Research ArticleExperimental Studies
Open Access

Short-term Effects of Non-invasive Physical Plasma Treatment on Genomic Stability

NADINE GELBRICH, RAZAN MUHTADI, HARRY SCHERTHAN and MATTHIAS B. STOPE
In Vivo January 2024, 38 (1) 82-89; DOI: https://doi.org/10.21873/invivo.13413

Abstract

Background/Aim: The application of non-invasive physical plasma (NIPP) generates reactive oxygen species. These can lead to chemical oxidation of cellular molecules including DNA. On the other hand, NIPP can induce therapeutically intended apoptosis, which also leads to DNA fragmentation in the late phase. Therefore, to assess unwanted genotoxic effects, the formation of DNA damage was investigated in this study in discrimination from apoptotic processes. Materials and Methods: Mutation events after NIPP application were analyzed in CCL-93 fibroblast cells using the hypoxanthine phosphoribosyl transferase assay. Additionally, DNA single-strand breaks (SSB) and double-strand breaks (DSB) were quantified by performing the alkaline comet assay, and terminal deoxynucleotidyl transferase dUTP nick-end labeling assay. DSBs were quantified by phospho-histone 2AX-p53-binding protein 1 co-localization DSB focus assay. The data were compared with cell death quantification by the caspase-3/7 apoptosis assay. Results: Treatment with NIPP led to exceedingly rapid damage to genomic DNA and the appearance of DNA SSBs and DSBs in the initial 4 h. However, damage decreased again within the first 4-8 h, then the late phase began, characterized by DNA DSB and increasing caspase-3/7 activation. Conclusion: Although NIPP treatment leads to extremely rapid damage to genomic DNA, this damage is reversed very quickly by efficient DNA-repair processes. As a consequence, only those cells whose genome damage can be repaired actually survive and proliferate. Persistent genotoxic effects were not observed in the cell system used.

Key Words:

Physical plasma is a highly energized gas containing numerous free charge carriers. The latter possess high kinetic energies and can therefore generate further free charged particles by shock ionization. The high kinetic energy of this type of physical plasma generally induces high temperatures, restricting its use to technical processes. The development of devices for the generation of non-thermal physical plasma has allowed their use in medical applications (1). Medical non-thermal plasma only achieves temperatures slightly above body temperature. Due to its low temperature and low energy input into treated tissue, this physical plasma technology is tissue-protective and is also referred to as non-invasive physical plasma (NIPP). At the interface of NIPP and the ambient atmosphere, a further transfer of energy occurs between NIPP particles and particles of the atmosphere. As a consequence, mainly reactive oxygen (ROS) and nitrogen species are produced, which are primarily responsible for the biomedical efficacy of NIPP (2).

At present, NIPP is being tested with regard to its possible application in various medical fields. The longest experience exists in dermatology, where NIPP is used to reduce the bacterial load of miscolonized and infected skin regions to improve wound healing (2, 3). Oncology is another current field of research for the application of NIPP (4). Antiproliferative, antimetastatic and immunostimulatory effects of NIPP have been demonstrated in various malignancies and thus point to a novel and innovative application horizon in oncological therapy (5-9).

However, the composition of NIPP, namely its excited and highly reactive particles, suggests its strong chemical reactivity, especially with respect to oxidative processes. In fact, it has been shown that NIPP can lead to the chemical modification of biomolecules, including of lipids, proteins, and nucleic acids (10, 11). Nucleic acid modifications include the oxidation of DNA bases, as well as DNA single-strand breaks (SSB) and double-strand breaks (DSB). Moreover, DNA–protein cross-links induced by NIPP treatment can cause further damage as these are disruptive for replication and transcription processes (10). NIPP-induced nucleic acid modifications may also eventually lead to mutations and consequent harmful effects.

