Abstract

   Cadmium (Cd), a carcinogenic component of tobacco, is a recognized risk
   factor for oral squamous cell carcinoma (OSCC). However, the molecular
   mechanisms underlying Cd-induced cytotoxicity in OSCC remain largely
   undefined. Here, we demonstrate that acute Cd exposure triggers
   ferroptosis in CAL27 OSCC cells derived from never-smokers, but not in
   SCC154 cells derived from smokers. Mechanistically, Cd outcompetes Fe,
   causing early iron depletion and activating the nuclear receptor
   coactivator 4 (NCOA4)-mediated ferritinophagy. This process enhances
   the labile iron pool, promotes mitochondrial reactive oxygen species
   (ROS) generation, lipid peroxidation, and ferroptotic cell death.
   Notably, iron supplementation rescues CAL27 cells from Cd-induced
   damage, while exacerbating iron deficiency through transferrin receptor
   CD71 silencing amplifies cytotoxicity. Conversely, OSCC cells from
   smokers exhibit resistance to Cd toxicity, likely due to the
   overexpression of metallothionein 2A (MT2A), a heavy metal
   detoxification protein. Collectively, this study provides the evidence
   that ferritinophagy may act as a critical upstream driver of Cd-induced
   ferroptosis in OSCC cells derived from never-smokers, paving the way
   for potential ferroptosis-targeted therapeutic strategies in
   Cd-associated malignancies.

   Keywords: Cadmium, Ferroptosis, Ferritinophagy, NCOA4, Oral Cancer,
   Iron Metabolism, Smokers

Introduction

   The homeostasis of intracellular metal ions is essential for
   maintaining cellular integrity and function. However, disturbances in
   metal balance can trigger a cascade of toxic events, including
   oxidative damage to proteins and DNA, disruption of cell membranes, and
   activation of regulated cell death (RCD) pathways [59]^1^,[60]^2. In
   recent years, a growing body of evidence has delineated distinct forms
   of metals-induced RCD, namely ferroptosis, cuproptosis, and
   calcicoptosis, each characterized by the accumulation of specific
   divalent cations, namely iron (Fe²⁺), copper (Cu²⁺), and calcium
   (Ca²⁺), respectively [61]^3^,[62]^4. In addition, exogenous metals such
   as zinc (Zn²⁺), manganese (Mn²⁺), and cadmium (Cd²⁺) have been shown to
   induce cell death through context-dependent mechanisms, often
   converging on oxidative stress and mitochondrial dysfunction
   [63]^5^,[64]^6.

   Cd^2+ is a well-recognized environmental pollutant, primarily
   originating from industrial processes, agricultural activities, and
   tobacco-consumption [65]^5^,[66]^7^-[67]^9. Although Cd^2+ is a
   non-Fenton-like metal and does not directly generate reactive oxygen
   species (ROS), it can induce oxidative stress through several indirect
   mechanisms [68]^10^,[69]^11. These include (i) depletion of antioxidant
   molecules such as glutathione (GSH), (ii) inhibition of ROS-detoxifying
   enzymes, (iii) displacement of essential redox-active metals (Zn²⁺ and
   Fe²⁺), and (iv) impairment of mitochondrial electron transport chain,
   collectively resulting in mitochondrial dysfunction and ROS
   overproduction [70]^10^-[71]^13.

