensed and/or fragmented chromatin after Hoechst staining. Data are means ± SEM from three independent assays, normalized to the DMSO control (set to 100%; horizontal dashed line). Statistical significance: **p < 0.01; ***p < 0.001 vs. DMSO; #p < 0.05, ##p < 0.01 vs. B[a]P alone; ⌧p < 0.05 vs. brusatol alone.</figcaption></figure>

period preceding detectable apoptosis. NRF2, SIMH, and delayed apoptosis, therefore, appear to be associated elements of the cellular response. This hypothesis is further supported by the fact that in the presence of an NRF2 inhibitor, apoptosis becomes detectable as early as 3 h of B[a]P exposure. Conversely, an NRF2 activator completely inhibits apoptosis at 24 h of exposure. Further investigations using a molecular approach should allow a definitive conclusion on the involvement of NRF2.

## 5. Conclusion

Even at low concentrations, the mitochondrion remains a sensitive intracellular organelle to B[a]P toxicity. To more thoroughly assess the mitochondrial cellular response, we explored the three key components of mitochondrial homeostasis: biogenesis, mitophagy, and mitochondrial dynamics. B[a]P bioactivation induces mitochondrial dysfunction and an early survival response relying on mitochondrial dynamics characteristic of SIMH. Mitochondrial biogenesis does not appear to be induced, and mitophagy seems compromised. In HMEC-1 cells exposed to low doses of B[a]P, we observe NRF2 accumulation as well as a decrease in the expression of the fission proteins DRP1 and MFF. This early response is accompanied by mitochondrial hyperfusion (due to impaired fission) and increased cellular ATP content. These SIMH features, while early and transient, allow the cell to delay apoptosis (seen at 24 h of exposure). Altogether, our findings suggest that, among the three determinants of mitochondrial homeostasis, alterations in mitochondrial dynamics represent an early signal of a cellular response to low doses of B[a]P.

## CRediT authorship contribution statement

**Joan Guillouzouic:** Writing – review & editing, Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization.
**Camille Chauvin:** Writing – review & editing, Validation, Data curation.
**Valentine Genet:** Writing – review & editing, Validation, Data curation.
**Simon Martarello:** Writing – review & editing.
**Agnes Burel:** Writing – original draft, Validation.
**Lydie Sparfel:** Writing – review & editing.
**Normand Podechard:** Writing – review & editing.
**Dominique Lagadic-Gossmann:** Writing – review & editing.
**Catherine Lavau:** Writing – review & editing, Funding acquisition, Conceptualization.
**Eric Le Ferrec:** Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Data curation, Conceptualization.

## Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Eric Le FERREC reports financial support was provided by French National Research Agency (ANR-23-CE34-0001 (ENDOMITOPAH)). Joan Guillouzouic reports financial support was provided by Brittany Region. Joan Guillouzouic reports financial support was provided by Rennes Faculty of Pharmacy. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.envpol.2025.127538.

## Data availability

Data will be made available on request.

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