Cellular responses to prolonged non-thermal plasma exposure in Schizosaccharomyces pombe
Maria Petkova, Sandra Durcanyova, Martin Kutka, Ivana Kyzekova, Katarina Gaplovska-Kysela, Katarina Soltys, Stanislav Kyzek, Veronika Medvecka, Andrea Sevcovicova
TL;DR
This study explores how fission yeast cells respond to prolonged non-thermal plasma exposure, revealing significant oxidative stress and cellular disruptions.
Contribution
The study provides new insights into the cellular and molecular effects of prolonged non-thermal plasma exposure in a eukaryotic model organism.
Findings
Extended non-thermal plasma exposure increases intracellular reactive oxygen and nitrogen species and mitochondrial superoxide.
Prolonged plasma exposure causes tubulin depolymerisation and cell cycle arrest in fission yeast.
NTP exposure alters the expression of genes involved in post-transcriptional regulation.
Abstract
Non-thermal plasma (NTP) generates a complex mixture of reactive oxygen and nitrogen species (RONS) that can impose strong oxidative stress on eukaryotic cells. While the antimicrobial potential of NTP has been widely explored, much less is known about how eukaryotic cells respond to prolonged NTP-induced stress at the cellular and molecular level. Here, we investigated the cellular effects of extended NTP exposure using the fission yeast Schizosaccharomyces pombe as a non-pathogenic eukaryotic model. Our results indicate that extended exposure to NTP significantly reduces cell viability and is associated with increased oxidative stress, as evidenced by increased levels of intracellular RONS and mitochondrial superoxide. These oxidative changes were accompanied by pronounced cellular responses including tubulin depolymerisation, cell cycle arrest, and impaired cell division. In…
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Figure 7- —Comenius University in Bratislava
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Taxonomy
TopicsPlasma Applications and Diagnostics · Surface Modification and Superhydrophobicity · Plasma and Flow Control in Aerodynamics
Introduction
Plasma is an ionised gas consisting of electrons, ions, neutral molecules, atoms, radicals, and excited species (Coutinho et al. 2018). Depending on the temperature of the particles within the plasma, it can be classified as high temperature, thermal, and non-thermal. Non-thermal plasma (NTP) is characterised by its non-equilibrium state in which electrons reach temperatures of approximately 10^6^–10^8^ K, while heavier particles remain near room temperature. This unique property enables NTP to interact with heat-sensitive materials, including biological systems, without thermal damage (Fridman et al. 2005; Sakudo et al. 2019).
NTP’s efficacy in biological and environmental applications stems from its generation of highly reactive species (including reactive oxygen and nitrogen species; RONS), alongside ultraviolet (UV) radiation and the generation of an electric field (Mishra et al. 2011). These RONS are primarily formed from the dissociation of neutral molecules and reactions of working gases like atmospheric oxygen and nitrogen (Adamovich et al. 2017; Machala et al. 2018). The environmentally friendly and sustainable nature of NTP drives its increasing adoption in environmental research and public health (Adamovich et al. 2017). Its applications range from antimicrobial treatment in the food industry (Lin et al. 2021) and agriculture (Gutiérrez-Leon et al. 2022; Molina-Hernandez et al. 2022a, b) to the degradation of hazardous pollutants in water and soil (Aggelopoulos et al. 2018; Giardina et al. 2024). A key advantage of NTP is its non-residual nature, non-thermal operation, and energy efficiency, offering a sustainable approach to microbial control and environmental management (Todorova et al. 2022).
Despite the well-documented applications and quantitative efficacy of NTP, particularly in microbial inactivation, a deeper, mechanistic understanding of its specific interactions with eukaryotic cells remains an area requiring further fundamental investigation (Nwador et al. 2022). While general antimicrobial effects of NTP are known (Kelly-Wintenberg et al. 1999; Sun et al. 2012), a comprehensive elucidation of underlying cellular and molecular mechanisms is often generalised. This study aims to provide fundamental biological insights crucial for optimising and refining NTP’s environmental and biomedical uses. Understanding how NTP impacts critical eukaryotic structures like tubulin and gene expression on a transcriptomic level can facilitate the development of more targeted and efficient decontamination strategies and inform broader environmental impact assessments by predicting and mitigating off-target effects on beneficial eukaryotic microorganisms.
The selection of Schizosaccharomyces pombe, an established model in environmental stress research, for the present study is specifically justified by the availability of specialised S. pombe strains, engineered with fluorescently labelled tubulin and endoplasmic reticulum. This critical feature allows real-time, high-resolution visualisation of specific cellular structures and processes, enabling the analysis of environmental stressor impacts at a subcellular level. This approach permits real-time visualisation of intracellular structures, revealing how NTP affects key eukaryotic processes such as cell division and cytoskeletal integrity. It offers high-resolution insight into cellular vulnerabilities to plasma, essential for assessing the broader biological impact of NTP.
