Replication stress induced exposure to methotrexate in root meristem cells of Vicia faba

Abstract

Replication stress (RS) and oxidative stress (OS) are two main types of endogenous stress (ES) which, by inducing various forms of DNA damage lead to genome destabilization and disruption of cell division control mechanisms. Methotrexate (MTX) is a compound that inhibits dihydrofolate reductase (DHFR), thereby blocking DNA replication and exhibiting antiproliferative effects. The aim of the study was to investigate how 72-h exposure to 0.75 mM MTX on meristematic cells of Vicia faba roots affects the morphology of cell nuclei and mitotic chromosomes, population change of cells in interphase, DNA replication dynamics, cell viability, and hydrogen peroxide (H₂O₂) production. Furthermore, in order to assess epigenetic changes induced by MTX, associated with DNA damage and the replication process, histone H3 acetylation at lysine 56 (H3K56Ac) and histone H4 acetylation at lysine 5 (H4K5Ac) were examined. It was demonstrated that root meristematic cells treated with MTX exhibited abnormal chromosome structure, sustained DNA biosynthesis, and elevated intracellular H₂O₂ levels. Immunocytochemical studies revealed an increased number of fluorescent H3K56Ac foci. It was observed that in the case of H4K5Ac, MTX significantly reduced the frequency of cell populations characterized by euchromatin immunofluorescence and limited the occurrence of heterochromatin-type nuclei. Furthermore, in cells treated with MTX, a significant increase in the number of nuclei with marked nucleoli was observed in addition to the gap 1 (G1) phase. In summary, continuous 3-day exposure to low concentrations of MTX induced a cellular response to reactive oxygen species and DNA replication stress conditions.

Introduction

In eukaryotic cells, accurate DNA replication is crucial for maintaining genome integrity, as it ensures the faithful transmission of genetic information to daughter cells (Saxena and Zou 2022). While the mechanism of DNA biosynthesis is highly accurate, its fidelity is often compromised by exogenous (external) and endogenous (internal) factors, leading to disruption of replication forks (RFs), reduced accuracy of genetic material copying, and DNA breaks (Techer et al. 2017; Zeman and Cimprich 2014). This phenomenon, commonly referred to as replication stress (RS), is a significant source of genome instability in precancerous changes and a characteristic feature of cancer cells (Bester et al. 2011; Hanahan and Weinberg 2011; Burrell et al. 2013; Macheret and Halazonetis 2015). RS can result from many sources, including direct obstacles to RFs progression, such as DNA damage and secondary DNA structures, trinucleotide structures consisting of DNA–RNA hybrids and displaced single-stranded DNA (ssDNA), known as R loops (Aguilera and Garcia-Muse 2012; Helmrich et al. 2013), errors in DNA polymerase function, collisions with transcriptional complexes, and disturbances in cellular nucleotide pools (Zeman and Cimprich 2014). One of the most common causes of DNA replication blockages is DNA damage or adducts (Ashour and Mosammaparast 2021; Ciccia and Elledge 2010), which are often the result of endogenous factors [e.g., reactive oxygen species (ROS), noncanonical nucleotides, or cellular metabolic byproducts] and exogenous factors (e.g., UV radiation and chemical mutagens) (Tubbs and Nussenzweig 2017). DNA damage, such as that caused by UV radiation or alkylating agents, can block DNA replication polymerases. In response to these blockages, the cell employs a mechanism called translesion synthesis (TLS), forcing RFs to bypass the damaged sites (Lehmann et al. 2007). Damage resulting from the attachment of reactive aldehydes or DNA-crosslinking agents (e.g., cisplatin and mitomycin C) to DNA, DNA–protein crosslinks (DPCs), and interstrand crosslinks (ICLs) also pose significant barriers to moving RFs (Ide et al. 2011; Vare et al. 2012; Voulgaridou et al. 2011). Furthermore, the encounter of RFs with single-strand DNA breaks (SSBs) generated by ionizing radiation, free radicals or topoisomerase I causes RFs to “collapse” with the formation of a single-ended double-strand break (seDSB) (Nielsen et al. 2009; Strumberg et al. 2000; Vrtis et al. 2021).

