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Article

Time-Resolved Repair of Clustered DNA Damage in γ-Irradiated Yeast Cells

by
Ana G. Sánchez
1,* and
Deborah J. Keszenman
1,2,*
1
Grupo de Biofisicoquímica, Departamento de Ciencias Biológicas, Centro Universitario Regional Litoral Norte, Universidad de la República, Salto 50000, Uruguay
2
Citizen Science Program, Bard College, Annandale-on-Hudson, NY 12504, USA
*
Authors to whom correspondence should be addressed.
DNA 2026, 6(2), 17; https://doi.org/10.3390/dna6020017
Submission received: 31 January 2026 / Revised: 23 March 2026 / Accepted: 27 March 2026 / Published: 1 April 2026
Browse Figures
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Abstract

Background/Objectives: Exposure of cells to ionizing radiation induces isolated DNA lesions, including single-strand breaks, apurinic/apyrimidinic sites, and oxidized bases, as well as clustered damages of different complexity. The latter types of damage are difficult to repair, and the failure to process them accurately and efficiently is related to the induction of mutagenesis, genomic instability, cancer, and aging. Since various types of clustered lesions may occur simultaneously after radiation exposure, leading to a complex architecture of DNA damage, the study of the concomitant formation and the removal kinetics of clustered DNA damage is important to determine the mutagenic and, consequently, the carcinogenic potential of ionizing radiation. Methods: With the aim of capturing real-time coexisting lesion types and assessing the repair kinetics of clustered damages, the simultaneous determination of double-strand breaks, apurinic/apyrimidinic site clusters, and oxypurine clusters induced by γ-irradiation of Saccharomyces cerevisiae yeast cells was performed immediately after exposure and at time intervals during incubation in Liquid Holding Recovery conditions. Results: Ionizing radiation induced lethal and mutagenic events, leading to a dose-dependent linear increase in double-strand breaks, apurinic/apyrimidinic site clusters, and oxypurine clusters. The kinetic study showed that double-strand break frequencies declined during Liquid Holding Recovery, although a transient increase was detected at early time points. At 160 Gy, apurinic/apyrimidinic site clusters repair was evident, whereas at 400 Gy the frequency of damage increased before returning to the initial value at 24 h. In contrast, oxypurine clusters showed no net increase in repaired lesions over 24 h. Conclusions: The complex nature and topological characteristics of ionizing radiation-induced clustered DNA damage may influence lesion processing. Also, ionizing radiation may disrupt redox cellular homeostasis, leading to DNA damage and delayed effects.

Graphical Abstract

1. Introduction

The biological consequences of ionizing radiation (IR) exposure, such as cell death and mutagenesis, are related to a broad variety of DNA damage. Approximately 30% of the IR-induced damage results from the local transfer of energy, causing ionizations and excitations distributed over nanoscale distances within cellular DNA [1,2,3,4]. However, 70% of IR damage is produced by reactive free radicals generated by the radiolysis of water of solvation that surrounds the DNA molecule and the water in the vicinity within 6 to 9 nm [5,6,7]. As a result, isolated DNA lesions, including single-strand breaks (SSBs), apurinic/apyrimidinic sites (AP sites), and oxidized bases, as well as clustered damage (also called multiple damaged sites), are produced. Bistranded clustered DNA damage is defined as two or more strand breaks, AP sites, and oxidized bases on opposite strands within a few helical turns [8,9,10]. These types of damage are difficult to repair, and the failure of the cellular processes to repair them accurately and efficiently is related to the induction of mutagenesis, genome instability, cancer, and aging [11,12,13,14,15].
Among the different types of clustered DNA damage, double-strand breaks (DSBs) are well-recognized deleterious lesions that must be repaired to prevent chromosomal fragmentation and preserve genome integrity [16,17,18]. IR-induced disruption of the phosphodiester backbone may cause SSBs to generate DSBs by coincidence of two SSBs on opposite strands separated by up to 10 bp. The complexity of these DBSs depends on the type of DNA strand ends and on the presence of neighboring DNA lesions, such as AP sites, among others [10,19,20,21,22,23]. IR also induces non-DSB clusters containing base damage, AP sites, and SSBs, which may or may not generate DSBs immediately after IR exposure [24,25,26,27,28]. In any case, the nature of the damages, as well as their aleatory combination, may lead to DNA damage of high complexity [13,29,30]. In addition, during processing of non-DSB clustered damage via different repair pathways, intermediate lesions such as SSBs may be generated, which can subsequently form DSBs. The AP sites and oxypurines are mainly processed by base excision repair (BER), while DSBs are subjected to recombinational repair [31,32]. In fact, the processing of a damaged base by BER opposite to an SSB may generate a DSB. Similarly, the processing of bistranded clustered base lesions or AP sites can also lead to DSBs [11,13,19].
Clustered damage repair kinetics is a key factor in determining the types, frequencies, and complexity of the different damages that may remain in cells. Repair mechanisms are triggered immediately after damage induction, and some types of lesions are rapidly processed [33,34,35]. In fact, the rapid repair kinetics of SSBs may, at least in part, prevent the formation of DSBs [36]. To gain more insight into the kinetics of clustered DNA damage processing and to study the formation of intermediate damages that may contribute to lethality and mutagenesis, we have investigated the repair kinetics of DSBs, AP site clusters, and oxypurine clusters induced in Saccharomyces cerevisiae by γ-radiation. Budding yeast is a well-known and widely used model organism for studying DNA damage and repair in eukaryotic cells [37].
The study of the formation and removal kinetics of clustered DNA damage is important to determine the mutagenic and, consequently, possible carcinogenic potential of exposure to a genotoxic agent such as IR. Several lines of evidence have shown the conversion of one type of damage into another type due to sequential steps of repair (oxidized base clusters into AP site clusters and/or DSBs) [11,13,20,38]. Based on the hypothesis that the frequency of AP site clusters and DSBs increase due to the generation of intermediate lesions during the early steps of clustered DNA damage processing, we have performed simultaneous determinations of DSBs, AP sites, and oxypurine clustered damages induced by γ-irradiation of stationary phase yeast cells immediately after exposure and at time intervals during incubation in Liquid Holding Recovery (LHR) conditions. Cell γ-irradiation induced lethal and mutagenic events, leading to a dose-dependent linear increase in DSBs, AP site clusters, and oxypurine clusters. The analysis of DSB repair over 24 h in LHR showed a decrease in DSB frequency; however, the levels of damage increased at early time points. At 160 Gy, AP site clusters repair was detectable, whereas at 400 Gy, AP site clusters frequencies initially rose and then returned by 24 h to the levels measured at t = 0 of LHR. The determination of the oxypurine repair kinetics showed no significant increase in the repaired oxypurine cluster frequency after 24 h, but transiently decreased frequencies were observed at early time points of LHR. The findings of an apparent lack of DNA damage repair and the increase in the level of damage during the early time points of LHR could be due to the complexity of the clustered damage, the generation of repair intermediates, such as SSBs, and/or the possible interference of competing repair proteins. Besides considering the repair of genomic damage, other cellular functions may be altered by IR that may have an impact on DNA, and thus lead to a delayed increase in the level of damage. In fact, exposure to IR triggers a rapid surge in reactive oxygen- and nitrogen-derived species (ROS and RNS) generated both directly by water radiolysis and indirectly through perturbed cellular redox pathways.

