Acrylamide-mediated errors in the cell cycle regulation are associated with altered TORC2 signaling in Schizosaccharomyces pombe

Abstract

Acrylamide (AA) poses a significant risk to living organisms as it is linked to serious health concerns. AA exposure triggers oxidative stress in cells through elevated ROS and modulation of antioxidant enzymes activities and expression of genes encoding antioxidant enzymes. AA-induced cell proliferation defects are linked to affected cell cycle regulation demonstrated by changes in the expression of genes encoding the major cell cycle regulators cdc2, cdc13, and cdc25, Additionally, cell division defects can be linked to changes in the expression of ark1 and cdc15, and AA-induced errors in chromosome segregation. The stress response involves signaling pathways like MAPKs (Mitogen-activated protein kinases) or the target of rapamycin (TOR) constituting two complexes TORC 1 and 2. As TORC2 manages the cell response to various stresses, its involvement in AA-mediated stress has been demonstrated by changes in the expression of tor1, wat1, ste20, sin1, bit61 encoding TORC2 members, and gad8 encoding a direct Tor1substrate, Gad8. To our surprise, AA has not affected the expression of sty1, which encodes the major stress-regulating kinase of the MAPK pathway in S. pombe. In the presented study we demonstrate, for the first time, that exposure to AA disrupts cellular homeostasis by altering TORC2 signaling and cell cycle regulation ultimately leading to carcinogenesis.

Introduction

The presence of acrylamide (AA), a possible human carcinogen, in significant quantities in a wide range of commonly consumed human foods, raises serious concerns about its potential health risks. This harmful compound is commonly found in carbohydrate-rich heat-processed foods such as chips, bread, or breakfast cereals, but also coffee, at high levels (150–5000 µg/kg)¹, which are higher than other well-known food carcinogens. Although the maximum limits for AA in food have not been established, according to the World Health Organization (WHO) guideline for AA in drinking water, the amount considered as not harmful is 0.5 µg/L². The average daily intake of AA in adults is estimated to be 0.6 µg/kg of body weight per day, but this value can range widely from 0.2 to 1 µg/kg of body weight per day making the intake of AA not optimistic^3,4.

Although cell toxicity and genotoxicity of AA have been well documented in numerous experimental studies^{5, 6, 7–8}, an in-depth systematic investigation of its perturbations with multiple nodes within the cellular signaling network is yet to be clarified.

The toxic effect of the food-derived AA is primarily mediated through its ability to induce oxidative stress⁹. Reactive oxygen species (ROS), like superoxide radical anion (O₂^•−), hydroxyl radicals (OH^•), and hydrogen peroxide (H₂O₂) are formed as a result of energy and redox imbalance in cells¹⁰. Cells exposed to oxidative stress experience degradation of lipid components in biomembranes¹¹, protein oxidation¹², nucleic acid damage, and mutations^10,13 which consequently damages the cell and accelerates aging. During evolution, cells have developed sophisticated defence mechanisms against ROS toxicity through scavenging pathways, manifested by both enzymatic and non-enzymatic antioxidant action. Antioxidants that counteract ROS to facilitate their physiological functions while minimizing the oxidative damage they cause include enzymes such as superoxide dismutase (SOD)¹⁴, catalase (CAT)¹⁵, and thiol-dependent peroxidases like glutathione peroxidase (GPx) or thioredoxin peroxidase (TRx)^16,17. In addition, cell detoxification from ROS utilizes glutathione, which is converted by glutathione reductase (GR) from its oxidized form to the reduced form. This mechanism is essential for cell survival under stress¹⁸. Stress generally leads to an increase in the activity of antioxidant enzymes as a response to elevated ROS. However, various studies have indicated that the enzyme activity may decrease depending on the type, strength, and duration of the stress^19,20. Thus, it is still an open question whether the modulation of ROS levels during stress is due to changes in enzyme activity or the amount of antioxidant enzymes. Another consequence of increased ROS formation is the disturbance in cell metabolism mediated by ROS, leading to cell growth arrest and ultimately resulting in cell death²¹. It has been demonstrated that cells exposed to AA exhibit growth retardation and delayed cell cycle progression^{7,22, 23–24}. However, the impact of AA on cell cycle regulation is still poorly understood.

The fission yeast Schizosaccharomyces pombe is a popular model system for biological studies, frequently used to investigate the toxicity of harmful compounds, including AA, on eukaryotic organisms at a molecular level^7,25. In particular, cell cycle regulation has been extensively studied using S. pombe as a model organism²⁶. The regulation of the cell cycle is under strict control, when the cell reaches a certain size, it undergoes mitosis followed by cell division²⁷. The major regulator of the cell cycle in S. pombe is the cyclin-dependent kinase, Cdc2. It is associated with specific cyclins at different cell cycle stages and requires a coordinated series of phosphorylation and dephosphorylation events to ensure accurate cell cycle progression²⁸. The control system is crucial for proper chromosome segregation during mitosis, ensuring equal distribution of genetic material to daughter cells and allowing each phase of the cell cycle to follow the next one smoothly²⁹. In addition, correct cell division can only occur when spindle microtubules are properly attached to kinetochores. Kinetochore biorientation is crucial for the error-free interaction of kinetochores with microtubules in a bi-polar manner. This is accomplished by the Aurora B kinase, which phosphorylates kinetochore components to activate the spindle assembly checkpoint (SAC) which is crucial for accurate chromosome segregation^30,31. The final step in the cell division cycle, resulting in the physical separation of the newly formed daughter cells, is cytokinesis. The F-BAR protein Cdc15, an essential protein for cytokinesis, facilitates the attachment of the cytokinetic ring (CR) to the plasma membrane and directs the formation of a division septum³². Such multi-step regulation may be disrupted by AA-induced stress, leading to error-prone cell cycle progression by interfering with regulatory molecules at various stages of the cell division cycle. To prevent this, cell defence mechanisms utilize MAPKs (Mitogen-activated protein kinases) and the target of rapamycin (TOR) signaling pathways to mitigate stress. In S. pombe, various environmental stimuli activate Sty1 kinase, which in turn triggers the activation of MAPKs to arrange intercellular signaling³³. In addition, cell viability under stress conditions largely depends on the TOR signaling. TOR is a serine/threonine protein kinase that plays a key role in modulating cell growth in response to different environmental conditions^{34, 35–36}. TOR is present from yeast to humans in two multiprotein complexes, TORC1 containing Tor2, and TORC2 containing Tor1 as the major catalytic subunit in S. pombe. Of the two complexes, rapamycin directly inhibits TORC1, whereas TORC2 is not affected. While the TORC1 kinase complex is vital for the vegetative growth of S. pombe cells, TORC2 is not essential. However, TORC2 is required for survival under stress conditions, proper G1 arrest, and sexual development^37,38. In addition, TORC2 has been shown to influence the assembly and function of the actin cytoskeleton, regulate cell wall integrity, and guide the successful completion of cytokinesis^39,40. Moreover, TORC2 plays a critical role in regulating chromatin metabolism and is involved in the propagation of mitosis, especially following recovery from DNA-damaging conditions^40,41. TORC2 signaling is mediated through its subunits Sin1, Ste20, Wat1, Bit61, the primary catalytic subunit Tor1 kinase, and its target, the AGC kinase Gad8 in S. pombe⁴².

We have previously shown that oxidative stress induced by AA leads to cell cycle regulation changes. In this study, we present that exposure to AA not only increases the production of individual ROS and disrupts the function of antioxidant enzymes, but also induces modifications in the expression of genes encoding antioxidant enzymes and molecules. In addition, errors in the cell cycle progression are mediated through AA-induced downregulation of genes involved in the regulation of cell division. Finally, we demonstrate that AA-induced cell disruptions are mainly caused by changes in TORC2 signaling rather than MAPK signaling in the fission yeast.