RS induces three main responses: activation of the replication checkpoint, remodeling of blocked RFs, and engagement of DNA repair or tolerance pathways (Saxena and Zou 2022). RS can disrupt the coordinated work of replication helicase and DNA polymerases in RFs, which can lead to increased exposure of single-stranded DNA and accumulation of replication protein A (RPA), as well as activation of the checkpoint kinase ataxia telangiectasia and Rad3-related (ATR) (Zou and Elledge 2003; Byun et al. 2005; Saxena and Zou 2022). Various types of damage and stress activate ATR, including stress caused by UV radiation, the action of DNA polymerase inhibitors, topoisomerase poisons, or stress induced by depletion of the deoxyribonucleoside triphosphate (dNTP) pool (Saldivar et al. 2017). ATR activation leads to the stabilization of stalled RFs, protects them from collapse, prevents premature activation of subsequent replication origins, and initiates cell cycle arrest, thereby mitigating the harmful effects of RS (Saldivar et al. 2017). Furthermore, the conversion of RFs in response to stress is a key cellular mechanism that regulates DNA replication. This process, known as fork reversal, is a crucial protective mechanism that allows forks to reverse their course when they encounter DNA damage, and resume DNA synthesis (Neelsen and Lopes 2015; Quinet et al. 2017). Furthermore, when RFs are under stress or arrest, several DNA repair or tolerance pathways are activated. These pathways include repair by homologous recombination (HR), nonhomologous end joining (NHEJ), repair by nucleotide excision (NER), base excision repair (BER), mismatch repair (MMR), DNA synthesis through damage (TLS), template switching (TS), break-induced replication (BIR), and reprimand via primase and DNA-directed polymerase (PrimPol) (Berti et al. 2016; Cortez 2019). Collectively, these coordinated responses in RFs play a crucial role in ensuring timely DNA replication and maintaining genome stability, especially in the face of RS.

Methotrexate (MTX) is a folic acid (FA) analogue that inhibits dihydrofolate reductase (DHFR), thereby reducing the availability of tetrahydrofolate (Verberne et al. 2019; Hyoun et al. 2012; Fig. 1). Since tetrahydrofolate is an important cofactor in thymidylate synthesis and de novo purine synthesis, MTX inhibits DNA replication and exhibits antiproliferative effects (Hyoun et al. 2012). Using root apical meristems of Vicia faba, the experiments conducted enabled, first, to establish links between MTX-induced RS and oxidative stress (OS), which causes DNA damage via ROS. Second, we attempted to identify the extent of cellular responses to RS at the cytophysiological and molecular levels. For this purpose, meristematic cells of field beans were incubated for 72 h in 0.75 mM MTX and analyzed for: morphology of cell nuclei and mitotic chromosomes, population changes of cells in the G1 phase, syntheses (S) and gap 2 (G2) phases, DNA replication dynamics, cell viability (using Evans blue), and intranuclear hydrogen peroxide (H₂O₂) production using 3,3′-diaminobenzidine (DAB) staining. In order to evaluate MTX-induced epigenetic changes involved in responses to RS, histone H3 acetylation at lysine 56 (H3K56Ac) and H4 acetylation of lysine 5 in nucleosomal histone H4 (H4K5Ac) were examined by immunocytochemistry.

Fig. 1.

Mechanism of action of MTX. Methotrexate inhibits dihydrofolate reductase (DHFR), thereby blocking the conversion of dihydrofolate (DHF) to tetrahydrofolate (THF), which is essential for the synthesis of deoxythymidine monophosphate (dTMP) from deoxyuridine monophosphate (dUMP). The lack of dTMP leads to impaired DNA replication

Materials and methods

Plant material

Seeds of field bean (Vicia faba subsp. minor L.) were surface-sterilized with 70% (v/v) ethanol for 3 min and 10% (v/v) bleach with 0.01% (v/v) Triton X-100 (Merck Life Science, Poznan, Poland) for 5 min. After several rinses with sterile water, the plants were sown in Petri dishes on moist tissue paper sheets and cultured at 24 °C. After 96 h, seedlings with roots approximately 1.5 cm long were placed in subsequent dishes containing distilled water (control) and 0.75 mM methotrexate (MTX; Merck Life Science, Poznan, Poland) solution. Incubation was carried out for 72 h in the dark.