2. Materials and Methods

2.1. The Yeast Strain and Growth Conditions

The Saccharomyces cerevisiae wild-type haploid strain BY4742 (MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0, Euroscarf, Frankfurt, Germany) was used. Yeast cell cultures were initiated from a single colony and incubated in liquid nutrient medium YPD [1% yeast extract (US Biological, Cleveland, OH, USA), 2% bactopeptone (US Biological), and 2% glucose (Sigma, St. Louis, MO, USA)] at 30 °C with shaking aeration. After 72 h of incubation, an aliquot was inoculated into fresh YPD medium, and cells were grown to the stationary phase over a cell concentration of 1–1.5 × 108 cells/mL. Cell concentration was assessed microscopically by counting using a Neubauer chamber.

2.2. Ionizing Radiation Treatment

Cell samples were irradiated in double-distilled water at a concentration of 1 × 108 cells/mL and kept on ice before and after radiation treatment. Cells were exposed to γ-rays using a 60Co source (dose rate = 0.8 Gy/s, assessed by Fricke dosimetry, Chemistry Division, Brookhaven National Laboratory, USA; LET: 1.17 MeV and 1.33 MeV [39]) over a dose range from 0 to 400 Gy.

2.3. Survival Assay Determination

Immediately after IR treatment, cell aliquots were appropriately diluted, plated on solid nutrient medium YPDA (YPD solidified with 2% agar (US Biological)), and incubated at 30 °C. After 4 days, the number of colonies was counted, and the surviving fraction (S) was calculated as S(x) = Ns/No, where Ns is the number of surviving cells capable of generating clones per mL, No is the total number of treated cells per mL, and (x) is the absorbed dose of radiation.

2.4. Mutation Frequency Analysis

To determine mutagenesis, aliquots of the cells exposed to γ-rays were immediately platted on synthetic complete solid medium SC (0.67% Bacto-yeast nitrogen base without amino acids (Difco Laboratories, Detroit, MI, USA), 2% glucose (Sigma, St. Louis, MO, USA), 2% agar (US Biological, Salem, MA, USA) and 2% Drop-out mix (US Biological, Salem, MA, USA) supplemented with canavanine (60 mg/L, Sigma, St. Louis, MO, USA). Plates were incubated for 15 days at 30 °C. Thereafter, canavanine-resistant mutants arising from a forward mutation were scored [40,41]. The mutation frequency was calculated as M(x) = Nm/Ns, where Nm is the number of mutants, Ns is the number of treated cells, and (x) is the dose of ionizing radiation as described in Keszenman et al. [42]. The induced mutation frequency was derived by subtracting the spontaneous mutation frequency (0 Gy) from the mutation frequency quantified in irradiated cells.

2.5. DNA Repair Kinetics Assay

Immediately after radiation, cells were kept on ice until further processing. A close monitoring of the time between the end of radiation exposure and the start of cluster damage assessment was performed in all the experiments. The transport time from the radiation source facility to the laboratory where the samples were processed ranged from 3 to 5 min. In this manuscript, the start of the sample processing was considered as “immediately after radiation treatment (t = 0)”. Then, aliquots of cells exposed to 0, 160, and 400 Gy were suspended in double-distilled water and incubated at 30 °C with aeration by shaking for 24 h. The post-irradiation treatment, described by Weber, K.J. & Kiefer, J. [43] as Liquid Holding Recovery (LHR), was used. To assess DNA repair kinetics, samples were collected at different time points for DNA isolation (t = 0, 0.25, 0.5, 1.5, 3.0, 6.0, and 24 h).

2.6. DNA Damage Assessment

2.6.1. DNA Isolation

Aliquots of cells were washed once with phosphate-buffered saline (PBS), then twice with EDTA (50 mM, pH 7.5). DNA was isolated in agarose plugs as described in Keszenman and Sutherland [44]. In brief, cells were embedded in 2% low-melting-point agarose (InCert, FMC, Rockland, ME, USA), lysed with Lyticase (Sigma-Aldrich, St. Louis, MO, USA), incubated with RNase A (Sigma-Aldrich, St. Louis, MO, USA), and digested with Proteinase K (Roche Molecular Biochemicals, Indianapolis, IN, USA) at 37 °C. Three to five agarose plugs were generated for each experimental sample (controls, irradiated, and LHR time points).

2.6.2. Clustered DNA Damage Determination

DSB and non-DSB clusters (AP site clusters and oxidized purine clusters) were studied using the agarose plugs generated from each experimental sample. Non-DSB clusters were assessed as described in Bennett et al. [45] and Keszenman and Sutherland [44] by digestion with damage-specific enzymes: E. coli Endonuclease IV (Nfo protein) to identify AP site clusters, and the Fpg enzyme (formamidopyrimidine-DNA glycosylase, Sigma) for oxidized purine clusters. Their respective titrations and the optimal amounts of enzyme for yeast DNA were previously described in Keszenman and Sutherland [44]. Although Nfo protein recognizes and cleaves several types of AP sites (regular and oxidized) [44], specific configurations and/or the proximity of multiple lesions may impair enzymatic activity, thereby leading to undetected Nfo sites and underestimating the frequency of Nfo clusters [38]. The frequencies of DSBs, Nfo, and Fpg clusters were determined by neutral pulsed-field gel electrophoresis (PFGE) (running parameters: 2–160 s ramps, 165 V, 20 h, 9 °C), electronic imaging, and length average length analysis [44]. The clustered DNA damage frequencies quantified by the length average length analysis were calculated per Gbp. To relate the number of damages to the size of the yeast genome, the calculated frequencies are reported per Mbp.

2.7. Data Analysis

Mean values and the corresponding standard errors of the mean (S.E.M.) of at least three independent experiments were plotted using SigmaPlot 15. Statistical differences were assessed using One-way ANOVA and Dunnett’s post hoc to compare each time point to t = 0LHR (p < 0.05).