Materials and methods

Yeast cultivation and growth conditions

In all studies the wild type Schizosaccharomyces pombe strain SP72 h + ade6-M210 ura4-D18 leu 1–32 was used. For the immunostaining experiment, the JG 15,457 strain (cen2(D107)::KanR-ura4+-lacO his7+::lacI-GFP) carrying chromosome II labeled with GFP was used. Yeast cells were cultured as described by⁷. Briefly, yeasts were cultured in a complete YES liquid medium (0.5% yeast extract, 3% glucose, supplemented with adenine, L-histidine, L-leucine, L-lysine, and uracil), at 30 °C, and 150 rpm.

Growth intensity, generation time, and IC₅₀ value determination

Cells from the overnight (o/n) culture were diluted to the OD₆₀₀ = 0.3, transferred to 24 well plates, and exposed to indicated AA concentrations, 0, 0.1, 1, 10, 20, and 60 mM. Cells were incubated at 30 °C and 150 rpm for 9 h and changes in optical density at 600 nm were measured every 3 h by Glomax Multi Detection System (Promega Corporation, Madison, WI, USA). Since a single life cycle of S. pombe takes approximately three hours, the intensity of cell growth following the addition of AA was examined for three life cycles. The cell density at three, six, and nine hours is compared to the 0-hour time point to determine the total growth intensity (OD₆₀₀ ratio).

Generation time (gt) represents cell doubling time and is calculated by Eq. (1):

1

where m is the gradient of the regression line.

IC₅₀ value represents the concentration of AA that results in a 50% reduction in cell growth. Serial dilution of AA was used for IC₅₀ calculations. Cells without (w/o) AA treatment were used as a control. IC₅₀ was calculated using a four-parameter logistic regression using R software.

Characterization of the yeast morphology

Yeast morphology was determined as previously described⁷. Briefly, cells incubated with AA for 6 h were visualized with the use of bright-field microscopy at 40× magnification (Leica DMI 6000, Leica microsystems, Wetzlar, Germany). Using the ImageJ software version 1.52r, microscopy images were taken and cell morphometric analyses were performed (National Institutes of Health, USA).

Cell volume (V; µm³) was calculated according to Eq. (2):

2

where L represents the cell length, and W is the cell width.

Cell surface (S; µm²) was calculated according to Eq. (3):

3

where factor ε is calculated as .

Cell septation was analyzed in cells prepared as described above, incubated with calcofluor white and visualized under the fluorescence microscope (Leica DMI 6000, Leica microsystems, Wetzlar, Germany) at 40× magnification. The ratio of cells with septum was determined.

Preparation of the whole cell extract for biochemical analyses

Cell preparation for biochemical analyses was performed as previously described⁷. Briefly, control and AA-treated cells were washed and resuspended in PBS (pH 7.0). Afterwards, cells were homogenized by sonication (Digital Sonifier 450, Branson Ultrasonics Corp, Danbury, CT, USA) at 3 × 15 s intervals on ice in 1.0 s pulse interval, centrifuged 15 min at 14,000 g and 4 °C to remove cell debris. Protein level, MDA content, ROS generation, H₂O₂ formation, respiration, CAT, SOD, and GPx activities were determined in the collected supernatant.

Respiration activity

Respiration activity (RA) was calculated as described by⁷. Briefly, yeast cells were washed with PBS (pH 7.0) by 90 s centrifugation at 10,000 g, resuspended in 1 mL of 0.5% 2,3,5-triphenyltertrazolium chloride (TTC) diluted in PBS pH 7.4 and incubated 20 h in the dark at 30 °C. Cells were then washed twice with PBS, and generated red formazan was extracted by addition of 1 mL ethanol: acetone (2:1) mixture prior to cell lysis by sonication. Absorbance was measured at 485 nm by the Agilent Cary 60 UV/VIS spectrophotometer (Agilent Technologies, Santa Clara, CA, USA) and metabolic activity was calculated as relative absorbance units (r.u.) per milligram (mg) of protein.

Biochemical analysis

CAT activity was determined as previously described⁷. Briefly, the stepwise decrease in absorbance at 240 nm measured by Agilent Cary 60 UV/VIS spectrophotometer (Agilent Technologies, Santa Clara, CA, USA) over 90 s, indicates the H₂O₂ degradation, and represents CAT activity. The molar absorption coefficient 36 mM⁻¹ cm⁻¹ was used to calculate specific catalase activity.

SOD activity was determined as previously described⁷. Briefly, homogenized Sample was mixed with 50 mM phosphate buffer (pH 7.8) containing 1 mM EDTA, 13 mM L-methionine, and 75 µM NBT (nitroblue tetrazolium). Prior to reaction initialization by light irradiation (5000 lx) for 10 min at 20 °C, 2 mM of riboflavin was added. Absorbance was measured at 560 nm by the Agilent Cary 60 UV/VIS spectrophotometer (Agilent Technologies, Santa Clara, CA, USA).

GPx activity was determined as coupled to NAPDH oxidation via glutathione reductase enzyme according to⁴³ with a slight modifications. The reaction solution consisted of 50 mM Tris-HCl buffer (pH 7.6), 0.30 mM reduced glutathione, 0.09 mM EDTA, 0.15 mM NADPH, 0.25 mM H₂O₂, and 1 unit of glutathione reductase. GPx activity was measured spectrophotometrically at 340 nm using the Agilent Cary 60 UV/VIS spectrophotometer (Agilent Technologies, Santa Clara, CA, USA) at lab temperature.

Malondialdehyde content (MDA) was evaluated as previously described⁷. Briefly, the TBA solution [15% trichloroacetate (TCA) and 0.375% (w/v) thiobarbituric acid (TBA)] was added to each Sample, and incubated at 95 °C for 30 min. Then the Sample was cooled on ice, centrifuged at 8500 rpm for 60 s, and the absorbance was measured at 532 nm and 600 nm by the Agilent Cary 60 UV/VIS spectrophotometer (Agilent Technologies, Santa Clara, CA, USA). The molar absorption coefficient 153 mM⁻¹ cm⁻¹ was used to calculate MDA content in nmol µg⁻¹ protein.

Glutathione (GSH) concentration was determined using a GSH assay kit (Sigma Aldrich; CS0260). Briefly, an overnight yeast culture, adjusted to an optical density (OD) of 1.0 at OD600, was treated with 0, 1, 10, and 20 mM AA and incubated at 30 °C for either 3–6 h. Afterwards, 2 mL of the culture was collected and centrifuged at 5000 rpm for 5 min, the cell pellet was washed three times with PBS at pH 7.4. A repetitive freeze-thaw process (liquid nitrogen and 37 °C, repeat 3x) was used to disrupt the cell pellet. Subsequently, 5% sulfosalicylic acid was added to the pellet, and the Sample was centrifuged at 14,000 rpm for 10 min. Five microliters of each Sample were transferred to a 96-well plate and measured using a kinetic read mode for five minutes at 412 nm on a Varioskan Lux multimode microplate reader (Thermo Fisher Scientific, Inc., Waltham, MA, USA). To calculate GSH concentration, a standard curve of glutathione was generated according to the manufacturer’s instructions.

Protein concentration was determined at 600 nm by the Bradford assay⁴⁴ with bovine serum albumin (Sigma-Aldrich, St. Louis, MO) used as a standard at the Glomax Multi Detection System (Promega Corporation, Madison, WI, USA).

Total ROS formation was analyzed as previously described by⁴⁵ with slight modifications. Cell concentration in all samples was adjusted to OD₆₀₀ = 1, cells were washed with PBS pH 7.4, and incubated with 10 µM H₂DCFDA (Sigma-Aldrich) at 30 °C in the dark for 1 h. ROS interaction with H₂DCFDA forms highly fluorescent DCF detectable at 498 nm. A filter with 490 nm excitation wavelength and 510–570 nm emission wavelength was used in the Glomax Multi Detection System (Promega Corporation, Madison, WI, USA). Values were normalized to protein content.