Initial studies on the DNA-damaging effects of NIPP were performed with isolated DNA and bacteria (12, 13). However, such data cannot be extrapolated easily to eukaryotic cells with their significantly higher complexity, including complex DNA-damage detection and repair machinery. Recent studies on DNA-damaging effects of NIPP in eukaryotic model systems have so far not provided evidence of mutagenic effects (14), while it is well proven that NIPP treatment can induce apoptosis (4, 5, 8). Therefore, it is difficult to distinguish NIPP-related primary DNA damage in fragment read-out experiments (e.g., Comet assay) from NIPP-induced apoptotic DNA degradation. In the present study, DNA damage was examined within the first 24 h after NIPP treatment using a set of different experimental approaches including apoptosis assays to differentiate DNA-damaging and apoptotic effects of NIPP treatment.

Materials and Methods

Cell culture. The Chinese hamster cell line CCL-93 (lung fibroblast; American Type Culture Collection, Manassas, VA, USA) were propagated in Dulbecco’s modified Eagle’s medium (DMEM) containing 4.5 g/l glucose supplemented with 10% fetal bovine serum, 1 mM sodium pyruvate, stable glutamine, 1.5 g/l NaHCO3 and 1% penicillin/streptomycin (all PAN Biotech, Aidenbach, Germany) in a humidified atmosphere at 37°C with 5% CO2.

NIPP treatment. For generation of NIPP, Kinpen Med (Neoplas Tools, Greifswald, Germany) plasma device was used with argon as carrier gas (gas flow: 3 l/min; supply voltage=65 V DC; frequency: 1.1 MHz). For NIPP treatment, 0.2×105 or 0.4×105 CCL-93 cells were suspended in 200 μl DMEM (PAN Biotech) on an uncoated 24-well cell culture plate and treated for 10 s with NIPP and argon-only as a control, respectively. After NIPP treatment, cells were immediately transferred to uncoated 24-well plates [for cell proliferation assay, hypoxanthine phosphoribosyl transferase (HPRT) assay, comet assay or DNA DSB assay], 96-well plates (5.0×103 suspended in 200 μl for terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) assay, or caspase-3/7 assay) and were incubated in DMEM (PAN Biotech) for 0.01 min, 5 min, 1 h, 4 h, 8 h and 24 h.

Cell proliferation assay. Cellular growth of CCL-93 cells was examined by cell counting utilizing a CASY Cell Counter and Analyzer Model TT (Roche Applied Science, Mannheim, Germany). Treated cells were detached by trypsin treatment, suspended in CASYton (Roche Applied Science) at 1:100 dilution and three replicates of 400 μl of cell suspension were analyzed. Measurement was performed using a capillary of 150 μm in diameter and cell line-specific gate settings to discriminate between living cells, dead cells, and cellular debris (6.88 μm/12.38 μm).

HPRT assay. For investigating gene mutations, CCL-93 cells were incubated with 10 μg/ml 6-thioguanine (300 μM; Roche Applied Science) (15). After an incubation of 24 h, cellular growth was examined by cell counting utilizing a CASY Cell Counter and Analyzer Model TT (Roche Applied Science) as mentioned above.

Comet assay. Cells were analyzed with a reagent kit for single-cell gel electrophoresis assay (alkaline) (Trevigen, Gaithersburg, MD, USA). Adherent cells of a 24-well cell culture plate were suspended and centrifuged at 300 rcf. The cell pellet was washed with phosphate-buffered saline (PAN Biotech) and mixed with Comet LMAgarose (Trevigen) in a 1:5 ratio. Subsequently, the cell suspension was coated onto a CometSlide and stored in the dark at 4°C for 20 min. After a 1-h incubation in Lysis Solution (4 ml/slide) at 4°C, the slides were transferred to the Alkaline Unwinding Solution (pH>13, 200 mM NaOH, 1 mM EDTA) and incubated at room temperature in the dark for 20 min. After electrophoresis (15 min, 14 V) with Alkaline Electrophoresis Solution (pH>13, 200 mM NaOH, 1 mM EDTA), slides were washed, fixed and stained with diluted SYBR Gold (10,000×SYBR Gold in dimethyl sulfoxide, 10 mm Tris-HCl pH 7.5, 1 mM EDTA) at room temperature for 30 min. Images of fragmented DNA (‘comet tail’) were then taken with a BZ-9000 fluorescent microscope system (Keyence, Osaka, Japan) and analyzed with BZ II Analyzer software (Keyence).