   Cd²⁺-induced oxidative stress has been recognized as a key driver of
   several pathological conditions, including cancer [72]^12^-[73]^17. In
   1993, indeed, the International Agency for Research on Cancer (IARC)
   classified Cd²⁺ as a Group 1 carcinogen [74]^18. Mechanistically, Cd^2+
   exerts its carcinogenic activity through multiple pathways, including
   the induction of oxidative DNA damage in the form of DNA mutation,
   strand breaks, and chromosomal aberrations, as well as the inhibition
   of DNA repair systems - notably through suppression of p53 DNA binding
   capacity and the suppression of DNA repair-associated genes
   [75]^19^-[76]^21. Beyond its genotoxic effect, Cd^2+ also exerts
   epigenetic effects by altering DNA and histone methylation patterns.
   For instance, Cd^2+-induced hypermethylation of tumor suppressor genes
   promoters, such as p16, has been associated with malignant
   transformation of human prostate epithelial cells [77]^22. Furthermore,
   chronic exposure to sub-toxic concentrations of Cd^2+ can activate
   defense mechanisms against oxidative stress, including the stimulation
   of ROS-sensitive transcription factors, such as nuclear factor
   erythroid 2-related factor 2 (Nrf2), activator protein 1 (AP-1) and
   nuclear factor-kB (NF-kB), as well as mitogen-activated protein kinases
   (MAPKs)- dependent signaling pathways, which may ultimately promote
   cell survival and tumorigenesis [78]^23^-[79]^25.

   Over time, epidemiological studies have reported a significant
   association between Cd²⁺ exposure and increased risk of oral squamous
   cell carcinoma (OSCC), the most common subtype of head and neck
   squamous cell carcinoma (HNSCC) [80]^26^-[81]^28. In this context, both
   environmental factors and genetic alterations in oncogenes and tumor
   suppressor genes play central roles in OSCC pathogenesis
   [82]^29^-[83]^33. Notably, chronic and prolonged exposure to Cd^2+ -
   particularly through tobacco consumption - appears to contribute to
   carcinogenic transformation of the oral epithelial mucosa
   [84]^9^,[85]^30^,[86]^34^-[87]^37. However, the molecular mechanism
   underlying the Cd^2+-induced toxicity in oral epithelial cells remains
   incompletely understood and requires further study.

   In this study, we investigated the effects of Cd²⁺ acute exposure in
   OSCC cells derived from non-smoker and smoker patients - the latter
   being chronically exposed to roughly 4-5 times higher levels of Cd²⁺
   compared to non-smokers. Our findings reveal that Cd²⁺ toxicity
   selectively affects OSCC cells derived from non-smokers whereas OSCC
   cells derived from smokers display resistance, likely due to the
   overexpression of the heavy metal detoxification protein
   metallothionein 2A (MT2A). Notably, we demonstrate for the first time
   that, in OSCC cells derived from non-smokers, ferroptosis is involved
   in Cd²⁺-induced cytotoxicity. Mechanistically, Cd^2+ outcompetes Fe,
   thus leading to an early iron depletion, which in turn acts as a
   driving force for the nuclear receptor coactivator 4 (NCOA4)- mediated
   autophagic degradation of ferritin (ferritinohagy). Ferritinophagy,
   subsequently, determines an increase in labile iron pool (LIP),
   mitochondrial ROS production, and lipid peroxidation. Overall, this
   study uncovers a novel mechanism of Cd-induced cytotoxicity in OSCC
   cells, providing a basis for developing ferroptosis-based therapeutic
   strategies for Cd-associated diseases.

Materials and Methods

Cell lines and cell culture

   Human oral squamous cell lines (OSCC) - CAL27, OT1109, SCC090, and
   SCC154 - were purchased from the American Type Culture Collection
   (ATCC, Rockville, MD, United States). CAL27 and OT1109 cells were
   derived from never-smoker patients, while SCC090 and SCC154 originated
   from tobacco users. Following ATCC instruction, CAL27 cells were grown
   in DMEM medium (Sigma-Aldrich, St. Louis, Missouri, United States ),
   while SCC154 cells were cultured in MEM (Sigma-Aldrich, St. Louis,
   Missouri, United States), both supplemented with 10% (v/v) fetal bovine
   serum (FBS) (Invitrogen, San Diego, CA), L-glutamine and 1% (v/v)
   penicillin and streptomycin (Sigma-Aldrich, St. Louis, Missouri, United
   States) at 37°C in a humidified incubator with 5% CO[2] atmosphere. All
   cell lines were tested for mycoplasma contaminations and authenticated
   via short tandem repeat (STR) profiling.