Superoxide anion (O₂^•−) content was determined according to⁴⁶ with minor modifications by the chemiluminescent method, using the Superoxide Anion Assay Kit (CS1000, Sigma Aldrich). Yeast samples were stored in PBS (pH 7.8) in a deep-freeze box at − 80 °C until analysis. After thawing, the cells were collected by centrifugation at 10,000 rpm for 15 min, and 4 °C. Ice cold PBS (pH 7.8) was added to the pellet in a volume of 1000 µL, followed by sonication (3 × 15 s intervals on ice in 1.0 s pulse interval). After centrifugation (10000 rpm, 15 min, 4 °C), 100 µL of the supernatant was added to the microplate well containing assay solution (according to the manufacturer instruction) to a final volume of 200 µL. Changes in the luminescence intensity, representing O₂^•− concentration, were determined by Glomax Multi Detection System (Promega Corporation, Madison, WI, USA) with the luminescence mode at 540 nm for 5 min at 30 s intervals. Results were normalized to the protein content.

Hydrogen peroxide (H₂O₂) content was determined with the use of hydrogen peroxide assay kit (MAK165, Sigma Aldrich) according to the manufacturer instructions with minor modifications. A 100 µL of the cell lysate was added to the black 96-well microplate containing 25 µL of the assay buffer (comprising red peroxidase substrate) at a dim light. The mixture was incubated at room temperature for 30 min, in the dark. Fluorescence intensity at excitation/emission of 490 nm/510–570 nm, determining the H₂O₂ concentration, was measured using the Glomax Multi Detection System (Promega Corporation, Madison, WI, USA). Results were normalized to the protein content.

Hydroxyl radical (OH^•) concentration was determined by the fluorescence method using the selective probe OH580 (ab219931; Abcam) according to⁴⁷ with minor modifications. 100 µL of the cell lysate was added to the well of a black 96-well microplate containing 100 µL of the reaction buffer at dim light. The mixture was incubated at room temperature for 60 min, in the dark. Fluorescence intensity at excitation/emission of 525 nm/580–640 nm, determining the OH^• concentration, was measured using the Glomax Multi Detection System (Promega Corporation, Madison, WI, USA). Results were normalized to the protein content.

Detection of apoptosis and necrosis

Evaluation of the yeast cells apoptosis was performed according to⁷. Briefly, Annexin V-FITC (fluorescein isothiocyanate)-conjugated specifically binds to phosphatidylserine (PS) residues, and is used as an apoptosis marker. Propidium iodide (PI) (Sigma Aldrich) penetrates dead cells and serves as a marker to differentiate between apoptosis and necrosis. Cells exposed to AA for 1 h were washed twice with PBS, pH 6.8 and resuspended in sorbitol buffer (1.2 M sorbitol, 0.5 mM MgCl₂, and 35 mM K₂HPO₄), pH 6.8 to a final concentration of 1 × 10⁷ cells mL⁻¹. To remove cell wall, cells were incubated with 10 µg mL⁻¹ Zymolyase (Roche) for 1 h at 37 °C in sorbitol buffer. Afterwards, 1 mL of spheroplasts were centrifuged at 500 rpm for 5 min, mixed with the Annexin-V-FITC and PI and incubated for 10 min at room temperature (RT). Cells were visualised by the fluorescence microscope (Leica DMI 6000, Leica Microsystems, Wetzlar, Germany). Two independent experiments with a minimum of 150 cells each were assessed. To prevent counting the same cells, cells were viewed in the carefully chosen microscope fields.

Immunostaining and fluorescence microscopy

Chromosome segregation was analyzed using fluorescence microscopy as previously described⁴⁸. Shortly, yeast strain JG15457 with the chromosome II associated with GFP was cultured in YES medium at 30 °C and 150 rpm overnight. Next day, cells were diluted and collected after 6 h of incubation with 20 mM AA, fixed by 2% PFA, and stained with primary TAT1 mouse monoclonal anti-tubulin and rabbit polyclonal anti-GFP antibodies, DAPI was used to visualize DNA. The fluorescence microscope (Leica DMI 6000, Leica Microsystems, Wetzlar, Germany) with a digital camera was used for the analyses. The proper or improper segregation of chromosome II was assessed in at least 200 cells that were in the anaphase stage of the mitotic cell cycle.

Cell cycle analysis of S. pombe cells exposed to AA

For cell cycle analysis, cells were stained with propidium iodide (Thermo Fisher Scientific) at an initial cell concentration of 5 × 10^6 cells/mL, incubated in 1 mM, 10 mM, and 20 mM acrylamide for 3 and 6 h. As a positive control, cells were incubated in 12 mM hydroxyurea for 4 h. After incubation, 1 ml of each Sample was washed in 50 mM phosphate buffer (1 M KH₂PO₄, 1 M K₂HPO₄, pH 7.8) and fixed in 70% ethanol at 4 °C overnight. To prevent RNA staining, cells were treated with RNase A (Thermo Fisher Scientific) in a concentration of 100 µg mL⁻¹ in 20 mM EDTA (pH 8) for 3 h at 37 °C. Subsequently, cells were stained with propidium iodide (4 µg ml⁻¹) in 20 mM EDTA. Propidium iodide fluorescence was detected using a CytoFLEX S cytometer (Beckman Coulter, Indianapolis, Indiana, United States) with the PE channel (585/42). Data were analysed using CytExpert 2.4 software (Backman Coulter, Inc., 2011–2019).

Gene expression

Total RNA was extracted from cells exposed to indicated AA concentrations for 6 h with the use of RiboPure™-Yeast Kit (ThermoFisher Scientific, Waltham, MA, USA), following the manufacturer’s protocol. An additional DNase treatment step was performed to eliminate genomic DNA contamination. RNA quality and quantity were assessed using NanoPhotometer™ (Implen GmbH, Munich, Germany). The quality of RNA was evaluated by measuring the absorbance ratios at 260 nm/280 nm and 260 nm/230 nm. First-strand cDNA synthesis was performed using RT² First Strand Kit (QUIAGEN, Germantown, MD, USA), with 2320 ng of total RNA as a template. Gene expression levels were quantified using qPCR with the gene-specific primers (Supplementary Table 1) on Real-Time PCR Thermal Cycler qTOWER3 (Analytik Jena GmbH + Co. KG, Konrad-Zuse-Straße 1, 07745 Jena, Germany). The expression levels of target genes were normalized using the housekeeping gene act1 (actin, ID SPBC32H8.12c) serving as internal controls with the use of relative quantification by Pfaffl⁴⁹. EliZyme™ Green MIX AddROX (Elisabeth Pharmacon, Brno-Židenice, Czech Republic) was used according to the manufacturer’s instructions for the qPCR reactions.

PCR conditions:

95 °C for 2 min (preincubation), followed by: 40 cycles using the following conditions: denaturation at 95 °C for 5 s, annealing at 59 °C for 20 s for the target genes sod1, sod2, ctt1, pgr1, and 62 °C for 20 s for the target genes grx1, gst1, gpx1, cdc2, cdc13, cdc25, cdc15, ark1, tor1, ste20, sin1, wat1, sty1, gad8, ksg1, bit61. Melting curve analysis was performed after the completion of amplification cycles to confirm the specificity of the PCR products.

Each experimental reaction was performed in biological triplicate, and technical duplicate.