Material and methods
Cell cultivation
The wild-type S. pombe strain 972 h^−^ and strain 18487 (h90 ade6 + :patb2:mCherry-atb2:terminatoratb2:hphMX lys3 + :pBiP:SignalSequenceBiP-sfGFP-AHDL:bsdMX), with fluorescently labelled tubulin and endoplasmic reticulum, were used for the experiments described in this study. The strain 18487 was prepared by mating of the strains FY38512 (h + lys3 + :pBiP:SignalSequenceBiP-sfGFP-AHDL:bsdMX) and FY38525 (h- ade6 + :patb2:mCherry-atb2:terminatoratb2:hphMX) provided by the NBRP (YGRC), Japan.
After thawing glycerol stocks of S. pombe stored at −80 °C, the cells were cultivated overnight in liquid YES medium (0.5% yeast extract (Biolife Italiana, Milano, Italy), 3% glucose, 0.01% L-leucine, 0.01% L-lysine, 0.01% L-histidine, 0.01% uracil, and 0.01% adenine sulphate (all purchased from Sigma-Aldrich, St. Louis, MO, USA)) at 30 °C with appropriate shaking to reach the exponential growth phase. After reaching the exponential phase, the culture was divided into two conditions: untreated control cells and cells exposed to NTP. Additionally, for flow cytometry analysis, a sample treated with bisphenol A (used as a positive control; Sigma-Aldrich, St. Louis, MO, USA) was included, and for live cell imaging, the untreated sample was cultivated in liquid YES medium.
Non-thermal plasma treatment
For NTP treatment, commercially available multi-hollow surface dielectric barrier discharge (MSDBD) (RPS30, Roplass s.r.o., Brno, Czech Republic) was used (Homola et al. 2020). MSDBD can generate NTP using various working gases including ambient air, oxygen, or nitrogen. NTP is generated within the hollows of the device, with the active species blown onto the samples via the working gas flow. The source is powered by alternating high voltage with a frequency of 20 kHz, an input power of 30 W, and a gas flow rate of 5 l/min. The sample was suspended in sterile deionised water located 2 mm from the plasma discharge zone within the Petri dish and subjected to pulse plasma exposure at 30 s intervals for a total of 90 s (3 × 30 s), 180 s (6 × 30 s), 270 s (9 × 30 s), and 360 s (12 × 30 s), with mixing after each interval. Exposure times were selected based on yeast survival rates as reported by Medvecká et al. (2025). Control cells were treated identically to NTP-treated cells in all experiments, including incubation in sterile deionised water without plasma exposure. For each experiment, cells were derived from the same overnight culture, which was divided into control and NTP-treated fractions. This experimental setup was applied consistently across all experiments, including RNA isolation and RNA sequencing.
Intracellular RONS analysis
To analyse the presence of intracellular RONS in NTP-treated cells, 2′,7′-dichlorodihydrofluorescein diacetate (H_2_DCFDA; Sigma-Aldrich, St. Louis, MO, USA) staining was used. H_2_DCFDA is a small lipophilic, cell-permeable molecule that is metabolised by cellular esterases into 2′,7′-dichlorodihydrofluorescein (H_2_DCF) in metabolically active cells. In the presence of RONS, H_2_DCF is oxidised to the fluorescent 2′,7′-dichlorofluorescein (DCF) form (Eruslanov and Kusmartsev 2009).
NTP-treated and untreated cells were washed and resuspended in 50 mM phosphate buffer (1 M KH_2_PO_4_, 1 M K_2_HPO_4_, both from Sigma-Aldrich, St. Louis, MO, USA; pH 7.8). Staining was performed according to Mániková et al. (2014) with incubation in the dark for 45 min at 30 °C without shaking. To distinguish live and dead cells, additional staining with 7-aminoactinomycin D (7AAD) (Sigma-Aldrich, St. Louis, MO, USA) was performed. Mitochondrial superoxide in NTP-treated (9 × 30 s) and untreated cells was detected using Mitosox Red (Thermo Fisher Scientific, Dreieich, Germany), which contains the fluorescent probe dihydroethidine. Within the mitochondrial matrix, this probe is oxidised to ethidium, which then intercalates into mitochondrial DNA (Robinson et al. 2006). Staining was performed for 10 min at 30 °C according to Mániková et al. (2014). As a positive control for both intracellular RONS and mitochondrial superoxide detection, cells were treated with 1 mM bisphenol A (Sigma-Aldrich, St. Louis, MO, USA) for 60 min at 30 °C and 350 rpm. RONS-positive cells were detected using Cytoflex S flow cytometer (Beckman Coulter, Krefeld, Germany) with fluorescein isothiocyanate (FITC) channel (525/40) for DCF detection, PC5.5 channel (690/50) for 7AAD detection, and phycoerythrin (PE) channel (585/42) for Mitosox Red detection. For correct separation of DCF-positive and 7AAD-positive cells, compensation was manually set during measurement. The data were analysed with CytExpert 2.4 software (Beckman Coulter, Inc., Krefeld, Germany, 2011–2019).