Feulgen staining and DNA cytophotometry

Cut bean root meristems were fixed in Carnoy’s mixture (absolute ethanol and glacial acetic acid; 3:1, v/v; 60 min). After thorough washing with ethanol (Merck Life Science, Poznan, Poland), the material was rehydrated (30 min) and hydrolyzed in 4 M HCl (Merck Life Science, Poznan, Poland). After 60 min, the roots were stained with Schiff’s reagent (Merck Life Science, Poznan, Poland). After rinsing with SO₂ (Merck Life Science, Poznan, Poland) and distilled water, the cut meristems were crushed on basal slides (Merck Life Science, Poznan, Poland) in a drop of 45% acetic acid. After freezing the slides on dry ice (15 min), the slides were rinsed with 70% (v/v) ethanol. After thorough drying, the slides were embedded in Canada balsam (Merck Life Science, Poznan, Poland). The total number of analyzed cells was 4000 (from 10 root meristems) for each experimental series. The extinction of nuclei stained by the Feulgen method was measured at 565 nm using a Jenamed 2 microscope (Carl Zeiss, Jena, Germany) equipped with a computer system (Forel, Lodz, Poland) and calibrated in arbitrary units (a.u.). About 8000 nuclei were collected to assess the distribution of DNA content.

Identification of replicating nuclei with 5-ethynyl-2′-deoxyuridine (EdU)

Root meristems from all experimental series were incubated with 10 μM 5-ethynyl-2′-deoxyuridine (EdU; Thermo Fisher Scientific, Warsaw, Poland) under darkened conditions. After 30 min, the dissected meristems were fixed in 4% phosphate buffered saline (PBS)–buffered paraformaldehyde (4 °C; 20 min; pH 7.4; Merck Life Science, Poznan, Poland) and then macerated with 2.5% citrate-buffered pectinase (pH 5.0; 30 min; Merck Life Science, Poznan, Poland). Meristems were crushed on Polysine^TM slides (Merck Life Science, Poznan, Poland). Replicating nuclei were visualized using the Click-iT DNA Alexa Fluor^® 555 Imaging Kit (Thermo Fisher Scientific, Warsaw, Poland), according to the supplier’s instructions. In the final step of the procedure, DNA was stained for 10 min with 15 μM 4′,6-diamidino-2-phenylindole (DAPI; Merck Life Science, Poznan, Poland). The slides were mounted in a mixture of PBS/glycerol/diazabicyclo[2.2.2]octane (DABCO, 2.3%; Merck Life Science, Poznan, Poland).

Cell viability

Cell viability was determined using Evans blue assay according to Chen et al. (2021) with some modification. Roots of bean were incubated in 0.25% Evans blue (Merck Life Science, Poznan, Poland) for 15 min and rinsed twice in distilled water. Next, in order to better visualize the Evans blue-stained nuclei, nuclear isolation was performed according to the procedure described by Żabka et al. (2024). To achieve a positive control, roots were treated for 1 h in 10% H₂O₂ solution before Evans blue treatment.