Phosphorylation detection

For detecting phosphoproteins, harvested cells were disrupted with glass beads in 20% trichloroacetic acid (TCA) (Merck Life Science, Bratislava, Slovakia) solution. Precipitated proteins were resolved with 4× SDS sample buffer (Bio-Rad Laboratories, CA, USA) and then boiled for 5 min. The samples were separated by SDS-PAGE and Western blots prepared with the use of trans-blot turbo transfer system (Bio-Rad Laboratories, CA, USA) were probed for the presence of phosphorylated Rps6 was detected by Phospho-(Ser/Thr) Akt Substrate (PAS) Antibody (Cell Signaling Technology, MA, USA) at 1:2000. Detection of the proteins was carried out with standard chemiluminescent method using ECL (Merck Life Science, Bratislava, Slovakia) and the detection system (ChemiDoc, Bio-Rad Laboratories, CA, USA).

Statistical analysis

All presented findings are the result of two (biochemical and morphology parameters) to four independent experiments (growth parameter), with each experiment containing at least three (gene expression) or more biological replicates (all other methods) in two technical replicates. Statistical analysis was performed using R-Studio software (RStudio Team, PBC, Boston, MA, USA; http://www.rstudio.com/). Data are expressed as the mean ± standard deviation (SD). Statistical significance of obtained differences was analyzed by the ANOVA Duncan´s post-hoc test. Lavene´s and Cochran tests were used to evaluate data homogeneity and normality distribution of the results. Limits of the statistical significance were set up to p < 0.05 *, 0.01 **, 0.001 ***. Correlation relationships between individual traits were assessed by Pearson’s correlation coefficient (r_P) with indicated level of significance (p).

Results and discussion

Acrylamide alters cell growth and morphology

To investigate AA toxicity, we have determined the influence of increasing AA concentrations to cell growth. As expected, cell exposure to AA decreases cell growth in a dose-dependent manner. Increasing AA concentration caused stepwise inhibition of the cell growth until its almost complete interruption at 60 mM, constituting more than 2.75-fold growth inhibition compared to the control, p < 0.001, (Fig. 1A). This is in agreement with other studies, showing dose-dependent AA-mediated retardation of cell growth in various model systems including lung epithelial cell line (BEAS-2B), yeast Saccharomyces cerevisiae, and human umbilical vein endothelial cells (HUVECs)^6,50,51. The half-inhibition concentration of AA, the IC₅₀ value was determined as 26.54 mM, based on a concentration range of 1 to 1000 mM (Supplementary Fig. 1). Delayed growth of cells is associated with the AA-mediated concentration-dependent increase of the cell doubling time (Fig. 1B). Cell exposure to lower AA concentrations (0.1 and 1 mM) had none or only mild, but significant (p < 0.001), effects on cell doubling time while higher concentrations (10 and 20 mM) dramatically increased the doubling time of cells. The time required for one division cycle of the control cells (0 mM AA) was 2.64 h, while the time required for one cell division in the presence of 20 mM AA doubled compared to the unaffected control. Concentrations as high as 60 mM led to nearly complete cell growth arrest or severe cell growth retardation, causing the cell doubling time to increase more than 4 times, to 11.43 h, compared to the control. In line with our results, the effect of cell doubling prolongation after AA exposure has been observed in human lung adenocarcinoma cell line (A549) but also in HUVECs in which long-term exposure to low AA concentrations significantly prolonged cumulative population doubling^50,52. According to⁵³, delayed cell proliferation may be associated with the AA-mediated impairment of kinesin motors functionality that affects mitotic/meiotic cell cycle progression. Furthermore, as S. pombe cells elongate at the cell tip during the cell cycle to reach the optimal size for division, it is important to analyze changes in cell morphology as an indicator of an incorrect cell division process. The shape of cells exposed to AA changed slightly and depended on the concentration, with significant alterations noted at higher AA levels (Fig. 1C-D, and Supplementary Fig. 2). A statistically significant increase in the cell length and width in the presence of 20 mM AA led to a 15% increase in the cell surface and a 20% increase in cell volume (p < 0.001). These changes suggest that the cell cycle regulation is affected since cell size in S. pombe is crucial for normal cell cycle progression⁵⁴. Concomitant with our results, shape changes derived by AA were observed also in human colon adenocarcinoma cell line Caco-2⁵⁵. Given that cells exposed to AA exhibit stunted growth and cease dividing, we investigated how AA exposure affects cell fate decisions. Notably, despite fairly affected cell growth, AA exposure led to higher levels of apoptosis or necrosis only in high concentrations (40 mM), indicating that cells are more likely to stop dividing rather than die (Fig. 1E). In several human cell lines, such as the human astrocytoma U-1240 MG, human neuroblastoma cells (SH-SY5Y), and the human colon adenocarcinoma cell line Caco-2, exposure to AA-induced apoptosis, albeit to a much more pronounced extent^{55, 56–57}. Collectively, these data indicate that AA exposure affects the regulation of cell division, rather than causing cell death. We believe that the effect of AA exposure results from accumulated changes in the cellular regulatory processes, which we investigated in subsequent experiments on S. pombe cells.

Fig. 1 [Images not available. See PDF.]

AA-mediated alterations of cell division, cell morphology, and cell fate decisions. (A) Growth intensity was evaluated as the OD₆₀₀ ratio determined every three hours of incubation (3 h, 6 h, and 9 h) and calculated as the difference to the time point 0 h. Increasing AA concentration reduces cell division in a dose-dependent manner. The data are represented by a symbol that indicates the mean and standard deviation (SD) of three independent experiments with four biological replicates. (B) Generation time (gt) represents cell division time reflecting the time the cell population requires for its doubling. Increasing AA concentrations significantly increased cell doubling time in a dose-dependent manner. Each bar with a tick represents the mean and SD of three independent experiments with four biological replicates. Cell morphology: surface (C), and volume (D) were determined in S. pombe cells exposed to indicated AA concentrations. Each bar represents mean value ± SD (n=100). Statistical significance in A, B, C, and D is determined by Duncan’s post-hoc test, indicated as p < 0.05 *, 0.001 ***, nd indicates no statistically significant difference. (E) Cell death determination. To evaluate cell death by apoptosis, the Annexin V-FITC signal was determined with fluorescence microscopy upon AA exposure, cells were co-stained with PI to distinguish between apoptosis and necrosis. PI-positive cells were considered necrotic. Cell fractions of at least 150 counted cells represent a portion of cells undergoing apoptosis, necrosis, or staying alive.

Acrylamide-induced oxidative stress affects cell vitality

Affected cell growth is often associated with altered cellular respiration. We show here that AA exposure impaired the respiration activity of cells. Lower AA concentrations (1 and 10 mM) led to its slight increase (p < 0.05 or non-significant, respectively), while high AA concentration (20 mM) dramatically reduced (more than 2-fold, p < 0.001) the respiration activity of the cell (Fig. 2A) impairing cell vitality. As cellular respiration occurs in mitochondria, altered respiration activity suggests mitochondrial dysfunction or loss of mitochondrial quality. Together, the mitochondrial electron transport chain (ETC) and ATP-synthase provide most cellular energy. The ETC utilizes over 95% of the oxygen consumed during respiration; however, some electrons are directly transferred to O₂, resulting in the formation of reactive oxygen species (ROS). It is well-documented that stress disrupts the balance between the production and degradation of ROS. This disruption leads to excessive ROS production by the ETC, which can ultimately result in cell damage or death^10,58. In our experimental setup, AA exposure boosted ROS generation in a dose-dependent manner (p < 0.001) (Fig. 2B), as also demonstrated by others in the human primary glioblastoma cell line, U-87 MG, yeast Saccharomyces cerevisiae, or rats^{59, 60–61}. The considerable rise in MDA production (Fig. 2C), an end product of lipid peroxidation that indicates cell membrane damage, provides evidence that cells are under oxidative stress.