3. Results

3.1. Biological Outcomes of γ-Radiation Exposure: Survival and Mutagenesis

To quantitatively study DNA damage repair kinetics, we initially set the level of γ-ray-induced DNA damage by assessing the survival and mutagenic frequency dose responses in our experimental conditions. Stationary phase haploid yeast cells were exposed to γ-rays in the range of 0 to 400 Gy in double-distilled water at 4 °C.
The survival dose response shows a decrease in the surviving fraction (Figure 1A). The corresponding plot showed two exponential components, with a change in slope at 80 Gy and 20% survival. The calculated slope of the first component was 6 × 10−2 ± 9.4 × 10−3 Gy−1, while the slope of the second component was 6.4 × 10−3 ± 1.2 × 10−3 Gy−1. As shown, there was a 10-fold decrease in the calculated slope of the second exponential component of the dose response compared to the first. This indicated a decrease in cell sensitivity to IR, as higher doses were required to achieve the same level of lethality. In the assessment of mutagenesis, the observed spontaneous mutation frequency was 1.5 × 10−6 ± 8.9 × 10−7, in agreement with values previously reported for other yeast strains [42,46]. As shown in Figure 1B, the induced mutagenic frequency as a function of IR dose showed a linear increase. Based on the survival and mutagenesis responses, we selected two IR doses, 160 and 400 Gy, that, on the one hand, yielded survival fractions within the second exponential component of the dose–response curve and thus had similar γ-ray sensitivity. On the other hand, these doses induced different levels of mutagenesis, as evidenced by a 2-fold difference in the induced mutagenic frequency.

3.2. DNA Double-Strand Breaks, AP Site Clusters, and Oxypurine Clusters Induced Immediately After γ-Ray Exposure

Clustered DNA damage was assessed immediately after IR exposure in the range of 0–400 Gy, and the frequencies of DSB⋅Mbp−1, Nfo clusters⋅Mbp−1 and Fpg clusters⋅Mbp−1 were determined. Figure 2 shows that the frequency of the three types of clustered DNA damage increased linearly with the γ-ray dose. The calculated DSB yield, corresponding to the slope of the curve, was 1.42 ± 0.06 DSBs⋅Mbp−1 per Gy (Figure 2A). As described in Section 2, the IR-induced AP site clusters and oxypurine clusters were assessed by digestion with the damage-specific enzymes (Nfo protein and Fpg enzyme, respectively). Each enzyme recognizes a specific type of DNA damage and generates a strand break [44].
In our experimental system, the spontaneous level of AP site clusters was 0.010 ± 0.002 Nfo clusters⋅Mbp−1 (Figure 2B). The Nfo clusters frequencies at each IR-dose were notably lower than the DSB frequencies. Applying linear regression to the graph of Figure 2B resulted in a yield of 0.233 ± 0.017 Nfo clusters⋅Mbp−1 per Gy. In the case of oxypurine clusters, the spontaneous level was 0.062 ± 0.006 Fpg clusters⋅Mbp−1, and the Fpg frequencies were lower than the DSBs ones, but higher than the Nfo clusters frequencies. The calculated yield of oxypurine clusters was 0.653 ± 0.050 Fpg clusters⋅Mbp−1 per Gy (Figure 2C). The spontaneous levels of Nfo and Fpg clusters reported were in accordance with previously observed values [44].

3.3. Endogenous Clustered DNA Damage Before and During Liquid Holding Recovery

Clustered DNA damage is generated spontaneously at low levels in unirradiated cells [47,48]. Under our experimental conditions, both control and irradiated cells were maintained on ice before and after irradiation to minimize any DNA damage unrelated to the treatment. The delay between the end of radiation treatment (t = 0) and the start of the LHR repair assay was approximately 3–5 min. To determine whether there was a variation in the spontaneous damage level due to sample manipulation, the frequencies of DSBs, AP site clusters, and oxypurine clusters determined in the unirradiated samples at the start of LHR (t = 0 LHR) were compared to those measured in the control samples immediately after IR exposure. The difference in the damage levels for DSBs was 0.009 ± 0.03 DSBs⋅Mbp−1, and for AP site clusters was 0.008 ± 0.011 Nfo clusters⋅Mbp−1, but the differences were not statistically significant (DSB: p = 0.72; AP site clusters: p = 0.37). In the case of oxypurine clusters, a slight increase in the level of damage was observed (0.112 ± 0.043 Fpg clusters⋅Mbp−1, p = 0.002).

3.4. Repair Kinetics of DNA Clustered Damages Induced by IR Exposure

To study the repair kinetics of clustered DNA damage, irradiated cells were treated with the selected γ-ray doses (160 and 400 Gy) and allowed to repair under LHR conditions. Aliquots were collected at different time points to determine DSBs, AP site clusters, and oxypurine clusters. To analyze the magnitude of repair during LHR, the number of lesions at each time point was normalized to the one observed at t = 0 of the damage repair assessment (0LHR). Therefore, at t = 0LHR, the number of lesions repaired at this time point corresponds to zero. The values of clustered damage > 0 indicated repair, whereas a value < 0 represented an increase in the number of damages.

3.4.1. DSBs Repair Kinetics

As described for the unirradiated samples, the comparison of DSB frequencies induced by 160 and 400 Gy immediately after irradiation and at the onset of LHR showed no significant difference (p = 0.64 and p = 0.52, respectively). Since the disappearance of DSBs is interpreted as DSB repair [49], Figure 3A,B (160 Gy LHR and 400 Gy LHR) show an increase in the number of repaired DSBs as a function of LHR time. However, at the earliest time point tested (15 min of LHR), the magnitude of repaired DSBs was negative at 160 Gy, indicating an increase in DSBs (Figure 3A). Also, at both radiation doses, the LHR kinetics were hyperbolic, showing an initial phase of fast repair (see inserts in Figure 3). Interestingly, the ratio between the DSBs observed at 400 and 160 Gy (400:160 Gy) immediately after radiation was 2.38 ± 0.09, and remained at this level during LHR.

3.4.2. AP Site Clusters Repair Kinetics

Similarly to DSBs, no significant difference was observed in the AP site clusters frequencies induced by 160 Gy (p = 0.77) or 400 Gy (p = 0.17), assessed immediately after irradiation and at the onset of LHR. At 160 Gy, an initial rapid AP site repair kinetics was observed in the first 30 min of LHR, as shown by the increase in the number of repaired Nfo clusters (Figure 4A). On the contrary, at 400 Gy, no evident repair of AP site clusters was observed throughout the duration of LHR. In fact, an increase in AP site clusters was observed during the LHR. However, at 24 h, the level was similar to that at t = 0 (p = 0.43) (Figure 4B).

3.4.3. Oxypurine Clusters Repair Kinetics

The comparison of oxypurine cluster frequencies determined immediately after irradiation and at the start of LHR showed a significant difference at 160 Gy (p = 0.043), but was similar at 400 Gy (p = 0.92). At 160 Gy, the number of Fpg clusters decreased after 30 min in LHR treatment, although it was not statistically different from the value at t = 0 of LHR (p = 0.65). The linear regression showed a slight increase in the number of repaired oxypurine clusters, although the R2 value was 0.71 (Figure 5A and insert). At 400 Gy, the number of Fpg clusters increased during the first 1.5 h of LHR, and then a slow repair was observed, reaching at 24 h a similar level to that observed immediately after γ-irradiation (p = 0.18) (Figure 5B).