TUNEL assay. TUNEL analysis was performed using HT TiterTACS Assay Kit (Trevigen) following the supplier’s recommendations to detect 3′ ends of DNA strand breaks that occur during apoptosis. CCL-93 cells (5.0×103) were resuspended in 200 μl DMEM and treated with NIPP for 10 s. Argon-treated cells served as a control. After 24 h of culture, adherent cells were detached by 0.1% trypsin/0.04% EDTA. TUNEL data were acquired using an Infinite 200 PRO multimode reader (Tecan, Männedorf, Switzerland) and were analyzed using i-control 1.9 software (Tecan).

Phospho-histone 2AX (γ-H2AX)–p53-binding protein 1 (53BP1) assay. The formation of DNA DSBs was studied by fluorescence microscopy using the γ-H2AX) and p53-binding protein 1 (53BP1) co-localization DSB focus assay (16-18). γ-H2AX–53BP1 DSB foci were immunostained as described previously (19). Briefly, cell samples were exposed to NIPP, incubated for 0.01 min, 5 min, 1 h, 4 h, 8 h and 24 h and then fixed in 70% ethanol, followed by storage at −20°C. Cell suspensions (9.0×104 cells) were cytocentrifuged on glass slides and subjected to γ-H2AX–53BP1 co-immunostaining. Cells were then mounted in antifade (Rotimount; Carl Roth, Karlsruhe, Germany) and evaluated using a Zeiss Axioobserver 2 epifluorescence microscope (Carl Zeiss, Oberkochen, Germany) equipped with a green and red double-band pass filter (Chroma, AHF, Tübingen, Germany). γ-H2AX and 53BP1 co-localizing DSB foci in the nuclei were counted manually in 100 cells per sample by an experienced observer (R.M.). Values were expressed as average foci per cell (FPC).

Capase-3/7 activity assay. The caspase-3/7 activity assay was performed to distinguish apoptotic from live and necrotic cells using a CellEvent Caspase-3/7 Assay Kit (Thermo Scientific, Waltham, MA, USA). After NIPP treatment, cells were transferred to uncoated 96-well cell culture plates and were incubated in DMEM (PAN Biotech) for 0.01 min, 5 min, 1 h, 4 h, 8 h and 24 h. NIPP-treated CCL-93 cell culture plates were then washed, fixed and incubated with diluted green detection reagent CellEvent Caspase-3/7 (10 μM, phosphate-buffered saline, 5% fetal bovine serum) at 37°C for 45 min. Caspase-3/7 activity was determined using an Infinite 200 PRO multimode reader (Tecan) and were analyzed using i-control 1.9 software (Tecan).

Statistical analysis. Statistical analysis was performed using GraphPad Prism 5.0 software (GraphPad Software, La Jolla, CA, USA). Data shown are representative experiments or expressed as mean±standard deviation of at least three independent experiments. Significance was tested by t-test and set at p<0.05.

Results

NIPP inhibits cell growth but does not lead to mutations in the HRPT gene. In this study, we analyzed the effect of 10 s NIPP exposure on genomic DNA of CCL-93 cells. This fibroblast cell line is a well-accepted model for mammalian mutagenesis assays (20). If genotoxic noxae lead to the mutation of the HPRT gene, toxic 6-thioguanine is not incorporated into the DNA and mutant cells survive in the presence of the toxic purine analogue.

Compared to control cells, treatment with NIPP led to a significant reduction in the number of living cells over an incubation period of 120 h (Figure 1). Cells cultured in the presence of 6-thioguanine showed similarly inhibited cell growth with and without NIPP treatment (Figure 1). Thus, these experiments suggest that NIPP exposure had no mutagenic impact on the HPRT gene in CCL-93 cells.