When an electron from the ETC is transferred to oxygen, a superoxide anion (O₂^•−) may form through enzymatic reactions involving the ETC complex⁶² or through non-enzymatic reactions such as the Fenton and/or Haber-Weis reactions⁶³. In radical chemistry, O₂^•− is readily converted into other ROS. A quantitative assessment of individual ROS was performed to more accurately define stress mediated by acrylamide. Analytically, it was determined that the formation of O₂^•−, H₂O₂, and hydroxyl radical (OH^•) increased along with increasing AA concentration (Fig. 2D-F). Despite the slight increase in respiratory activity of cells exposed to low concentrations of AA (1 mM), no statistically significant increase in intracellular concentrations of O₂^•−, H₂O_2, and OH^• was observed. On the other hand, higher concentrations of AA (10 and 20 mM, respectively) induced a dramatic increase in the concentration of all studied radicals. In particular, the intracellular concentration of O₂^•− increased by over 5-fold after exposure to 20 mM AA (Fig. 2D). The reaction of O₂^•− with H₂O₂ (Haber-Weiss reaction) and/or H₂O₂ with divalent iron cations (Fenton reaction) generates the highly reactive OH^• radical, which is likely responsible for most of the oxidative damage attributed to ROS⁶⁴. As the existence and localization of membrane-bound extramitochondrial NADPH oxidase complexes (NOX) have been identified in single-celled living fungi, including S. cerevisiae^65,66, it is reasonable to speculate that intracellular ROS generation in S. pombe cells may not only be produced by ETC but may also result from the activation of NOX complexes⁶⁷ induced by AA. However, this hypothesis needs to be clarified in future studies.

Given that ROS play a dual role in cells, as they can cause oxidative damage¹⁰ while also acting as molecular signals that activate stress responses^68,69, a strict balance between their production and detoxification needs to be maintained. Thus, we have analysed the antioxidant defence mechanisms of S. pombe cells exposed to AA.

Fig. 2 [Images not available. See PDF.]

Determination of AA-induced oxidative stress. (A) Respiration activity (RA) determined as formazan production, measured at 485 nm and normalized to protein unit, indicates cell vitality and functionality of mitochondria. Lower AA concentration had only a mild effect on respiration activity, while high concentration dramatically decreased RA, indicating mitochondrial malfunction. (B) Determination of the total ROS was performed with the use of H₂DCFDA as a ROS formation indicator. The generation of ROS was compared between the control group of cells and cells exposed to AA for 6 h, indicating that increasing AA concentration led to an increase in ROS levels. (C) MDA is an end-product of lipid peroxidation that serves as an indicator of oxidative damage. As indicated by enhanced MDA production, cell exposure to AA causes peroxidation of membrane lipids in a dose-dependent manner. Determination of individual ROS formation revealed that levels of superoxide anion (O₂^•−) (D), hydrogen peroxide (H₂O₂) (E), and hydroxyl radical (OH^•) (F) increase along with increasing AA concentration indicating AA-mediated oxidative stress. In all experiments, individual bars represent the mean value of 8 samples from two independent experiments ± SD. Statistical significance, determined by Duncan’s post-hoc test, was set up as p < 0.05 *, 0.01 **, 0.001 ***, nd indicates no statistically significant differences.

Cell protection against AA-derived oxidative stress

Antioxidant defence machinery in cells serves to protect organism against oxidative damage. The primary defence system consists of antioxidant enzymes system and antioxidant molecules. Their role is to alleviate ROS or to reduce the reactivity of individual ROS. Superoxide dismutase (SOD) reduces superoxide radicals to hydrogen peroxide, and catalase (CAT) degrades H₂O₂ to O₂ and H₂O. The activity of the two enzymes dramatically increased in a dose-dependent manner upon AA exposure (Fig. 3A, B), which is mostly in contrast to the results of other authors, indicating a decrease in the SOD and CAT activities^51,70,71. These discrepancies may arise from the use of different model systems, including animal models such as rats or yeast Saccharomyces cerevisiae, and variations in the experimental setup. We assume that the acceleration of ROS formation acts as a signal and stimulates cells to increase the gene transcription and/or activity of the antioxidant enzymes SOD and CAT for self-protection against AA toxicity. As we have shown, cell exposure to AA increased ROS formation, primarily superoxide anion (O₂^•−) (Fig. 2D). This, in turn, leads to an increase in the expression of the genes sod1 and sod2, along with a significant rise (nearly 12-fold, p < 0.001) in SOD activity. Consequently, enhanced SOD activity results in increased H₂O₂ production, which subsequently causes a substantial increase in the expression of the gene ctt1 and the activity of CAT (5.5-fold, p < 0.001) (Fig. 3A, B, and Fig. 4A-C). In contrast to the increase in SOD and CAT activity, AA at higher concentrations of 10 and 20 mM significantly reduced the activity of glutathione peroxidase (GPx). However, a lower AA concentration of 1 mM led to a slight increase in GPx activity (p < 0.01) (Fig. 3C). GPx, similarly to CAT, detoxifies hydrogen peroxide and also organic peroxides utilizing glutathione (GSH) as an electron donor⁷². Despite the decrease in GPx enzyme activity in response to higher AA concentrations, the expression of the gpx1 gene increased upon exposure to AA (Fig. 4D). Low intracellular H₂O₂ concentration in the signalling pathway in the cell’s stress response to 1mM AA activates redox-sensitive proteins such as peroxiredoxins and antioxidant enzymes, including GPx. The decrease in GPx activity at high AA concentrations may occur either directly due to increased ROS formation and/or by modification of the enzyme structure⁷³. For optimal GPx activity, a balanced redox state is essential, unlike in the presence of AA, as GPx activity is directly dependent on the concentration of reduced glutathione (GSH). Thus, we believe that the imbalance of the redox state and GSH and its oxidized form (GSSH) levels results in decreased GPx activity.

Fig. 3 [Images not available. See PDF.]

AA affects antioxidant cell capacity through modulation of antioxidant enzyme activity. The antioxidant capacity of cells was assessed via the determination of antioxidant enzyme activity upon AA treatment. CAT (A) and SOD (B) activity increased in cells exposed to AA in a dose-dependent manner. GPx (C) activity slightly increased after cell exposure to low AA concentration (1 mM), whereas cell exposure to higher AA concentrations resulted in decreased GPx activity. Bars in the graph represent the mean ± SD of 4 individual samples. Statistical significance is determined by Duncan’s post-hoc test and was set up as p < 0.05 *, 0.01 **, 0.001 ***.

In S. pombe glutathione S-transferases are induced by H₂O₂ and are regulated by the stress-activated protein kinase Sty1. Three genes, gst1, gst2, and gst3 encode glutathione S-transferases in S. pombe, of which gst1 and gst2 encode closely related proteins with related functions⁷⁴. Surprisingly, we found that AA downregulates gst1 expression (Fig. 4E), suggesting that the cell prefers to utilize other defence mechanisms to cope with AA-induced stress or that AA modulates transcription factors responsible for the regulation of gst1 expression.

Among reducing thiols, the tripeptide glutathione (GSH), consisting of l-γ-glutamyl-l-cysteinylglycin, is one of the most common antioxidant molecules due to its ability to detoxify ROS and to serve as an electron donor for antioxidant enzymes. ROS, metals, and disulfides all react with GSH to oxidize it, producing oxidized glutathione (GSSG). Glutathione reductase (GR) effectively mediates the reduction of GSSG to GSH by employing NADPH as a reducing equivalent^18,75. In S. pombe, the GR, encoded by the pgr1 gene, is essentially required for cell growth under aerobic conditions and is essential for GSSG reduction to GSH under stress conditions⁷⁶. In our experimental setup, cells exposed to AA displayed elevated pgr1 expression (Fig. 4F). We propose that acrylamide induces the oxidation of glutathione, prompting the cell to elevate the expression of the pgr1 gene in order to convert the oxidized glutathione back to its reduced form. Notably, given that GPx activity directly depends on GSH levels, we assume that upregulated pgr1 expression suggests elevated GSSG concentration, which is responsible for the decrease in the GPx activity (Fig. 3C) despite enhanced expression of the gpx1 gene (Fig. 4D) under high AA concentrations. Additionally, as GSH serves as an electron donor for glutaredoxins to reduce protein disulfides or glutathione–protein mixed disulfides, we have determined the expression of the major dithiol glutaredoxin in S. pombe, grx1. The two glutaredoxins in S. pombe, Grx1 and Grx2, function in distinct cellular compartments: Grx1 operates in the cytosol, while Grx2 is located in mitochondria⁷⁷. However, the changes in the mRNA expression of only grx1 have been determined under oxidative stress⁷⁸. Concomitant with this, AA exposure resulted in the increase of grx1 expression (Fig. 4G), suggesting a stress-induced expression of the gene to contribute to cell protection against AA toxicity.