4. Discussion

A quantitative study of the formation and removal of clustered DNA damage is important for predicting lethality and mutagenicity caused by IR. This information may be useful for estimating radiation risks or for radiotherapy planning, alone or in combination with chemotherapy. As the first step toward gaining more insight into clustered damage formation, we characterized the survival response and mutagenicity of the BY4742 haploid Saccharomyces cerevisiae yeast strain under γ-radiation exposure to assess the extent of lethal and mutagenic effects arising from DNA damage. As expected, survival decreased exponentially with the IR dose (Figure 1A), and the induced mutagenic frequency increased linearly (Figure 1B). Several studies have shown that IR of different qualities induces a dose-dependent increase in lethality and mutagenesis in yeast cells as a consequence of DNA damage [50,51,52].
To investigate the DNA damage underlying the observed lethal and mutagenic responses, the formation of DSBs, AP site clusters, and oxypurine clusters as a function of γ-ray dose was studied (Figure 2). It is noteworthy that, under our experimental conditions, all types of damage were assessed using the same DNA sample embedded in agarose plugs. Each plug was divided into equal parts, which were used to determine DSBs, Nfo, or Fpg clusters. The three different types of damage show a linear dose–response with different slopes. The DNA damage frequencies decreased in the order: DSBs > Fpg clusters > Nfo clusters, as shown by the yield ratios: DSB:Fpg = 2.18, DSB:Nfo = 6.09, Fpg:Nfo = 2.80. In our experimental conditions, the comparison of DSBs with non-DSB damage showed that DSBs were the most frequent (61.4% vs. 38.6%). However, because pyrimidine oxidative lesions were not quantified in this study, the absence of these measurements may account for the apparent predominance of DSBs relative to non-DSB damage. Interestingly, results obtained after γ-irradiation of bacteriophage T7 DNA in phosphate or Tris buffers showed oxypurine clusters as the most frequent type of damage [53]. In addition, our findings need to be considered in light of the well-recognized limitation of the Nfo enzyme-based assay used to detect AP sites in the present study. Georgakilas et al. [38], using human monocytes, reported that high-density abasic clusters and specific configurations are not cleaved by the Nfo protein and, therefore, the Nfo assay underestimates the frequencies of AP site clusters. Also, when several lesions reside within a few helical turns, the initial incision can alter local DNA structure or generate strand discontinuities that hinder subsequent enzymatic access. As a result, only a subset of AP sites within a complex cluster may be detected. This is particularly relevant when considering that repair of complex clusters involves a hierarchical pattern, in which certain lesions are preferentially processed while others persist because of steric or biochemical constraints [19,38]. More recently, using AFM (Atomic Force Microscopy) to quantify X-ray base lesions (oxypurines and oxypyrimidines) in TK6 cells, Nakano et al. [21] reported a yield of 0.25 base damage per 106 bp per Gy. The difference in the observed yields of the various types of clustered damage induced by low-LET radiation may be related to the size of the targeted DNA.
Cell lethality and mutagenesis result from unrepaired or misrepaired DNA damage. Therefore, it is important to assess the dynamics of how cells handle the initial injury induced by IR, which may compromise cell integrity. To examine the kinetics of clustered DNA damage repair, the levels of DSBs, AP site clusters, and oxypurine clusters were assessed in cells with similar radiosensitivity after irradiation with 160 or 400 Gy of γ-rays, and then allowed to repair under LHR conditions.
The DSB repair kinetics studied after 160 Gy or 400 Gy γ-ray exposure showed an initial fast decrease in the damage frequency (Figure 3A,B). The DSB-repaired frequencies remained at the same extent of damage in the window time between 6 and 24 h of LHR (Figure 3A,B), indicating that the level of DSBs detected at 6 h (independently of the DSB source) persisted with no measurable additional repair or increase in generated lesions. It should be noted that the level of DSBs at 24 h was lower than t = 0 of LHR. These findings indicate that an efficient DSB repair process may underlie the levels of survival observed. The ratio of DSBs at 400 Gy/160 Gy at each time point held steady during the LHR study, indicating a dose-independent repair kinetics. However, at the earliest LHR time point examined (15 min), an increase in DSBs was detected at 160 Gy. This rise likely reflects DSBs generated during the processing of other forms of DNA damage, such as non-DSB clustered lesions, either alone or in combination with IR-induced SSBs [11,13,19]. Indeed, at 160 Gy, the AP site clusters repair kinetics was evident within the first 15 min of LHR (Figure 4A), a process expected to generate transient intermediate SSBs through the resolution of AP sites. If these newly formed SSBs, as well as the ones directly produced by the radiation exposure, are not rapidly repaired, they can be converted into DSBs. Also, the IR-induced oxidized base repair may contribute to the formation of newly SSBs [13]. Georgakilas et al. [38] reported that, in 28SC monocytes, additional DSBs possibly generated during cluster processing were detected after 2 days of radiation. It is well documented that oxidized base clusters can be converted to AP site clusters and/or DSBs by the generation of SSBs through the sequential action of glycosylase/lyase or endonuclease [13,38]. However, no significant repair of oxypurines was observed after γ-ray exposure to 160 Gy in our experimental conditions during the 24 h of LHR (Figure 5A). It has been shown that clusters containing tandem 7,8-dihydro-8-oxoguanine (8-OxoGua) opposite to an AP site exhibited delayed AP-site repair, although both lesion types are BER substrates [19]. Among the oxypurines, 8-OxoGua is a well-studied type of lesion, and it is recognized by 8-OxoGua DNA glycosylase 1 (OGG1). However, it has been reported that OGG1 substrate recognition and excision efficiency can be hindered by DNA conformational distortions, such as those generated within complex lesions. These structural constraints may impair the processing of oxypurines and contribute to their persistence within clustered damage sites [38,54,55]. But the observed unchanged levels of Fpg clusters during LHR may not be solely interpreted as the absence of damage repair, as they may arise from additional oxidized purines formed via ROS and RNS generated both directly by water radiolysis and indirectly through perturbed cellular redox pathways. In fact, IR may induce an oxidative shift, disrupting metabolic homeostasis and contributing to genomic instability. Importantly, beyond these radiochemical events, mitochondria play an important role in other IR biological effects, including sustained ROS/RNS production and altered redox signaling that amplify and prolong the cellular response to IR [56,57].
The analysis of AP site clusters and oxypurine clusters repair kinetics, determined after cell IR exposure to 400 Gy, showed a significant decrease in the frequency of repaired lesions (Figure 4B and Figure 5B). However, for both, the level of damage after 24 h of LHR was not significantly different from the one observed at t = 0 of LHR. Interestingly, the repair timeframe for each type of damage varies widely, ranging from seconds to hours [58,59]. Molecular dynamics studies using the divisive hierarchical clustering (DIANA) approach have revealed seven clusters of DNA repair proteins where the ones including BER and NER (nucleotide excision repair) proteins are recruited to damage sites within the first 10 s after damage occurrence, the cluster of some DSBs repair proteins within 83 s and the cluster including homologous recombination proteins approximately within 20 min [33]. This kinetics-based clustering of proteins involved in DNA repair sheds light on the notion that, at a given time after DNA damage, multiple proteins from different repair pathways coordinate the sequential molecular events of DNA repair. In this complex DNA repair scenario, where damage processing timing interacts with topological molecular changes, other variables have to be considered, such as the possible persistence of non-deleterious damage. These unrepaired or misrepaired lesions may remain unrepaired or be subject to further processing. Evidence indicates that multiple AP sites positioned in close proximity generate intrinsic structural inhibition by imposing steric and physical constraints on repair enzymes. In mammalian cells, it has been shown that the optimal activity of APE1 is compromised by these conformational constraints [38,60,61]. Also, several studies reported that the clustering of DNA lesions may cause interference in the recruitment of different components or factors of the different repair pathways, facilitating the generation of DSBs [13,21,30,62,63]. In fact, the nature, orientation, and spacing of base damages, AP sites or SSBs, within a bistranded cluster determine the repair efficiency by BER [60,64]. Both in vitro and in vivo investigations have shown a delay in the processing of clustered DNA damage by BER, thereby facilitating the persistence of DNA lesions and leading to an increase in mutation frequency [13,15]. The balance between restoring DNA integrity and the persistence of damage will determine whether normal cell functions are modified or maintained.