Fluorescent microscopy
The yeast strain 18487 containing labelled tubulin and endoplasmic reticulum was analysed with live cell imaging. Samples were prepared according to Merlini et al. (2017) and analysed with an inverted Olympus IX 83 fluorescent microscope equipped with a UPlanXApo 60x/1.42 oil objective (Olympus, Tokyo, Japan). Cells were recorded every 10 min for a total of 120 min using an Andor Zyla-4.2PUSB3 camera (Andor Technology Ltd., Belfast, Northern Ireland, UK). Each image acquisition consisted of a z-stack of ten focal planes, each with a height of 0.67 μm. To ensure optimal conditions throughout the imaging process, a stable temperature of 30 °C was maintained using the ibidi Heating System™ (ibidi GmbH, Gräfelfing, Germany). Cells were observed in transmitted light with a BF-FAST (bright field) filter set, a FFITC/470 filter set for visualisation of the endoplasmic reticulum, and a FTRIC/550 filter set (excitation wavelength 580 nm) for visualisation of tubulin. The resulting images were evaluated in the CellSens Dimension programme (Olympus, Tokyo, Japan).
RNA sequencing
To analyse changes in the transcriptome, whole cellular RNA was isolated from cells treated with plasma for 9 × 30 s and from untreated cells according to Collart and Oliveiro (1993). To distinguish between early and late responses to plasma treatment, RNA was isolated from cells immediately after treatment and 1 h post-treatment. For DNA degradation, RNA samples were treated using the RNA Clean & Concentrator kit (Zymo Research, Freiburg im Breisgau, Germany), followed by quantitative PCR to detect any remaining DNA fragments in the RNA samples (Supplemental Table S1).
RNA library preparation
Extracted RNA was quantified spectrophotometrically. One microgram of total RNA was used as input for the ribosomal depletion reaction. mRNA enrichment was performed using NEB Next Poly(A) mRNA Magnetic Isolation Module (New England Biolabs, Ipswich, MA, USA). mRNA samples were then used for library preparation according to the original protocol from the NEB Next Ultra II Directional RNA Library Prep Kit for Illumina (New England Biolabs, Ipswich, MA, USA). After reverse transcription, the cDNA was amplified by PCR using NEB Next Multiplex Oligos for Illumina (New England Biolabs, Ipswich, MA, USA) with nine amplification cycles. The final library was purified using NEB Next Sample Purification Beads. The concentration of samples was determined using the Qubit™ dsDNA HS Assay Kit with Qubit Fluorometer v.2 (Thermo Fisher Scientific, Waltham, MA, USA). Fragment size was assessed on the Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). Finally, paired-end sequencing (2×40 bp) was performed using the Illumina NextSeq 550 platform.
High-throughput sequencing data analysis
FASTQ files for each sample were concatenated and filtered for length and a minimum quality of 20 using Cutadapt (v4.0) (Martin 2011), aligned to the reference genome of S. pombe 972 h^−^ (GCA_000002945.2_ASM294v2) using RNA STAR (v. 2.7.8a) (Dobin et al. 2013) followed by counting of the mapped fragments with featureCounts (v1.6.4) (Liao et al. 2014). The quality of reads was assessed with FastQC (Andrews 2010) with MulitQC tool applied for the visualisation of aggregating results (Ewels et al. 2016). Expression values were given as FPKM (fragments per kilobase millions). Differential gene expression was assessed using DESeq2 (v2.11.40.7) (Love et al. 2014).
Statistical analysis
Data were analysed in Microsoft Excel (Microsoft Office 365) and Statgraphics Centurion XV.I (StatPoint, Inc. Warrenton, VA, USA). For detection of normal distribution, the Shapiro-Wilk test was used. Significant differences in normally distributed data were analysed using one-way and multifactor analysis of variance (ANOVA) at a p-value ≤ 0.05, followed by Tukey’s HSD (honestly significant difference) test. Significant differences in data without normal distribution were analysed using the Kruskal-Wallis test at a p-value ≤ 0.05 followed by the analysis of median notches.