Fig. 4 [Images not available. See PDF.]

AA-mediated changes in the expression of genes encoding antioxidant enzymes. Total RNA was extracted from control cells and cells exposed to an indicated amount of AA for 6 hours. RT-qPCR was used to determine gene expression changes upon AA exposure. Quantification analyses revealed that AA exposure resulted in upregulation of sod1 (A), sod2 (B), ctt1 (C), gpx1 (D), downregulation of gst1 (E), upregulation of pgr1 (F), and grx1 (G) in a dose-dependent manner. To determine changes in expression, act1 was used as the housekeeping control, and fold change was evaluated by relative quantification according to Pfaffl (Pfaffl, 2001). The values in the bars are the means of 3 biological and two technical replicates ± standard deviations. p values were determined using Duncan’s post-hoc test, and statistical significance was set up as p < 0.05 *, 0.01 **, 0.001 ***, nd stands for no significant differences.

Correlation analysis confirmed highly significant positive correlations between increased expression of genes involved in the antioxidant cascade (except for gst1) and production of total and individual ROS (Supplementary Fig. 3). Cells thus utilize complex systems to manage AA toxicity, including GR-mediated reduction of oxidized glutathione (GSSG) to its reduced form (GSH) that can subsequently serve as an electron donor for the glutaredoxins. Nonetheless, the defence mechanisms are insufficient to protect cells against AA at concentrations that exceed the antioxidant capacity of the cell, leading to cell growth retardation or cell cycle arrest. To investigate the impact of AA exposure on the redox balance within cells, we analyzed the levels of glutathione (GSH) under conditions of AA-induced stress. AA exposure significantly reduced GSH levels as early as 3 h after incubation, and the decrease was persistent and maintained after 6 h of incubation (Fig. 5). These results support our earlier findings that exposure to AA disrupts the cellular antioxidant capacity, ultimately leading to redox imbalance and oxidative stress. Consistently, correlation analysis revealed that reduced GSH levels are negatively associated with oxidative stress markers such as SOD and CAT activity (r_P < −0.6), as well as the regulation of antioxidant genes. In contrast, GSH levels showed a positive correlation with GPx activity (r_P = 0.478), indicating that GPx function is largely dependent on GSH availability (Supplementary Fig. 3).

Fig. 5 [Images not available. See PDF.]

AA-induced oxidative stress is related to AA-mediated decrease of GSH levels. Control cells and those exposed to a specified amount of acrylamide (AA) for 3 and 6 hours were used to determine the concentration of reduced glutathione (GSH). The results showed that acrylamide caused a significant decrease in GSH levels in a dose-dependent manner. This reduction in GSH was noticeable 3 hours after exposure to AA (A) and continued to persist after 6 hours (B) of AA incubation. The values in the bars are the means of 8 biological replicates ± standard deviations. p values were determined using Duncan’s post-hoc test, and statistical significance was set up as p < 0.001 ***.

Acrylamide negatively influences the regulation of the cell division cycle

To investigate the effect of AA-induced oxidative stress on cell cycle regulation, we analysed changes in the expression of genes involved in regulating the cell cycle and examined chromosome segregation and cytokinesis.

As in higher eukaryotes, the cell cycle of S. pombe is divided into four phases, G1-phase, S-phase, G2-phase, and M-phase, with the major regulator a 34 kDa serine/threonine kinase Cdc2. The expression of the cdc2 gene remains constant throughout the cell cycle, while its activity changes depending on the cell cycle phase. Cdc2 activity drives progression through the cell cycle at both the G1/S and G2/M boundaries, depending on its association with a particular cyclin^79,80. Cdc2, Cdc13, and Cdc25 are positive cell cycle progression regulators at the G2/M interface⁸¹. Exposure of S. pombe cells to AA caused alterations in the expression of genes encoding those principal cell cycle regulators cdc2, cdc13, and cdc25. Although the expression of the cdc2 gene does not change during the cell cycle, the genotoxic effect of AA reduces its expression in a dose-dependent fashion (p < 0.001) (Fig. 6A). Similarly, the expression of the gene encoding the B-type cyclin Cdc13, which is required for Cdc2 activation, has been affected by AA. While lower concentration (1 mM) led to increased expression of cdc13 (more than 25%, p < 0.001), higher concentrations (10 and 20 mM) caused a decrease in the expression of the cdc13 gene (Fig. 6B). The protein phosphatase Cdc25 dephosphorylates Cdc2 on Y15 to allow entry into mitosis by activating the Cdc2/Cdc13 complex⁸². The expression of the cdc25 gene is reduced in cells exposed to AA (p < 0.001), similarly to the cdc2 gene (Fig. 6C). Changes in the expression of genes required for cell cycle regulation have been reported in various studies dealing with the response of the organism, including mice or legume plant Vicia faba, to oxidative stress^{83, 84–85}. Our results show that AA-induced oxidative stress downregulates the expression of genes involved in mitotic control, resulting in cell cycle arrest. Under physiological conditions, cell size control is closely linked with cdc25 expression as cells that grow in size express more cdc25, while cells overexpressing cdc25 enter mitosis in reduced size^86,87. Accordingly, we demonstrate that although AA exposure slightly increases the size of the cell (Fig. 1D), the expression of cdc25 decreases, and cell division is reduced (Fig. 1A, B). Under normal conditions, the expression of cdc25 increases in line with the cell cycle progression, and the activity of Cdc25 is essential for mitotic entry⁸⁸. Due to the AA-triggered downregulation of cdc25 expression, we can assume that Cdc2 becomes hyperphosphorylated, leading to the inhibition of the G2/M transition, ultimately preventing the progression of cell division during mitosis. To further investigate the impact of AA on the progression of the cell division cycle, we have looked at the expression of the gene that encodes the protein required for the mitotic exit and cytokinesis, the F-BAR protein Cdc15. During mitosis in eukaryotic cells, the cytoskeleton undergoes significant changes. Microtubules reorganize to separate the duplicated sister chromosomes, while filamentous (F)-actin reorganizes to facilitate cell division⁸⁹. Cytokinesis in S. pombe requires an actomyosin-based division apparatus linked to the plasma membrane, the contractile ring (CR), necessary for symmetrical cellular division⁹⁰. Cdc15 has been shown to play an essential role in regulating cytokinesis onset in S. pombe. The mRNA levels of this phosphoprotein change during the cell cycle, culminating at the septum formation initiation. S. pombe cells depleted from cdc15 are unable to form a functional medial ring and a division septum⁹¹. We show here that cell exposure to AA results in altered cell septation. Increasing AA concentration caused downregulation of cdc15 mRNA levels (20% downregulation, p < 0.001), resulting in a decrease in septum formation (Fig. 6D, E). Although low AA concentration (1 mM) caused elevation of the cdc15 expression, cells were not able to form a functional septum as depicted in Figs. 6D and E. The ability to generate a septum varied greatly across exposed cells. Higher AA concentrations led to a significant decrease in cdc15 expression, which reduced septum development while the size of cells increased (Fig. 1C, D, and Supplementary Fig. 2 A, B). Thus, we show here that AA exposure not only affects cell cycle regulation but also reduces the ability of the cell to undergo cytokinesis. In addition, the commitment to the cell cycle progression is presented by a series of feedback-controlled network processes that result in splitting the mother cell into two identical daughter cells, each possessing the same genetic information. Chromosome segregation during mitosis must be precisely coordinated in space and time with the process of cytokinesis. Improper chromosome separation before cytokinesis can cause aneuploidy, linked to cancer development⁹². Aurora kinases have been implicated in the regulation of distinct mitotic events. Aurora kinases share high similarity in their kinase domains and are classified into three types: A, B, and C. Aurora-A kinases are critical for centrosomal duplication, maturation, and separation, necessary for the formation and stabilization of the bipolar mitotic spindle⁹³. Aurora-B subfamily members function later in mitosis and belong to a class of proteins referred to as chromosomal passengers, essential for coordinating chromosome segregation with cytokinesis⁹⁴. Less studied is the Aurora-C kinase, which resembles Aurora-B in both location and function in mammals⁹⁵. The single Aurora kinase in fission yeast, Ark1, integrates the functions of Aurora-A and Aurora-B kinases found in higher organisms. It has been demonstrated that ark1 is an essential gene necessary for the spindle attachment response and for proper association with kinetochores, which is crucial for the correct segregation of sister chromatids during mitosis. Cells lacking the Ark1p exhibit fragmented nuclei and frequently display defects in chromosome condensation and migration^96,97. In consistent with this, we show here that exposure of cells to high concentrations of AA (10 and 20 mM) leads to reduced expression of the ark1 gene (almost 50%, p < 0.001) (Fig. 6F), which in turn causes chromosome missegregation during mitosis (Fig. 6G). Cell cycle progression in response to AA exposure, as determined by flow cytometry, showed an accumulation of 4 C DNA content in cells exposed to AA for three hours. Although cells exposed to AA for six hours did not exhibit any significant changes in DNA content (Fig. 6G, H), we believe that the 4 C accumulation observed at three hours caused a delay in the cell cycle. This suggests an S-phase arrest and the G2/M boundary checkpoint activation, which confirms the previously described data. We assume that high concentrations of AA exposure lead to DNA-damaging activities, which activate critical cell cycle progression checkpoints and hinder cell cycle progression, similar to what has been previously described by⁹⁸. The correlation analysis showed that changes in the cell cycle regulatory genes expression positively correlated (r_P > 0.5) with the growth activity, while a close negative correlation (r_P < −0.5) between these genes and morphological traits, individual ROS, and antioxidant enzyme activity, except for GPx, has been detected (Supplementary Fig. 3).