Figure 1.
Figure 1.

Hypoxanthine phosphoribosyl transferase (HPRT) assay. To detect mutations of the HPRT gene, CCL-93 cells were treated with argon (Ctrl), non-invasive physical plasma (NIPP), 6-thioguanine (TG), alone and in combination. Cellular growth was examined by live cell counting over 120 h. Statistically significantly different from the control by t-test as follows: *p<0.050, **p≤0.010, and ***p≤0.001.

Early phase induction of DNA SSB and DSB by NIPP. Since no mutagenic effects were detectable in the HPRT assay, we next tested the induction of DNA strand breaks using the alkaline comet assay. This assay detects SSB and DSB and quantifies DNA damage as a DNA fragment ‘cloud’, like a comet tail, outside of the nucleus in single-cell gel electrophoresis (21). The comet assay revealed a significantly higher amount of fragmented DNA in NIPP-treated cells 1 h after exposure (Figure 2). At 4 and 8 h after treatment, the comet signal had decreased to the level of the control signal but increased slightly but significantly again 24 h after NIPP treatment (Figure 2).

Figure 2.
Figure 2.

Comet assay. To detect DNA single-strand and double-strand breaks, CCL-93 cells were treated with argon gas (Ctrl), or non-invasive physical plasma (NIPP) and fragmented DNA was microscopically detected by alkaline comet assay over a period of 24 h. ***Statistically significantly different from the control by t-test at p≤0.001.

To confirm these data, DNA SSB and DSB formation by 3′-end labelling of DNA ends was performed by TUNEL assay (22). In the first 5 min after NIPP treatment, control and NIPP-treated cells showed equally high TUNEL signals (Figure 3). After incubation of 1 h and 4 h, TUNEL signals decreased in controls, but the NIPP-treated cells displayed significantly higher TUNEL signals, indicating DNA SSB and DSB formation. At 8 and 24 h after NIPP treatment, the proportion of cells with TUNEL signals increased, while signals in the control cells further decreased. The late TUNEL signals in the NIPP-treated cells may stem from ensuing apoptotic cell death (see below).

Figure 3.
Figure 3.

Terminal deoxynucleotidyl transferase dUTP nick-end labeling assay. To confirm of DNA single-strand and double-strand breaks, CCL-93 cells were treated with argon (Ctrl), or non-invasive physical plasma (NIPP) and fragmented DNA was measured applying a HT TiterTACS Assay Kit (Trevigen, Gaithersburg, MD, USA) over a period of 24 h. Statistically significantly different from the control by t-test as follows: **p≤0.010 and ***p≤0.001.

Since the alkaline comet assay and TUNEL assay do not discriminate between DNA SSBs and DSBs, we next performed microscopic quantification of DNA DSB foci by the enumeration of foci co-localizing the DSB markers γ-H2AX and 53BP1 (23). NIPP treatment induced a very rapid increase in foci per cell (FPC) in control cells from 0.6±0.3 FPC at 0.01 min to 1.9±0.8 FPC 5 min post NIPP. As incubation time progressed, the average FPC in controls decreased to 1.1±0.8 at 1 h, 1.2±0.6 FPC at 4 h, 1.0±0.3 at 8 h, and 0.9±1.4 at 24 h reflecting the progression of DNA repair (Figure 4). NIPP treatment induced a significant increase of the average foci during the first 4 h after treatment compared to controls (5 min: 3.6±0.5 FPC, 1 h: 2.9±0.8 FPC, 4 h: 3.2±0.6 FPC). This was followed by a decrease to the level of the control cells after 24 h (Figure 4).

Figure 4.
Figure 4.

Phospho-histone 2AX–p53-binding protein 1 (γ-H2AX–53BP1) assay. To specifically detect DNA double-strand breaks, CCL-93 cells were treated with argon (Ctrl) or non-invasive physical plasma (NIPP) and foci of nuclear γ-H2AX–53BP1 co-localization were analyzed by fluorescence microscopy over a period of 24 h.