5. Conclusions

The present study provides an integrated analysis of the formation and repair kinetics of multiple types of γ-radiation-induced clustered DNA damages in Saccharomyces cerevisiae. By simultaneously monitoring DSBs, AP sites clusters, and oxypurine clusters, we showed that IR generates these lesions in a dose-dependent manner and that their removal follows distinct kinetic patterns. While DSBs and AP site clusters exhibited measurable repair during LHR, their transient early increases suggest complex processing dynamics. On the other hand, oxypurine clusters persisted, revealing the challenge posed by oxidative lesions within complex damage architectures. Together, these findings support the mechanistic framework presented in Figure 6 and contribute to highlighting the importance of the structural complexity and topological constraints of low LET IR-induced clustered DNA damage that may influence repair efficiency. They also reinforce the notion that IR perturbs cellular redox homeostasis, leading to both immediate and delayed genomic effects. Overall, our results help to elucidate mechanisms underlying genomic instability and mutagenesis and may be relevant to carcinogenic risk, although they do not directly demonstrate carcinogenic potential.

Author Contributions

Conceptualization, D.J.K.; methodology, A.G.S. and D.J.K.; writing—original draft preparation, A.G.S. and D.J.K.; writing—review and editing, A.G.S. and D.J.K. All authors have read and agreed to the published version of the manuscript.

Funding

This research was part of A.G.S.’s internship under the supervision of D.J.K. in Betsy M. Sutherland’s laboratory (NASA Grant: NNJ07HC731 and US Department of Energy Grant: BO-089 to B.M.S.) at Brookhaven National Laboratory, NY, USA.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding authors.

Acknowledgments

In memoriam of Betsy M. Sutherland (Brookhaven National Laboratory, NY, USA) and Elia Nunes (Facultad de Medicina, Udelar, Montevideo, Uruguay). The authors (A.G.S. and D.J.K.) acknowledge all the scientific lessons learned from these outstanding women scientists.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
IRIonizing radiation
ABSsAbasic sites
SSBsSingle-strand breaks
DSBsDouble-strand breaks
BERBase excision repair
NERNucleotide excision repair
LHRLiquid Holding Recovery
ROSReactive oxygen species
RNSReactive nitrogen species