Results
Non-thermal plasma treatment leads to an increase in intracellular RONS levels in yeast cells
Intracellular RONS levels were measured immediately after the treatment, 1 h post-treatment, and 2 h post-treatment. NTP treatment led to an increase in intracellular RONS only after longer treatment (Fig. 1). After the shortest treatment (3 × 30 s, total exposure time 90 s), the percentage of cells with elevated RONS levels (DCF-positive cells) and dead cells (7AAD-positive cells) did not differ from the untreated control (0 × 30 s) over the 2-h observation period. However, the 6 × 30 s (total exposure time 180 s) NTP treatment increased the percentage of DCF-positive cells by 47.77% compared to the untreated control immediately after the treatment. Nonetheless, 1 and 2 h post-treatment, the percentage of DCF-positive cells decreased to the level observed in the untreated control. The percentage of 7AAD-positive cells did not increase over the 2-h observation period (Fig. 1). In the 9 × 30 s (total exposure time 270 s) NTP-treated sample, an immediate increase in DCF-positive cells was observed, along with a slight increase in the percentage of 7AAD-positive cells compared to the untreated control. However, 1 h post-treatment, the percentage of DCF-positive cells remained at the same level as immediately after the treatment, whereas the percentage of 7AAD-positive cells increased by 19.89%. Two hours post-treatment, 75% of cells were 7AAD-positive (Fig. 1). Immediately after the longest treatment (12 × 30 s, total exposure time 360 s), the highest increase in the percentage of 7AAD-positive cells was detected, and this percentage continued to rise with prolonged incubation, reaching 97.65% 2 h post-treatment (Fig. 1).Fig. 1. The percentage of cells with a physiological level of intracellular RONS (DCF-negative cells), elevated levels of RONS (DCF-positive cells), and dead cells (7AAD-positive cells). 0 × 30 s treatment time represents the untreated control, while 3 × 30 s, 6 × 30 s, 9 × 30 s, and 12 × 30 s correspond to total plasma exposure times 90 s, 180 s, 270 s, and 360 s, respectively. 0, 1, and 2 h refer to incubation times post-treatment. The results were statistically evaluated separately for each incubation time (0 h, 1 h, or 2 h). Different letters indicate statistically significant differences at a p-value ≤ 0.05, according to Tukey’s HSD test. Bars sharing at least one common letter are not significantly different
According to multifactorial ANOVA, the duration of the treatment, post-treatment time, and their combination had a highly significant effect on cell viability and the formation of RONS. Taken together, the data suggest that longer exposure times led to an increase in intracellular oxidation levels and a reduction in cell viability. In contrast, longer post-treatment incubation times resulted in a decrease in intracellular oxidation levels as well as cell viability (Supplemental Table S2).
Based on the analysis of intracellular RONS level, the treatment time of 9 × 30 s was selected for mitochondrial superoxide measurement and subsequent experiments. NTP treatment led to a 35.3% increase in the percentage of cells with elevated mitochondrial superoxide levels immediately after the treatment, continuing to rise with longer incubation times (Fig. 2). Two hours post-treatment, 72.8% of the NTP-treated cell population was composed of cells with higher mitochondrial superoxide levels.Fig. 2. The percentage of cells with elevated levels of mitochondrial superoxide in untreated control (0 × 30 s) and NTP-treated cells (9 × 30 s) immediately after treatment (+ 0), 1 h post-treatment (+ 1), and 2 h post-treatment (+ 2). Different letters indicate statistically significant differences at a p-value ≤ 0.05, according to Tukey’s HSD test. Bars sharing at least one common letter are not significantly different
Non-thermal plasma treatment causes cell cycle arrest and tubulin depolymerisation
Cells with labelled tubulin and endoplasmic reticulum were analysed using fluorescent microscopy to study the effect of NTP treatment on cell division. To exclude the effect of cultivation in deionised water on cell division and survival, a control containing untreated cells incubated in YES medium (YES) was included. In untreated cells (0 × 30 s) or in YES, a significant fraction of actively dividing cells was observed. At the same time, there was no significant difference in cells incubated in deionised water and YES medium in any fraction. On the contrary, in the sample treated with NTP 9 × 30 s, less than 1% of cells divided during the 2-h observation period, and the percentage of dead cells increased significantly by 19% compared to the untreated control (Fig. 3a). In addition, tubulin depolymerisation throughout the entire cell cycle was observed in 69% of NTP-treated cells (Fig. 3b). Polymerised tubulin was visible as red lines in the intracellular space in both control samples throughout the entire analysis (Fig. 4a). Although transient depolymerisation occurred immediately after cell division even in control cells, microtubules re-polymerised shortly afterwards. In contrast, in NTP-treated cells, microtubules were not visible, and the red fluorescent signal remained dispersed throughout the whole cell for up to 2 h (Fig. 4b). After NTP treatment, only 30% of cells could polymerise microtubules compared to both control samples where nearly 100% of cells polymerized microtubules. This decrease was statistically significant (Fig. 3b).Fig. 3. Microscopic analysis of cell division and vitality (a), tubulin polymerisation dynamics (b), and tubulin localisation (c) in NTP-treated cells (9 × 30 s), untreated cells (0 × 30 s), and untreated cells incubated in YES medium (YES). Statistical analyses were performed separately for each panel. Within each panel, statistical comparisons were carried out across all groups, including different treatment conditions (0 × 30 s, 9 × 30 s, and YES) and cell categories (e.g. divided cells, non-divided cells, and dead cells). Different letters indicate statistically significant differences in medians at a p-value ≤ 0.05, according to the Kruskal-Wallis test followed by median notches analysis. Bars sharing at least one common letter are not significantly differentFig. 