Late-phase induction of apoptosis by NIPP. To investigate whether apoptotic processes are the source of TUNEL-positive cells at late time points due to DNA degradation, we performed a caspase-3/7 assay. The caspase-3/7 signal increased within the first 8 h, with the control and NIPP-treated cells being statistically similar (Figure 5). At 24 h, the caspase-3/7 signal was further increased in NIPP-treated cells (p≤0.001 compared with the control), while in the control cells, lends remained similar to that at 8 h (Figure 5), suggesting NIPP induced cell death 24 h after exposure.

Figure 5.
Figure 5.

Caspase-3/7 assay. To detect apoptosis, CCL-93 cells were treated with argon (Ctrl), or non-invasive physical plasma (NIPP) and activation of apoptotic caspases was measured applying a CellEvent Caspase-3/7 Assay Kit (Thermo Scientific, Waltham, MA, USA) over a period of 24 h. ***Statistically significantly different from the control by t-test at p≤0.001.

Discussion

One of the most promising possibilities for the clinical application of NIPP is its use in antineoplastic therapies (4). This requires extensive evaluation of potential genotoxic effects in terms of patient and operator safety.

In the present study, we investigated NIPP effects on genomic stability of CLL-93 fibroblast cells, a cell model widely used in genotoxicity studies (20). DNA strand breaks were detected via alkaline comet assay (SSB, DSB), TUNEL assay (SSB, DSB), and γ-H2AX–53BP1 co-localization focus assay (DSB). Cell death was quantified by caspase-3/7 apoptosis assay (21, 24, 25). In the late phase of apoptosis, fragmentation of genomic DNA occurs, which also leads to the formation of DNA DSBs and thus to positive comet and TUNEL signals, while such cells display a pan-γ-H2AX formation in their nuclei (18, 26, 27). Recent studies have shown that apoptosis is detectable as early as about 24 h after NIPP treatment (28, 29). Therefore, the challenge of the presented experimental approach was to temporally resolve DNA damage within the first 24 h after NIPP exposure to discriminate DNA strand breaks caused by NIPP and those caused by apoptosis.

Analysis of NIPP-induced mutations by HPRT assay demonstrated no mutagenic effects from NIPP treatment within the incubation period. This observation agrees with other HPRT studies that all failed to demonstrate mutagenic effects of NIPP treatment (14, 30, 31). Furthermore, long-term experiments in humans and animals, have failed to reveal genotoxic actions of NIPP (32-34). However, recent literature has shown that NIPP exposure does result in nucleic acid damage in eukaryotic cells (35-39).

In line with the latter, we observed that NIPP treatment does lead to pronounced DNA damage. After NIPP exposure, DNA SSB and DSB events were rapidly detectable. Comet and TUNEL assay showed significant DNA SSB and DSB signal increases within minutes. The induced DNA damaged thereafter declined and approached the control values at 24 h post exposure, suggesting efficient DNA repair. However, over the course of the remaining 24-h incubation, signals slowly increased again. DNA DSBs, as determined by the γ-H2AX–53BP1 DSB focus assay, decreased up to 8 h after exposure. while the 24 h measurements showed strong variability. Thus, a distinction can be made between the initial DNA breaks formed during the early phase and DNA damage that occurs in the late phase. With respect to DNA damage occurring later after exposure, it remains possible that the commencement of cell death typically seen after 24 h that is associated with DNA and nuclear fragmentation may have influenced the later measurements.

Apoptotic cell death was thus addressed by the caspase-3/7 activation assay. In control and NIPP-treated cells, caspase signals increased slowly and consistently during the first 8 h, but at 24 h after NIPP treatment, apoptosis was significantly increased relative to the control. The high apoptotic background early after exposure in control and exposed cells is likely due to the mechanical stress that occurs during the exposure (bubbling) when using a cell suspension (5, 40). Overall, two effects seem to overlap. In the early phase, direct NIPP treatment causes rapid DNA damage (41), as described above. However, this declines with ongoing repair over subsequent hours. In the late phase, NIPP-induced apoptosis occurs, causing DNA damage again (5, 42).