References

  1. Alizadeh, E.; Orlando, T.M.; Sanche, L. Biomolecular damage induced by ionizing radiation: The direct and indirect effects of low-energy electrons on DNA. Annu. Rev. Phys. Chem. 2015, 66, 379–398. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  2. Goodhead, D.T. The initial physical damage produced by ionizing radiations. Int. J. Radiat. Biol. 1989, 56, 623–634. [Google Scholar] [CrossRef] [Scilit]
  3. Cucinotta, F.A. Modeling Clustered DNA Damage by Ionizing Radiation Using Multinomial Damage Probabilities and Energy Imparted Spectra. Int. J. Mol. Sci. 2024, 25, 12532. [Google Scholar] [CrossRef] [Scilit]
  4. Rezaee, M.; Adhikary, A. The Effects of Particle LET and Fluence on the Complexity and Frequency of Clustered DNA Damage. DNA 2024, 4, 34–51. [Google Scholar] [CrossRef] [Scilit]
  5. Hill, M.A. Track to the future: Historical perspective on the importance of radiation track structure and DNA as a radiobiological target. Int. J. Radiat. Biol. 2018, 94, 759–768. [Google Scholar] [CrossRef] [Scilit]
  6. Nikjoo, H.; Khvostunov, I.K. Biophysical model of the radiation-induced bystander effect. Int. J. Radiat. Biol. 2003, 79, 43–52. [Google Scholar] [CrossRef]
  7. Sevilla, M.D.; Becker, D.; Kumar, A.; Adhikary, A. Gamma and Ion-Beam Irradiation of DNA: Free Radical Mechanisms, Electron Effects, and Radiation Chemical Track Structure. Radiat. Phys. Chem. 2016, 128, 60–74. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  8. Goodhead, D.T. Initial events in the cellular effects of ionizing radiations: Clustered damage in DNA. Int. J. Radiat. Biol. 1994, 65, 7–17. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  9. Sutherland, B.M.; Bennett, P.V.; Sidorkina, O.; Laval, J. Clustered damages and total lesions induced in DNA by ionizing radiation: Oxidized bases and strand breaks. Biochemistry 2000, 39, 8026–8031. [Google Scholar] [CrossRef] [Scilit]
  10. Ward, J.F. The complexity of DNA damage: Relevance to biological consequences. Int. J. Radiat. Biol. 1994, 66, 427–432. [Google Scholar] [CrossRef] [Scilit]
  11. Eccles, L.J.; O’Neill, P.; Lomax, M.E. Delayed repair of radiation induced clustered DNA damage: Friend or foe? Mutat. Res. 2011, 711, 134–141. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  12. Evstratova, E.S.; Petin, V.G.; Geraskin, S.A. DNA Repair Does Not Trigger the Genetic Instability of Yeast Cells. Russ. J. Genet. 2023, 59, 623–626. [Google Scholar] [CrossRef] [Scilit]
  13. Sage, E.; Shikazono, N. Radiation-induced clustered DNA lesions: Repair and mutagenesis. Free Radic. Biol. Med. 2017, 107, 125–135. [Google Scholar] [CrossRef] [Scilit]
  14. White, R.R.; Vijg, J. Do DNA Double-Strand Breaks Drive Aging? Mol. Cell 2016, 63, 729–738. [Google Scholar] [CrossRef] [Scilit]
  15. Kozmin, S.G.; Eot-Houllier, G.; Reynaud-Angelin, A.; Gasparutto, D.; Sage, E. Dissecting Highly Mutagenic Processing of Complex Clustered DNA Damage in Yeast Saccharomyces cerevisiae. Cells 2021, 10, 2309. [Google Scholar] [CrossRef] [Scilit]
  16. Goodarzi, A.A.; Jeggo, P.A. The repair and signaling responses to DNA double-strand breaks. Adv. Genet. 2013, 82, 1–45. [Google Scholar] [CrossRef] [Scilit]
  17. Her, J.; Bunting, S.F. How cells ensure correct repair of DNA double-strand breaks. J. Biol. Chem. 2018, 293, 10502–10511. [Google Scholar] [CrossRef] [Scilit]
  18. Mehta, A.; Haber, J.E. Sources of DNA double-strand breaks and models of recombinational DNA repair. Cold Spring Harb. Perspect. Biol. 2014, 6, a016428. [Google Scholar] [CrossRef] [Scilit]
  19. Eccles, L.J.; Lomax, M.E.; O’Neill, P. Hierarchy of lesion processing governs the repair, double-strand break formation and mutability of three-lesion clustered DNA damage. Nucleic Acids Res. 2010, 38, 1123–1134. [Google Scholar] [CrossRef] [Scilit]
  20. Kozmin, S.G.; Sedletska, Y.; Reynaud-Angelin, A.; Gasparutto, D.; Sage, E. The formation of double-strand breaks at multiply damaged sites is driven by the kinetics of excision/incision at base damage in eukaryotic cells. Nucleic Acids Res. 2009, 37, 1767–1777. [Google Scholar] [CrossRef] [Scilit]
  21. Nakano, T.; Akamatsu, K.; Tsuda, M.; Tujimoto, A.; Hirayama, R.; Hiromoto, T.; Tamada, T.; Ide, H.; Shikazono, N. Formation of clustered DNA damage in vivo upon irradiation with ionizing radiation: Visualization and analysis with atomic force microscopy. Proc. Natl. Acad. Sci. USA 2022, 119, e2119132119. [Google Scholar] [CrossRef] [Scilit]
  22. Stewart, R.D. Induction of DNA Damage by Light Ions Relative to 60Co gamma-rays. Int. J. Part. Ther. 2018, 5, 25–39. [Google Scholar] [CrossRef] [Scilit]
  23. Yajima, H.; Fujisawa, H.; Nakajima, N.I.; Hirakawa, H.; Jeggo, P.A.; Okayasu, R.; Fujimori, A. The complexity of DNA double strand breaks is a critical factor enhancing end-resection. DNA Repair 2013, 12, 936–946. [Google Scholar] [CrossRef] [Scilit]
  24. Cunniffe, S.; O’Neill, P.; Greenberg, M.M.; Lomax, M.E. Reduced repair capacity of a DNA clustered damage site comprised of 8-oxo-7,8-dihydro-2′-deoxyguanosine and 2-deoxyribonolactone results in an increased mutagenic potential of these lesions. Mutat. Res. 2014, 762, 32–39. [Google Scholar] [CrossRef] [Scilit]
  25. Georgakilas, A.G.; O’Neill, P.; Stewart, R.D. Induction and repair of clustered DNA lesions: What do we know so far? Radiat. Res. 2013, 180, 100–109. [Google Scholar] [CrossRef] [Scilit]
  26. Gulston, M.; de Lara, C.; Jenner, T.; Davis, E.; O’Neill, P. Processing of clustered DNA damage generates additional double-strand breaks in mammalian cells post-irradiation. Nucleic Acids Res. 2004, 32, 1602–1609. [Google Scholar] [CrossRef] [Scilit]
  27. Harrison, J.C.; Haber, J.E. Surviving the breakup: The DNA damage checkpoint. Annu. Rev. Genet. 2006, 40, 209–235. [Google Scholar] [CrossRef] [Scilit]
  28. Nikitaki, Z.; Nikolov, V.; Mavragani, I.V.; Plante, I.; Emfietzoglou, D.; Iliakis, G.; Georgakilas, A.G. Non-DSB clustered DNA lesions. Does theory colocalize with the experiment? Radiat. Phys. Chem. 2016, 128, 26–35. [Google Scholar] [CrossRef] [Scilit]
  29. Lomax, M.E.; Folkes, L.K.; O’Neill, P. Biological consequences of radiation-induced DNA damage: Relevance to radiotherapy. Clin. Oncol. 2013, 25, 578–585. [Google Scholar] [CrossRef] [Scilit]
  30. Schipler, A.; Iliakis, G. DNA double-strand-break complexity levels and their possible contributions to the probability for error-prone processing and repair pathway choice. Nucleic Acids Res. 2013, 41, 7589–7605. [Google Scholar] [CrossRef] [Scilit]
  31. Nickoloff, J.A.; Sharma, N.; Taylor, L. Clustered DNA Double-Strand Breaks: Biological Effects and Relevance to Cancer Radiotherapy. Genes 2020, 11, 99. [Google Scholar] [CrossRef] [Scilit]