4Representative images of the tubulin state in S. pombe cells immediately after NTP treatment (0 min) and at 60 and 120 min during analysis. a Untreated cells (0 × 30 s) incubated in deionised water. b Cells treated with NTP for 9 × 30 s. Images were captured every 10 min for a period of 120 min, with tubulin labelled by fusion with the mCherry fluorescent protein. In the untreated cells (a), the formation of microtubules was visible as red filaments throughout the observation period. After NTP treatment (b), the standard tubulin organisation and cell division were no longer observed in the cells during the entire observation period. Scale bar represents 5 µm
In addition to tubulin depolymerisation, changes in tubulin localisation in cells treated with NTP for 9 × 30 s were observed. In untreated cells, a transient depolymerisation of microtubules was observed immediately after cell division, during which tubulin monomers were evenly distributed throughout the entire cell. However, microtubules rapidly repolymerised, and polymerised tubulin was subsequently visualised as red linear structures in the cytoplasm (Fig. 4a). In contrast, in NTP-treated cells, microtubules remained depolymerised for the entire duration of imaging, and the red fluorescent signal was not only dispersed throughout the cell but also accumulated in the nuclear region, as defined by the green fluorescent labelling of the endoplasmic reticulum (Fig. 5). This phenomenon was observed in approximately 27.4% of NTP-treated cells. Compared to both control groups (0 × 30 s and YES), where this localisation shift was not detected, the increase in nuclear tubulin signal was statistically significant (Fig. 3c).Fig. 5. Tubulin localisation in the nuclear region of S. pombe immediately after NTP treatment (0 min) and at 60 and 120 min during analysis. a Untreated cells (0 × 30 s) incubated in deionised water. b Cells treated with NTP for 9 × 30 s. Images were captured every 10 min for a 120-min observation period. Tubulin was labelled with an mCherry fluorescent protein fusion, and the endoplasmic reticulum was labelled with green fluorescent protein. In NTP-treated cells, a brighter circular red signal was observed approximately in the centre of the cell, corresponding to the nucleus, as indicated by endoplasmic reticulum localisation. Scale bar represents 5 µm
Non-thermal plasma treatment (9 × 30 s) does not have a significant effect on the transcriptome of S. pombe
To investigate the complex cellular response to NTP treatment, total RNA sequencing was performed. As in previous experiments, the 9 × 30 s treatment time was selected to examine both the immediate cellular response to NTP treatment and the subsequent response to NTP generated particles.
In cells analysed immediately after NTP treatment (+ 0), a statistically significant change in expression was detected in only one transcript, SPNCRNA.128 (rrk1), the expression of which decreased to 42% of the level observed in the untreated control. In contrast, cells incubated in NTP-treated water for 1 h (+ 1) exhibited significant changes in the expression of five transcripts, including SPNCRNA.128, the expression of which decreased to 36% of the untreated control incubated in deionised water for 1 h post-treatment (Fig. 6). In addition to rrk1, RNA sequencing revealed changes in the expression of four long non-coding RNAs (lncRNAs), each showing an approximately twofold increase in expression relative to untreated control cells (Fig. 6).Fig. 6. Statistically significant changes in transcript expression in S. pombe after NTP treatment. 9 × 30 s + 0 h refers to samples from which RNA was isolated immediately after NTP treatment, 9 × 30 s + 1 h refers to samples from which RNA was isolated 1 h post-treatment. Transcript expression is shown as relative expression normalised to the untreated control. Gene identifiers are given as PomBase systematic IDs (PomBase, 2025): SPNCRNA.128–rrk1 gene, SPNCRNA.1224–lncRNA, SPNCRNA.1467–lncRNA, SPNCRNA.1033–lncRNA, SPNCRNA.990–lncRNA
Discussion
Non-thermal plasma (NTP) generates a rich cocktail of reactive oxygen and nitrogen species (RONS) including hydroxyl radicals (^•^OH), ozone (O^3^), nitric oxide radicals (NO^•^), hydrogen peroxide (H_2_O_2_), and superoxide radicals (O_2_^•−^) (Adamovich et al. 2017; Medvecká et al. 2025). These highly reactive chemical species are the primary mediators of NTP's biological effects, interacting directly with various cellular macromolecules such as DNA, proteins, and lipids, leading to their oxidative modification and subsequent functional impairment (Birden et al. 2012). Understanding these specific chemical reactions at the cellular and molecular level is crucial for elucidating the precise mechanisms of NTP-induced biological responses.