Reagents and treatments

   Cadmium chloride (CdCl[2]), ferrostatin-1 (Fer-1) and bafilomycin (Baf)
   were purchased from Sigma Aldrich (Sigma-Aldrich, St. Louis, MO, USA).
   Ferlixit (62.5 mg/5 mL, sodium ferric gluconate complex in sucrose,
   SANOFI) has been obtained from the outpatient pharmacy at the Unit of
   Cardiology, “Magna Graecia” University of Catanzaro. Cells were seeded
   in a 12- and 6-well plate in complete medium. Each compound was used at
   the following final concentrations: CdCl[2] at 0.1, 1, 5, 10, 50 and
   100μM for 12h; CdCl[2] at 26.1μM for 30', 1h, 6h and 12h; Fer-1 at
   100μM for 24h; Baf at 1μM for 12h; ferlixit at 25, 50 and 100μM for
   12h. Treatments were performed at least three times on independent
   biological replicates. CAL27 were exposed to 10μM CdCl[2] for 30 days
   to induce metal tolerance (CAL27T); this concentration was replenished
   every 2-3 passages to maintain tolerance.

Patients and clinical samples

   Fourteen OSCC patients, 7 non-smokers and 7 smokers, underwent surgery
   at the Oral Pathology and Oral Surgery Unit of “Magna Graecia”
   University, between December 2020 and December 2022 [88]^32^,[89]^33.
   For each patient, primary tumor tissue specimens were collected within
   the macroscopic lesion boundaries defined visually and by palpation.
   All patients provided a written informed consent at the time of data
   collection. No information that could identify individual participants
   are available. The procedures reported in this study were performed in
   accordance with the Helsinki Declaration guidelines (2008) on human
   experimentation and good clinical practice (good clinical practice or
   GCP).

PI staining analysis

   Cells were incubated with propidium iodide (PI) at 37°C for 15 min in
   the dark, washed twice with PBS, and analyzed using a BD LSRFortessa™
   X-20 flow cytometer (BD Biosciences, San Jose, CA, USA). A total of
   2×10^4 events were acquired for each sample. Data analysis was carried
   out using FlowJo™ v10 Software (BD Biosciences, San Jose, CA). Each
   experiment was performed in triplicate.

Apoptosis assay

   Apoptotic cells populations were identified using the Alexa Fluor®488
   Annexin V/Dead Cell Apoptosis Kit (Thermo Fisher Scientific, Waltham,
   MA, USA [90]^38. Briefly, 1×10^5 single-cell suspensions from CAL27 and
   SCC154 cell lines were centrifuged and resuspended in 100μL 1X
   annexin-binding buffer. To each sample, 5μL Alexa Fluor®488 Annexin V
   and 1μL PI working solution (100μg/mL) were added. Samples were then
   incubated for 15' at room temperature in the dark. Each tube was
   diluted with 200 μL of Annexin Binding Buffer. Flow cytometry assays
   were performed using the BD LSRFortessa™ X-20 (BD Biosciences, San
   Jose, CA, USA). Data were acquired from three independent biological
   replicates and analyzed out using FlowJo™ v10 Software (BD Biosciences,
   San Jose, CA).

Cell viability assay (MTT)

   Cell viability was assessed using the
   3-[4,5-dimethylthiazolyl]-2,5-diphenyltetrazolium bromide (MTT) assay
   (Sigma-Aldrich, St. Louis, MO, USA) assay. Briefly, CAL27 and SCC154
   cells (5 × 10^4 cells/well) were seeded in a 24-well plates. Following
   exposure to CdCl[2], cells were incubated with freshly prepared MTT
   solution (2 mg/mL) for 4h at 37 °C. Then, the supernatant was removed
   and replaced with 200μL of isopropanol to solubilize the resulting
   formazan crystals. Absorbance was measured at 595nm using a microplate
   spectrophotometer. Cell viability was expressed as a percentage
   relative to untreated control cells, which were set as 100%. The assay
   was performed at 0, 12, and 24h post-treatment. All experimental
   conditions were tested in triplicate across three independent
   experiments.