Our findings indicate that AA modulates cell division control at multiple stages, leading to altered cell growth and potential errors in chromosome segregation, which could contribute to carcinogenesis. The detrimental impact of AA on cell cycle regulation is illustrated in the schematic drawing depicted in Fig. 6J.

Fig. 6 [Images not available. See PDF.]

AA exposure disturbs cell cycle regulation. Control cells and cells exposed to an indicated amount of AA for 6 hours were used for the total RNA isolation and gene expression was determined using RT-qPCR. Quantification analyses revealed that AA exposure resulted in moderate upregulation of cdc2 (A), low AA concentration induced slight cdc13 upregulation, while higher amounts caused downregulation of cdc13 (B), AA exposure caused downregulation of cdc25 in a dose-dependent manner (C). AA exposure negatively influences cell separation by modulation of cdc15 expression (D) resulting in the accumulation of cells with defective septation (E). Chromosome segregation is affected upon AA exposure as indicated by the dose-dependent downregulation of ark1 (F) resulting in the enhancement of cells displaying chromosome missegregation (G). The values in the bars (in A, B, C, D, F) are the means of 3 biological and two technical replicates ± standard deviations. (E). (G) Comparison of the cell cycle in Schizosaccharomyces pombe cells exposed to 0 mM, 1mM, 10 mM, and 20 mM acrylamide for 3 and 6 hours. The X-axis represents DNA content in the cells, measured as the fluorescence intensity of propidium iodide. In the untreated sample, most cells showed a 2C DNA content after incubation for both 3 hours and 6 hours. Exposure to AA caused an accumulation of cells in the S phase at the 3-hour incubation, leading to a delay in cell cycle progression. (H) The graphs represent the percentage of DNA content in both control cells and cells treated with AA after 3 and 5 h of incubation. In (I) bars represent the percentage of cells with the defect in the segregation of chromosome II, n=100. p values were determined using Duncan’s post-hoc test, and statistical significance was set up as p < 0.05 *, 0.01 **, 0.001 ***, nd stands for no significant differences. (J) The schematic drawing represents the impact of AA exposure on individual steps of the cell cycle regulation, cell separation, and chromosome segregation, all leading to cell growth retardation.

Involvement of the TOR signaling in cell response to AA

The TOR (target of rapamycin) pathway is a crucial nutrient and stress sensor conserved from yeasts to humans. In mammals, its deregulation is often associated with an increased risk of cancer, diabetes, and aging processes^99,100. In S. pombe, loss of Tor1 kinase, the catalytic subunit of TORC2, results in defective cell growth under various stresses. Although Tor1 is not essential for vegetative growth under nutrient-rich conditions, it is required under starvation and other stress conditions, including oxidative stress¹⁰¹. Loss of function in other components of the TORC2 complex, such as Ste20, Sin1, or Bit61, leads to similar phenotypes as the Tor1 mutation, highlighting the crucial role of TORC2 in the cellular stress response¹⁰². In our study, we have investigated the response of AA-induced stress on the expression of genes encoding individual members of the TORC2 complex under nutrient-rich conditions. Cells exposed to AA displayed a significant decrease in the expression of tor1, ste20, bit61, and wat1 genes, while the decrease of the sin1 gene expression did not reach statistical significance (Fig. 7A-E). A nearly 50% reduction in expression (p < 0.001) has been observed in the wat1 gene (Fig. 7D), which is a critical component of both TOR complexes^103,104. Functionally, Wat1 a mammalian Lst8 homolog, has been connected to cell shape maintenance, mRNA maturation, and more recently, to oxidative stress response¹⁰⁵. Wat1 interacts with the C-terminal region of Tor1, this binding, stabilized by the FATC domain of Tor1, and together with the phosphorylation of Wat1 is required for the regulation of Gad8 function. In addition, the AGC kinase Gad8 requires phosphorylation of its activation loop (T-loop) by Ksg1, a PDK1 homolog in S. pombe, for full functionality¹⁰⁶. Strikingly, genes encoding Gad8 kinase and Ksg1 kinase have both been downregulated in cells exposed to AA (Fig. 7F, G). This slight but substantial AA-mediated downregulation of the TORC2 complex genes, along with the regulatory kinases, disrupts the cell stress response, leading to cell growth retardation. Given that the major catalytic subunit of TORC2, Tor1 kinase, is activated during starvation, our findings imply that the alterations in gene expression and the ensuing cell responses directly relate to AA-induced stress rather than to nutrient-derived dysregulation of TORC2 function. On the other hand, AA-induced downregulation of the TORC2 genes compromises the function of the complex and thus contributes to the disturbed cell conditions. Disruption of the TORC2 function has been associated with delayed entrance into mitosis and the formation of slightly elongated cells¹⁰⁷, which is in accordance with our results (Supplementary Fig. 2 A). Moreover, the fact that the TORC2-Gad8 module is required for cell survival under DNA-damaging conditions⁴¹ explains to a large extent the response of cells to AA-induced stress, as AA has been shown to possess DNA-damaging properties in mammals¹⁰⁸. Besides, as the coordination of the stress-activated MAPK pathway contributes to TORC2 activation³⁵, we have examined the expression of sty1, the major stress regulatory kinase of the MAPKs. Surprisingly, sty1 expression remained unaffected upon AA-induced stress (Fig. 7H). We assume that sty1 expression remained stable because the AA-mediated formation of individual ROS led to more than 5-fold increase in the O₂^•− levels, while the increase of H₂O₂ was only twice as high as in the control group of cells, which was likely not sufficient for the need to increase the sty1 gene expression.