The early phase post treatment correlates with the growth behavior of NIPP-treated cells. Bioinformatic analysis of NIPP-treated cells revealed two distinct effects that led to a reduced cell population growth. Firstly, a small fraction of treated cells is eliminated, likely by mechanical stress. As a result, the initial cell number is slightly reduced and total cell growth decreases compared to untreated control cells. Subsequently, surviving cells show a reduced growth rate throughout the further incubation period, likely due to cell-cycle arrest because of DNA damage and repair, additionally enhancing growth-inhibitory efficacy of NIPP (18, 43).

The initial rapid effect of cell elimination may be due to the very rapid and strong induction of DNA SSBs and DSBs. Application of NIPP leads to the formation of ROS, which dissolve in the liquid of the biological sample being treated. There, due to their extremely short half-life, ROS decay, chemically attack, or oxidize cellular structures (44, 45). NIPP-treated fluids therefore lead to effects in untreated cells comparable to direct NIPP treatment of the cells (6). However, redox stress alone can also lead to oxidative DNA damage (46, 47). Depending on the half-life of the ROS, NIPP efficacy can therefore last even after the initial NIPP treatment has ended. Due to high reactivity, ROS concentration decreases rapidly. Longer-lasting cellular and molecular effects are subsequently transmitted through intracellular damage, and signaling and effector cascades (4, 48). This explains why cellular responses provoked after a single short NIPP treatment are still detectable days after treatment.

Rapidly occurring NIPP-induced DNA SSBs and DSBs decrease due to various mechanisms of DNA repair over time before apoptotic cell death at late time points induces further DNA DSBs. This is also indicated by studies demonstrating activation of DNA-repair enzymes (37, 49, 50) and cell-cycle arrest (8, 51) after NIPP exposure. The rapid decrease in DNA SSB and DSB signals within the first 4-8 h after NIPP exposure indicates efficient DNA repair. This is particularly pronounced in the comet assay, indicating that NIPP induced a higher load of DNA SSBs, which agrees with oxidative stress being a major mechanism involved in NIPP effects.

Conclusion

Our data give rise to the following hypothesis: Treatment with NIPP leads to exceedingly rapid ROS-mediated damage to genomic DNA and the appearance of DNA SSBs and DSBs. Thereby, NIPP exposure leads to the activation of the DNA-damage response and repair, leading to the repair of extensive damage after a few hours. At more advanced time points after exposure, NIPP-induced apoptosis occurs and apoptotic DNA fragmentation in turn again leads to an increase in the formation of DNA DSB signals. Some of the NIPP-treated cells will initially die during early time points, which may be a response to initial DNA damage. However, in the remaining cells, this initial DNA damage can be repaired, allowing this subpopulation to survive. Depending on the duration of NIPP treatment and thus on the ROS concentration within the treated sample, apoptosis may be induced later in some of the cells. Our data show that NIPP induces DNA damage, which largely leads to cell death or is successfully repaired, showing that cells with gene mutations cannot survive or proliferate.

Footnotes

  • Authors’ Contributions

    Conceptualization: MBS and HS; methodology: NG and RM; formal analysis: NG, RM, HS, and MBS; supervision: MBS and HS; writing original draft: NG and MBS; original draft review and editing: MBS and HS.

  • Conflicts of Interest

    The Authors declare that they have no competing interests.

  • Received July 17, 2023.
  • Revision received September 4, 2023.
  • Accepted September 6, 2023.

This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY-NC-ND) 4.0 international license (https://creativecommons.org/licenses/by-nc-nd/4.0).

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Short-term Effects of Non-invasive Physical Plasma Treatment on Genomic Stability
NADINE GELBRICH, RAZAN MUHTADI, HARRY SCHERTHAN, MATTHIAS B. STOPE
In Vivo Jan 2024, 38 (1) 82-89; DOI: 10.21873/invivo.13413
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