  32. Thompson, P.S.; Cortez, D. New insights into abasic site repair and tolerance. DNA Repair 2020, 90, 102866. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  33. Aleksandrov, R.; Dotchev, A.; Poser, I.; Krastev, D.; Georgiev, G.; Panova, G.; Babukov, Y.; Danovski, G.; Dyankova, T.; Hubatsch, L.; et al. Protein Dynamics in Complex DNA Lesions. Mol. Cell 2018, 69, 1046–1061.e5. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  34. Mortusewicz, O.; Amé, J.-C.; Schreiber, V.; Leonhardt, H. Feedback-regulated poly(ADP-ribosyl)ation by PARP-1 is required for rapid response to DNA damage in living cells. Nucleic Acids Res. 2007, 35, 7665–7675. [Google Scholar] [CrossRef] [Scilit]
  35. Polo, S.E.; Jackson, S.P. Dynamics of DNA damage response proteins at DNA breaks: A focus on protein modifications. Genes. Dev. 2011, 25, 409–433. [Google Scholar] [CrossRef] [Scilit]
  36. Ogorek, B.; Bryant, P.E. Repair of DNA single-strand breaks in X-irradiated yeast: II. Kinetics of repair as measured by the DNA-unwinding method. Mutat. Res./DNA Repair Rep. 1985, 146, 63–70. [Google Scholar] [CrossRef] [Scilit]
  37. Botstein, D.; Fink, G.R. Yeast: An Experimental Organism for 21st Century Biology. Genetics 2011, 189, 695–704. [Google Scholar] [CrossRef] [Scilit]
  38. Georgakilas, A.G.; Bennett, P.V.; Wilson, D.M., III; Sutherland, B.M. Processing of bistranded abasic DNA clusters in γ-irradiated human hematopoietic cells. Nucleic Acids Res. 2004, 32, 5609–5620. [Google Scholar] [CrossRef] [Scilit]
  39. Buchtela, K. RADIOCHEMICAL METHODS|Gamma-Ray Spectrometry. In Encyclopedia of Analytical Science, 2nd ed.; Worsfold, P., Townshend, A., Poole, C., Eds.; Elsevier: Oxford, UK, 2005; pp. 72–79. [Google Scholar]
  40. Brendel, M.; Marisco, G.; Ganda, I.; Wolter, R.; Pungartnik, C. DNA repair mutant pso2 of Saccharomyces cerevisiae is sensitive to intracellular acetaldehyde accumulated by disulfiram-mediated inhibition of acetaldehyde dehydrogenase. Genet. Mol. Res. 2010, 9, 48–57. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  41. Cassier, C.; Chanet, R.; Henriques, J.A.P.; Moustacchi, E. The effects of three pso genes on induced mutagenesis: An novel class of mutationally defective yeast. Genetics 1980, 96, 841–857. [Google Scholar] [CrossRef] [Scilit]
  42. Keszenman, D.J.; Carmen Candreva, E.; Nunes, E. Cellular and molecular effects of bleomycin are modulated by heat shock in Saccharomyces cerevisiae. Mutat. Res. 2000, 459, 29–41. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  43. Weber, K.J.; Kiefer, J. Recovery from transcription inhibition in irradiated yeast cells. Br. J. Cancer Suppl. 1984, 6, 157–161. [Google Scholar] [PubMed]
  44. Keszenman, D.J.; Sutherland, B.M. Yields of clustered DNA damage induced by charged-particle radiations of similar kinetic energy per nucleon: LET dependence in different DNA microenvironments. Radiat. Res. 2010, 174, 238–250. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  45. Bennett, P.V.; Cintron, N.S.; Gros, L.; Laval, J.; Sutherland, B.M. Are endogenous clustered DNA damages induced in human cells? Free Radic. Biol. Med. 2004, 37, 488–499. [Google Scholar] [CrossRef] [Scilit]
  46. Gou, L.; Bloom, J.S.; Kruglyak, L. The genetic basis of mutation rate variation in yeast. bioRxiv 2018. [Google Scholar] [CrossRef] [Scilit]
  47. Bennett, P.; Ishchenko, A.A.; Laval, J.; Paap, B.; Sutherland, B.M. Endogenous DNA damage clusters in human hematopoietic stem and progenitor cells. Free. Radic. Biol. Med. 2008, 45, 1352–1359. [Google Scholar] [CrossRef] [Scilit]
  48. Keszenman, D.J.; Kolodiuk, L.; Baulch, J.E. DNA damage in cells exhibiting radiation-induced genomic instability. Mutagenesis 2015, 30, 451–458. [Google Scholar] [CrossRef] [Scilit]
  49. Blöcher, D.; Pohlit, W. DNA Double Strand Breaks in Ehrlich Ascites Tumour Cells at Low Doses of X-rays. II. Can Cell Death be Attributed to Double Strand Breaks? Int. J. Radiat. Biol. Relat. Stud. Phys. Chem. Med. 1982, 42, 329–338. [Google Scholar] [CrossRef] [Scilit]
  50. Bennett, C.B.; Lewis, L.K.; Karthikeyan, G.; Lobachev, K.S.; Jin, Y.H.; Sterling, J.F.; Snipe, J.R.; Resnick, M.A. Genes required for ionizing radiation resistance in yeast. Nat. Genet. 2001, 29, 426–434. [Google Scholar] [CrossRef] [Scilit]
  51. Löbrich, M.; Ikpeme, S.; Haub, P.; Weber, K.J.; Kiefer, J. DNA double-strand break induction in yeast by X-rays and alpha-particles measured by pulsed-field gel electrophoresis. Int. J. Radiat. Biol. 1993, 64, 539–546. [Google Scholar] [CrossRef] [Scilit]
  52. Magni, G.E.; Panzeri, L.; Sora, S. Sensitivity to ionizing radiations and damage repair in yeast. Int. J. Radiat. Oncol. Biol. Phys. 1979, 5, 1085–1088. [Google Scholar] [CrossRef] [Scilit]
  53. Sutherland, B.M.; Bennett, P.V.; Weinert, E.; Sidorkina, O.; Laval, J. Frequencies and relative levels of clustered damages in DNA exposed to gamma rays in radioquenching vs. nonradioquenching conditions. Environ. Mol. Mutagen. 2001, 38, 159–165. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  54. Wang, J.; Li, C.; Han, J.; Xue, Y.; Zheng, X.; Wang, R.; Radak, Z.; Nakabeppu, Y.; Boldogh, I.; Ba, X. Reassessing the roles of oxidative DNA base lesion 8-oxoGua and repair enzyme OGG1 in tumorigenesis. J. Biomed. Sci. 2025, 32, 1. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  55. Whitaker, A.M.; Stark, W.J.; Freudenthal, B.D. Processing oxidatively damaged bases at DNA strand breaks by APE1. Nucleic Acids Res. 2022, 50, 9521–9533. [Google Scholar] [CrossRef] [Scilit]
  56. Azzam, E.I.; Jay-Gerin, J.P.; Pain, D. Ionizing radiation-induced metabolic oxidative stress and prolonged cell injury. Cancer Lett. 2012, 327, 48–60. [Google Scholar] [CrossRef] [Scilit]
  57. Yang, S.; Li, Y.; Zhang, J.; Shen, A.; Hu, B.; Yan, J. The interplay between DNA damage response and mitochondrial dysfunction in radiotherapy. Front. Oncol. 2025, 15, 1642100. [Google Scholar] [CrossRef] [Scilit]
  58. Greenberg, M.M. Abasic and oxidized abasic site reactivity in DNA: Enzyme inhibition, cross-linking, and nucleosome catalyzed reactions. Acc. Chem. Res. 2014, 47, 646–655. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  59. Kitsera, N.; Rodriguez-Alvarez, M.; Emmert, S.; Carell, T.; Khobta, A. Nucleotide excision repair of abasic DNA lesions. Nucleic Acids Res. 2019, 47, 8537–8547. [Google Scholar] [CrossRef] [Scilit]
  60. Paap, B.; Wilson, D.M., 3rd; Sutherland, B.M. Human abasic endonuclease action on multilesion abasic clusters: Implications for radiation-induced biological damage. Nucleic Acids Res. 2008, 36, 2717–2727. [Google Scholar] [CrossRef] [Scilit]