The first part of our study focused on detecting RONS generated by NTP treatment within the intracellular space, as these molecules are known to cause oxidative damage. Elevated intracellular RONS levels following NTP exposure have been reported in several studies (Kim et al. 2025; Molina-Hernandez et al. 2022a, b; Pan et al. 2022). Given that NTP generates a wide spectrum of RONS with distinct half-lives, its effects may persist for extended periods, thereby contributing to long-term cellular response (Tsoukou et al. 2021). Consistent with this notion, elevated levels of hydrogen peroxide, nitrates, and nitrites were detected in our experimental setup for a prolonged period (up to 2 h) (Medvecká et al. 2025). Therefore, intracellular RONS levels were measured not only immediately after the treatment but also 1 and 2 h post-treatment. Our results revealed dynamic changes in the DCF-positive cell population across different treatment times. Shorter treatments (3 × 30 s and 6 × 30 s) neither significantly elevated RONS levels nor caused severe cellular damage, allowing the cells to effectively eliminate them. In cells subjected to prolonged NTP exposure (9 × 30 s and 12 × 30 s), the observed intracellular RONS levels appear to exceed the cells’ repair capacity, which was associated with increased cell death. This suggests that cells were not only unable to repair the damage, but that the level of intracellular RONS continued to rise. Importantly, previous temperature measurements performed under experimental conditions same as those used in this study showed no increase in sample temperature during plasma treatment, excluding thermal damage as a contributing factor (Medvecká et al. 2025). Measurements of mitochondrial superoxide further supported this observation, as NTP treatment was associated with increased superoxide levels, suggesting perturbation of mitochondrial redox balance. This may lead to oxidative damage of mitochondrial DNA, lipids, and proteins of the electron transport chain, potentially impairing ATP production. Such bioenergetic stress would have cascading effects on energy-dependent processes vital for cell division and growth (Palma et al. 2024). Moreover, superoxide can directly react with iron-sulphur clusters in proteins, leading to their inactivation (Djaman et al. 2004). Although yeasts do not undergo classical apoptosis due to the absence of several key components, such as caspases (Váchová and Palková 2007), elevated intracellular RONS levels and loss of mitochondrial membrane potential have been linked to regulated, apoptosis-like cell death pathways (Chaves et al. 2025). Previous studies investigating the biological effect of NTP have reported divergent outcomes with respect to apoptosis-like cell death. While some studies have demonstrated activation of apoptosis-like processes following plasma exposure, often associated with mitochondrial dysfunction and elevated intracellular RONS levels (Xu et al. 2020; Zhu et al. 2025), other studies did not detect activation of apoptosis-like cell death (Polčic et al. 2018). These discrepancies likely reflect differences in experimental conditions, including plasma source parameters and cell type. In the context of our study, the observed increase in mitochondrial superoxide levels and cell death is therefore consistent with the possible involvement of apoptosis-like processes; however, due to the lack of direct apoptotic markers, this interpretation remains hypothetical. As RNA sequencing was performed at early post-treatment time points, the transcriptomic data likely reflect transcriptional responses preceding the advanced execution of cell death pathways.