As reported by¹⁰⁹, high levels of H₂O₂ are required for the induction of sty1 expression and Sty1 activation. According to our results, we believe that AA-induced downregulation of wat1 and tor1, combined with ksg1, results in decreased Gad8 phosphorylation, which limits the function of Gad8 and thus reduces Pyp1 and Pyp2 activity. This, in turn, results in a decreased dephosphorylation of hyperactivated MAPK Sty1/Spc1¹¹⁰, leading to an altered response to the stress of affected cells.

Correlation analysis confirmed that expression of the TORC2 genes closely positively correlates (r_P > 0.5) with the growth activity. Additionally, it is noteworthy that there is a high positive correlation between GSH and the gene expression of cell signaling components, particularly tor1 (rP = 0.834) and ste20 (rP = 0.747) (Supplementary Fig. 3). Our results indicate that the response of cells to AA-induced oxidative stress is ultimately related to the alterations in the expression of genes of the TORC2 signalling pathway.

Fig. 7 [Images not available. See PDF.]

Cell response to AA-mediated stress is mediated through TORC2 rather than MAPK. After RNA extraction from control cells and cells exposed to an indicated amount of AA for 6 hours, RT-qPCR was used to determine gene expression changes. AA-induced downregulation of tor1 (A), ste20 (B), bit61 (C), and wat1 (D), while expression of sin1 remained unaffected (E). Moreover, gad8 encoding the direct substrate of Tor1 was downregulated upon AA exposure (F). Similarly, ksg1 which encodes the essential kinase Ksg1 required for full Gad8 activation, was slightly upregulated upon low AA concentration, however high AA concentrations downregulated its expression substantially (G). Surprisingly, the expression of sty1, which encodes the major stress-activated MAPK kinase in S. pombe, remained unaffected upon AA exposure (H). To determine changes in expression, act1 was used as the housekeeping control, and fold change was evaluated by relative quantification according to Pfaffl (Pfaffl, 2001). The values in the bars are the means of 3 biological and two technical replicates ± standard deviations. p values were determined using Duncan’s post-hoc test, and statistical significance was set up as p < 0.05 *, 0.01 **, 0.001 ***, nd stands for no significant differences.

To further investigate and validate the involvement of TORC2 signaling, we performed a phosphoprotein assay to assess the phosphorylation status of the ribosomal Rps6 protein. Although Rps6 was initially identified as a TORC1-specific substrate¹¹¹, Du et al.¹¹² demonstrated that its phosphorylation is regulated not only by TORC1 but also by TORC2 via Gad8 at serine 546 (Gad8.S546). Previous studies have shown that the anti-phospho-Akt substrate (PAS) antibody, which recognizes the phosphorylated [R/K]X[R/K]XX[S/T] consensus motif found in mammalian S6K1 and other AGC kinase substrates, can detect phosphorylated Rps6 in fission yeast. In S. pombe, Rps6 is encoded by two genes, rps601 and rps602, with the latter—co-purifying with Gad8—contributing predominantly to the observed Rps6 phosphorylation signal. Accordingly, we employed the anti-PAS antibody in AA-treated cells to confirm the role of TORC2, via Gad8-mediated phosphorylation of Rps6, in the cell response to AA-induced toxicity. Notably, Rps6 phosphorylation was almost completely abolished after 3 h of exposure to high concentrations of AA (10 and 20 mM), whereas low-dose treatment (1 mM) led to enhanced phosphorylation. Interestingly, by 6 h post-treatment, Rps6 phosphorylation levels increased compared to the untreated control (Fig. 8), suggesting the activation of alternative signaling pathways that may compensate for TORC2 activity. These findings clearly indicate that the cellular response to AA-induced stress is multifaceted, involving TORC2 but likely also other pathways such as TORC1. Further investigation is thus needed to fully elucidate the molecular mechanisms underlying the cellular response to AA-induced stress.

Fig. 8 [Images not available. See PDF.]

AA exposure modulates Gad8 mediated phosphorylation of Rps6 protein. TCA isolated protein extracts from control cells and cells exposed to an indicated amount of AA for 3 and 6 hours were subjected to SDS-PAGE, the proteins were transferred to a PVDF membrane and incubated with an anti-PAS antibody that recognizes the phosphorylated Rps6 protein by TORC2 through Gad8 kinase. The anti-beta-tubulin antibody TAT1 was used as a loading control. Our results indicate that exposure to AA affects Gad8, a direct substrate of Tor1-mediated phosphorylation. This suggests that TORC2 signaling is involved in the cell response to AA-induced stress.

Conclusion

We demonstrate that AA-induced oxidative stress leads to reduced metabolic activity, delayed cell growth, altered cell cycle regulation, and chromosome missegregation. Abnormalities in chromosome segregation are linked to the AA-induced downregulation of the genes connected to cell cycle regulation, septation, and the spindle checkpoint in S. pombe. Furthermore, cell response to stress caused by AA exposure is, to a certain extent, mediated through the TORC2 signalling pathway, indicated by AA-derived downregulation of genes encoding individual members of TORC2 along with the Gad8 and Ksg1 kinases. Additionally, the modulation of Rps6 phosphorylation further supports the involvement of TORC signaling. To our surprise, the expression of sty1, the primary MAPK stress response kinase in S. pombe, remained unaffected. This indicates that TORC2, unlike MAPK, is directly involved in the cell response to AA-induced stress. Moreover, suppression of the TORC2-dependent signalling upon AA exposure might be associated with AA-induced cellular toxicity, including aberrant cell proliferation, cell cycle arrest, or DNA damage, ultimately resulting in error-prone cell cycle regulation, which is a hallmark of cancer. Taken together, the results of our experiments point to a close relationship between AA toxicity and its capability to cause oxidative stress by increasing ROS production and interfering with cell defence mechanisms, including TORC2 signalling (Supplementary Fig. 3). This, in turn, results in alterations in the cell cycle regulation, causing chromosome missegregation. Thus, we believe that these functional impairments of the fundamental cellular processes may be directly related to the possible carcinogenic effect of AA (Fig. 9).

Fig. 9 [Images not available. See PDF.]

AA-mediated stress refers to the cellular response that occurs at multiple molecular levels. AA exposure triggers oxidative stress in cells by increasing the formation of ROS and disrupting the antioxidant response. Additionally, AA alters the regulation of the cell cycle, which leads to delayed cell growth and errors in chromosome segregation. The cellular response to AA toxicity involves the TORC2 signaling pathway rather than the MAPK pathway. The disruption of the antioxidant defense system, downregulation of cell cycle regulatory genes, and alterations in the TORC2 signaling pathway all contribute to the toxic effects of AA on cells. Taken together, these factors ultimately lead to conditions that disrupt the normal progression of cell division, which is a hallmark of cancer.

Author contributions

M.K.: Data curation, Methodology, Validation, Conceptualization. A.N.: Data curation, Validation. L.K.: Data curation, Validation. A.B.: Data curation. Mária.P.: Data curation. Miroslava.P.: Writing – original draft, Supervision, Funding acquisition. All authors reviewed the manuscript.

Funding

This research was financially supported by the Slovak Research and Development Agency (APVV) under contract no. APVV-22-0294, and VEGA under grant number 1/0583/23.

Data availability

Data is provided within the manuscript or supplementary information files.

Declarations

Competing interests

The authors declare no competing interests.

Publisher’s note

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

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