  61. Singh, V.; Kumari, B.; Das, P. Repair efficiency of clustered abasic sites by APE1 in nucleosome core particles is sequence and position dependent. RSC Adv. 2015, 5, 23691–23698. [Google Scholar] [CrossRef] [Scilit]
  62. Asaithamby, A.; Chen, D.J. Mechanism of cluster DNA damage repair in response to high-atomic number and energy particles radiation. Mutat. Res. Fundam. Mol. Mech. Mutagen. 2011, 711, 87–99. [Google Scholar] [CrossRef] [Scilit]
  63. Schwan, L.; Averbeck, N.B.; Durante, M.; Jakob, B. Base-excision repair increases DNA double-strand break clustering within heavy-ion tracks and modulates repair at δ-electron-induced breaks. Sci. Rep. 2026, 16, 1339. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  64. Hsu, C.W.; Conrad, J.W.; Sowers, M.L.; Baljinnyam, T.; Herring, J.L.; Hackfeld, L.C.; Hatch, S.S.; Sowers, L.C. A combinatorial system to examine the enzymatic repair of multiply damaged DNA substrates. Nucleic Acids Res. 2022, 50, 7406–7419. [Google Scholar] [CrossRef] [Scilit] [PubMed]
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Figure 1. Survival and mutagenic response of yeast cells to γ-rays exposure. (A) Cell survival dose–response to γ-rays (0–400 Gy) of stationary phase yeast cells. Data points correspond to mean values ± S.E.M. of at least three independent experiments, and the solid line represents the non-linear regression. (B) Induced mutation frequency as a function of γ-rays dose. Each symbol represents the mean ± S.E.M. of at least three independent experiments. Linear regression is plotted as a solid line, and the 95% confidence interval is represented by dashed lines.
Figure 1. Survival and mutagenic response of yeast cells to γ-rays exposure. (A) Cell survival dose–response to γ-rays (0–400 Gy) of stationary phase yeast cells. Data points correspond to mean values ± S.E.M. of at least three independent experiments, and the solid line represents the non-linear regression. (B) Induced mutation frequency as a function of γ-rays dose. Each symbol represents the mean ± S.E.M. of at least three independent experiments. Linear regression is plotted as a solid line, and the 95% confidence interval is represented by dashed lines.
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Figure 2. Clustered DNA damages per Mbp induced by γ-rays. Frequency of DSBs (A), AP sites (Nfo) clusters (B), and oxidized purines (Fpg) clusters (C), induced by IR exposure of yeast cells to 0–400 Gy. Linear regressions are plotted as a solid line, and the 95% confidence interval is shown as dashed lines. Note: the y-axis scale in (B) is reduced by a factor of 5 compared to (A,C).
Figure 2. Clustered DNA damages per Mbp induced by γ-rays. Frequency of DSBs (A), AP sites (Nfo) clusters (B), and oxidized purines (Fpg) clusters (C), induced by IR exposure of yeast cells to 0–400 Gy. Linear regressions are plotted as a solid line, and the 95% confidence interval is shown as dashed lines. Note: the y-axis scale in (B) is reduced by a factor of 5 compared to (A,C).
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Figure 3. DSBs repair kinetics of γ-irradiated yeast cells after Liquid Holding Recovery. Repaired DSBs per Mbp at 160 Gy (A) and 400 Gy (B) as a function of LHR time. Means ± S.E.M. of at least three independent experiments are presented. Inserts: Non-linear regressions are plotted as a solid line, and the 95% confidence interval is shown as dashed lines.
Figure 3. DSBs repair kinetics of γ-irradiated yeast cells after Liquid Holding Recovery. Repaired DSBs per Mbp at 160 Gy (A) and 400 Gy (B) as a function of LHR time. Means ± S.E.M. of at least three independent experiments are presented. Inserts: Non-linear regressions are plotted as a solid line, and the 95% confidence interval is shown as dashed lines.
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Figure 4. Repair of γ-ray induced AP site clusters during LHR. Repaired Nfo clusters per Mbp at 160 Gy (A) and 400 Gy (B) as a function of LHR time. Means ± S.E.M. of at least three independent experiments are presented. (*) indicates a statistically significant difference compared to t = 0 LHR (p < 0.05).
Figure 4. Repair of γ-ray induced AP site clusters during LHR. Repaired Nfo clusters per Mbp at 160 Gy (A) and 400 Gy (B) as a function of LHR time. Means ± S.E.M. of at least three independent experiments are presented. (*) indicates a statistically significant difference compared to t = 0 LHR (p < 0.05).
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Figure 5. Repair of γ-ray induced oxypurine clusters during LHR. Repaired Fpg clusters per Mbp at 160 Gy (A) and 400 Gy (B) as a function of LHR time. Insert: Linear regression is plotted as a solid line, and the 95% confidence interval is shown as dashed lines. Means ± S.E.M. of at least three independent experiments are presented. Where not shown, the error bars are of the size of the symbol. (*) indicates a statistically significant difference compared to t = 0LHR (p < 0.05).
Figure 5. Repair of γ-ray induced oxypurine clusters during LHR. Repaired Fpg clusters per Mbp at 160 Gy (A) and 400 Gy (B) as a function of LHR time. Insert: Linear regression is plotted as a solid line, and the 95% confidence interval is shown as dashed lines. Means ± S.E.M. of at least three independent experiments are presented. Where not shown, the error bars are of the size of the symbol. (*) indicates a statistically significant difference compared to t = 0LHR (p < 0.05).
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Figure 6. Schematic diagram highlighting the mechanistic framework involved in IR-induced clustered DNA damage processing.
Figure 6. Schematic diagram highlighting the mechanistic framework involved in IR-induced clustered DNA damage processing.
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Sánchez, A.G.; Keszenman, D.J. Time-Resolved Repair of Clustered DNA Damage in γ-Irradiated Yeast Cells. DNA 2026, 6, 17. https://doi.org/10.3390/dna6020017

AMA Style

Sánchez AG, Keszenman DJ. Time-Resolved Repair of Clustered DNA Damage in γ-Irradiated Yeast Cells. DNA. 2026; 6(2):17. https://doi.org/10.3390/dna6020017

Chicago/Turabian Style

Sánchez, Ana G., and Deborah J. Keszenman. 2026. "Time-Resolved Repair of Clustered DNA Damage in γ-Irradiated Yeast Cells" DNA 6, no. 2: 17. https://doi.org/10.3390/dna6020017

APA Style

Sánchez, A. G., & Keszenman, D. J. (2026). Time-Resolved Repair of Clustered DNA Damage in γ-Irradiated Yeast Cells. DNA, 6(2), 17. https://doi.org/10.3390/dna6020017

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