Although hydroxyl radicals are a plausible contributor given their known reactivity and presence in NTP-generated RONS mixtures, no RONS-specific scavengers, inhibitors, or genetic approaches were employed in this study. Therefore, the contribution of individual reactive species, including •OH, cannot be experimentally distinguished and should be regarded as speculative. Since the percentage of cells with elevated superoxide levels did not decrease during the post-treatment incubation period, it is likely that the damage was not efficiently repaired, or that intracellular RONS remained present and biologically active. Similar findings were reported by Molina-Hernandez et al. (2022a, b), who observed elevated intracellular RONS levels in Aspergillus chavalieri following NTP treatment, detected using a H_2_DCFDA probe. In their study, increased RONS levels were observed even 6 h post-treatment, suggesting that intracellular RONS can be continuously generated after exposure to NTP. Our results from mitochondrial and intracellular RONS detections align with this hypothesis, as elevated RONS levels persisted in our samples for up to 2 h post-treatment. Comparable findings were also reported by Itooka et al. (2018), who demonstrated sustained intracellular oxidative stress using a transcription factor responsive to the presence of hydrogen peroxide. Similarly, Čtvrtečková et al. (2019) detected superoxide radicals in the intracellular space of Saccharomyces cerevisiae following NTP treatment. Interestingly, the highest amount of superoxide was observed 8 days after NTP treatment, supporting the hypothesis of long-lasting RONS effects. In their study, NTP was also applied to strains lacking a mitochondrial genome, where a lower percentage of cells with elevated superoxide levels were detected. Moreover, these strains exhibited a higher survival rate compared to strains with an intact mitochondrial genome. These findings suggest that intracellular RONS detected in NTP-treated cells are likely generated endogenously by the cells themselves, rather than being directly introduced by NTP. Additionally, their results highlight the critical role of mitochondria in cell death, which aligns with conclusions from other studies (Itooka et al. 2018; Molina-Hernandez et al. 2022a, b; Strížová et al. 2023).
NTP-induced RONS negatively affect various cellular compartments and organelles, including the cytoplasmic membrane (Xu et al. 2021) and endoplasmic reticulum (Itooka et al. 2018), in addition to mitochondria. In our study, cells exposed to NTP also exhibited tubulin depolymerisation and were unable to form relatively stable microtubules during the 2-h observation period. It should be emphasised that our live-cell imaging captures only early cellular responses to NTP treatment within a limited observation window of up to 120 min post-exposure. Therefore, the observed tubulin depolymerisation reflects short-term effects, and no conclusions can be drawn regarding long-term microtubule recovery, repolymerisation, or adaptive remodelling at later timepoints. A similar phenomenon has been reported in myocytes treated with hydrogen peroxide (Drum et al. 2016), and in osteosarcoma cells subjected to oxidative stress induced by hydrogen peroxide treatment (Lee et al. 2005). In addition to tubulin depolymerisation, NTP-treated cells exhibited tubulin localisation in the nuclear region. In untreated cells, tubulin monomers were primarily localised in the cytoplasm, where they formed interphase microtubules. Prior to cell division, these monomers polymerised to form spindle filaments, which depolymerise following cell division and reorganise into new interphase microtubules after septum formation (Loncar et al. 2020). Localisation of tubulin in the nuclear region in untreated samples was observed only in dead cells. However, in NTP-treated cells, this relocalisation was visible even in live cells, some of which displayed regeneration during the observation period.
The observed tubulin depolymerisation and nuclear retention in NTP-treated cells represents a notable biological response. It is well-established that tubulin is highly susceptible to oxidative modification, particularly at cysteine residues, which can lead to structural changes, impaired polymerisation, and altered microtubule dynamics (Goldblum et al. 2021; Shields et al. 2024). Based on previous studies demonstrating the high susceptibility of tubulin cysteine residues to oxidative modification, we hypothesise that RONS generated by NTP may contribute to the observed impairment of tubulin polymerisation. However, this interpretation remains indirect and is based on analogy with oxidative stress models rather than direct evidence from the present study. In addition to the question of impaired polymerisation, the origin and persistence of tubulin monomers in the nuclear region require further consideration. Based on our results, we could not conclusively determine whether tubulin monomers observed in the nuclear region were remnants of the mitotic spindle (since S. pombe undergoes closed mitosis (McCully and Robinow 1971)) or whether they accumulate in the nuclear region through an active transport process. Although nuclear accumulation of tubulin has been described under specific stress conditions in other systems (Akoumianaki et al. 2009; Schwarzerová et al. 2006; Xu and Ludueña 2002), our data do not allow us to distinguish whether the observed nuclear signal represents persistent remnants of the mitotic spindle, altered nucleocytoplasmic trafficking, or passive redistribution following oxidative damage. Consequently, mechanistic interpretation of this phenomenon remains limited. This limitation is further reinforced by the experimental scope of the present study. It should be emphasised that our study captures only early cellular responses to NTP treatment, within 120 min post-exposure, and does not include RONS-specific inhibition strategies or direct biochemical assays targeting tubulin oxidation or microtubule dynamics. While our observations clearly demonstrate rapid cytoskeletal disruption following NTP exposure, the underlying molecular mechanisms remain to be experimentally validated. These limitations define important directions for future studies employing extended time-lapse imaging, targeted scavengers, and biochemical or genetic approaches.