Wound healing assay

   Cells (3 × 10⁵) were seeded in 12-well plates. A scratch was introduced
   using a sterile pipette tip, and wound closure was monitored at 0, 12,
   24, 48, and 72 h using using the Leica THUNDER Microscope DMi8 (Leica
   Microsystems S.r.l., Wetzlar, Germany). The gap area was quantified
   using by using ImageJ software. All experiments were conducted in
   triplicate.

Total protein extraction and western Blot analysis

   Total protein extracts were prepared using RIPA lysis buffer composed
   of 1M Tris HCl, Triton X-100, 3M NaCl, 0.5M EDTA, 10% SDS supplemented
   with cOmplete™ Protease Inhibitor Cocktail provided in EASYpacks (Roche
   Diagnostics, Mannheim, Germany) to prevent proteolytic degradation
   [91]^39. Briefly, cells were lysed in ice-cold RIPA buffer and lysates
   were centrifuged at 12.000g for 30' at 4°C to remove insoluble debris.
   Protein concentration was determined using the Bio-Rad Protein Assay
   Dye according to manufacturer's instructions (Bio-Rad Laboratories,
   Hercules, California, United States). Equal amounts of protein (50μg)
   from each sample were separated by 8%-12% SDS-PAGE, run at 200V for 1h
   and 30'. Proteins were then transferred onto nitrocellulose membranes
   (Sigma-Aldrich, St. Louis, MO, United States) at 50V for 2h. Membranes
   were blocked with 5% non-fat milk or 5% BSA for 1h at room temperature,
   followed by overnight incubation at 4°C with the appropriate primary
   antibodies. The antibodies against ferritin heavy subunit (FtH1)
   (1:200, sc-376594), NCOA4 (1:500, sc-373739) and hypoxia inducible
   factor-1 alpha (HIF-1ɑ) (1:500, sc-10790) were purchased from Santa
   Cruz Biotechnology (Santa Cruz Biotechnology, Dallas, Texas, United
   States); antibody against glutathione peroxidase 4 (GPX4) (1:1000,
   ab19534) was purchased from Abcam (Abcam, Cambridge, UK), while
   antibodies against mechanistic target of rapamycin complex 1 (mTORC1)
   (1:500, 2972s), phosphorylated mTORC1 (p-mTORC1) (1:500, 5536s),
   microtubule associated protein 1 light chain 3B (LC3B) (1:500, #2775)
   and iron regulatory protein 1 (IRP1) (1:1000, 20272) were obtained from
   Cell Signaling Technology (Danvers, Massachusetts, United States).
   Membranes were washed for 30' and then incubated for 1h at room
   temperature with peroxidase-conjugated secondary antibodies (Peroxidase
   AffiniPure Sheep Anti-Mouse IgG, 1:10,000; Peroxidase AffiniPure Donkey
   Anti-Rabbit IgG, 1:10,000; Peroxidase AffiniPure Donkey Anti-Goat IgG,
   1:10,000; Jackson ImmunoResearch Europe Ltd). Signals were detected
   using chemiluminescence reagents (ECL Western blotting detection
   system, Santa Cruz Biotechnology, Dallas, Texas) and acquired by Uvitec
   Alliance Mini HD9 (Uvitec Cambridge, United Kingdom). To calculate the
   relative expression of specific protein a mouse monoclonal IgG
   glyceraldehyde 3-phosphate dehydrogenase (GAPDH) HRP (1:3000; sc-47724)
   serves as references for sample loading. The protein band intensity on