To detect the complex cellular response to NTP treatment, we also focused on the transcriptome of NTP treated cells. RNA sequencing was intentionally performed at relatively early post-treatment time points (immediately and 1 h post-treatment), at which the proportion of non-viable cells remained low. This experimental design was chosen to minimise the potential loss of transcriptional information due to extensive cell death and to capture early transcriptional responses, including potential activation of stress-response or cell death-related pathways. Several studies have observed changes in gene expression, for instance in oxidative response, metabolism, transport, and DNA damage repair (Gao et al. 2025; Otsubo et al. 2023; Zhao et al. 2024). In our study, the limited changes in global gene expression, particularly in genes associated with oxidative stress response, are intriguing given the clear evidence of intracellular oxidation. These findings suggest that the primary cellular responses to NTP in S. pombe under the experimental conditions described in the present study may not involve extensive transcriptional reprogramming and could instead be associated with post-translational processes. This is consistent with a rapid chemical impact on existing cellular machinery, potentially preceding slower transcriptional responses. For example, direct oxidation of enzymes involved in detoxification or repair could compromise cellular defences without necessarily altering their gene expression (Kneeshaw et al. 2017; Tiwari et al. 2019). Expression changes were detected in five lncRNAs and three genes (rrk1, set2, and* mot2*). Only one of these genes, rrk1, showed a significant decrease in expression both immediately after the treatment and 1 h post-treatment. This gene encodes the catalytic subunit of the RNAse P enzyme, which is primarily responsible for processing 87 different pre-tRNAs into mature tRNAs. In addition to tRNA processing, RNase P also cleaves 4.5 S rRNA, riboswitches, and, in Escherichia coli, even mRNA (Phan et al. 2021). Based on this information, the observed decrease in rrk1 expression may be associated with reduced availability of the functional enzyme and altered tRNA maturation, which could potentially affect protein synthesis. Plasma-associated alternations in protein levels have been reported previously in S. cerevisiae (Chen et al. 2010); however, the molecular basis of these effects was not investigated in that study.
Based on the observed changes in gene expression, we hypothesise that NTP generated by MSDBD under our experimental conditions does not broadly alter gene expression at the transcriptomic level. Instead, it is more likely to impact global protein production by destabilising mRNA and impairing tRNA maturation. In addition, altered expression of lncRNAs can influence gene regulation through chromatin remodelling (Herman et al. 2022; Zhu et al. 2020) or splicing regulation (Pisignano and Ladomery 2021; Teng et al. 2021). Such disruptions could compromise the cell’s ability to synthesise functional proteins and enzymes, which may contribute to cell cycle arrest and cell death. It should be noted that these functional implications are inferred indirectly from transcriptomic data obtained at early post-treatment time points and require further experimental validation.
To further clarify whether NTP treatment induces oxidative damage, alters metabolic pathways, or compromises membrane integrity, alternative treatment times of shorter post-treatment incubation periods should be considered, as the conditions used in our study primarily resulted in cell death. Our findings, along with observations from other studies, indicate that NTP effects are highly variable and influenced by several factors, including the type of plasma source, the working gas, the distance from the source, and the treatment time. Therefore, it is difficult to generalise cellular responses to this stressor. Nonetheless, all studies consistently report that NTP exposure generates RONS, which are likely responsible for cellular damage. Given the variability in cellular responses, it is essential to investigate a wide range of conditions to better predict the effects of NTP in practical applications, not only on pathogens but also on host cells and tissues.
Future studies employing mass spectrometry–based redox proteomics would substantially strengthen mechanistic insight by enabling direct identification and quantification of oxidative protein modifications induced by NTP. Such analyses could identify specific oxidised residues, reveal the spectrum of affected proteins, and directly link NTP-generated reactive species to functional protein impairment.
Conclusion
Our study demonstrates that prolonged NTP exposure induces oxidative stress in S. pombe, characterised by increased intracellular RONS and mitochondrial superoxide. These chemical perturbations are consistent with oxidative damage to cellular components such as tubulin, although direct identification of oxidatively modified proteins was beyond the scope of the present study. While the transcription response appears limited, the observed effects underscore a direct and rapid chemical–biological interaction where RONS generated by NTP act as immediate triggers for profound cellular dysfunction, particularly affecting cytoskeletal integrity.
Supplementary Information
Below is the link to the electronic supplementary material.ESM 1(PDF 153 KB)
The reference list from the paper itself. Each links out to its DOI / PubMed record.
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