Detection of H₂O₂ using DAB

Hydrogen peroxide (H₂O₂) was detected with 3,3′-diaminobenzidine tetrachloride (DAB) reagent (Merck Life Science, Poznan, Poland) as described (Thordal–Christensen et al. 1997). Control seedlings and seedlings previously incubated in MTX were immersed for 3 h in 1 mg mL⁻¹ DAB–HCl in PBS buffer (pH 7.4). The reaction was stopped by transferring the seedlings to distilled water. After staining, the cut meristems were fixed for 45 min in 4% paraformaldehyde buffered with PBS and then macerated in a 2.5% pectinase (Merck Life Science, Poznan, Poland) solution buffered with citric acid (pH 5.0; 37 °C, 40 min; Merck Life Science, Poznan, Poland). The meristems were crushed on glass slides (Merck Life Science, Poznan, Poland) and then fixed in a mixture of glycerol and PBS (9:1; v/v). The presence of H₂O₂ was visualized as brown staining. The average intensity of DAB staining was measured using ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Immunocytochemical detection of histone H3 acetylation on lysine 56 (H3K56Ac) and H4 acetylation of lysine 5 in nucleosomal histone H4 (H4K5Ac)

The cut meristems of the control roots and those treated with MTX were fixed for 40 min (4 °C) in in PBS-buffered 4% paraformaldehyde. Cell nuclei were then isolated by crushing the root meristems between two basal slides according to Żabka et al. (2021). Following pretreatment with PBS-buffered 8% bovine serum albumin (BSA) and 0.1% Triton X-100 (50 min), slides were incubated with rabbit polyclonal antiH4K5Ac antibodies (Merck Life Science, Poznan, Poland) and rabbit monoclonal antihistone H3K56Ac antibodies (Abcam, Cambridgeshire, UK) dissolved in PBS containing 1% BSA (Merck Life Science, Poznan, Poland) at a dilution of 1:500. The specificity of the antibody has been confirmed by validation data provided by the manufacturer for the specified applications, as described in the relevant product data sheets. After an overnight incubation in a humidified atmosphere (4 °C), slides were washed with PBS and incubated for 2 h (20 °C) with secondary goat anti-rabbit immunoglobulin (IgG) conjugated to Alexa Fluor^® 488 antibody in PBS (1:500; Cell Signaling, Leiden, The Netherlands). Antibody dilutions were optimized, and negative controls (without primary antibody) were performed. Cell nuclei were counterstained with DAPI (Merck Life Science, Poznan, Poland). The slides were mounted in a mixture of PBS/glycerol/DABCO (2.3% diazabicyclo[2.2.2]octane; Merck Life Science, Poznan, Poland).

Observations and microscope analyses

Observations of cells stained with EdU and with antibodies against histone modifications H3K56Ac and H4K5Ac were made under a Nikon Eclipse E600W fluorescence microscope (Nikon, Tokyo, Japan) equipped with blue light (B2) filter (λ = 465–496 nm) for Alexa Fluor^® 488, green light (G2 filter) (λ = 540/25 nm) for Alexa Fluor^® 555 and UVB light (U2) filter (λ = 340–380 nm) for DAPI. In addition, images of fluorescent H4K5Ac signals within the nucleolus were acquired using a 0.3 μm step size under oil immersion using Leica Laser Scanning Confocal Microscopy (LSCM) SP8 platform (Leica Microsystems, Wetzlar, Germany) using 100 × /1.40 OIL lens, with the use of Laser Line Supercontinuum Visible (488 nm) and UV (405 nm) Diode laser and visualized using Leica LAS X software ver. 2.0.2.15022 (Leica Microsystems, Wetzlar, Germany).

In order to distinguish between cells in different phases of the cell cycle (G1, early, mid and late S phases, and G2), cytometric measurement of nuclear DNA content was employed. The analysis was performed by combining microfluorometric assessment of DNA content in DAPI-stained cell nuclei with visual analysis of the modifications: H3K56Ac and H4K5Ac.

All images were recorded at the same time of integration using a DS-Fi1 CCD camera (Nikon, Tokyo, Japan). Feulgen-stained cell nuclei DAB-stained cells, and nuclei stained with Evans blue were photographed using a Nikon Eclipse E600W microscope (Nikon, Tokyo, Japan). Quantitative analyses and nuclear DNA fluorescence measurements were made after converting color images into grayscale and expressed in arbitrary units as mean pixel value (pv) spanning the range from 0 (dark) to 255 (white).

In order to obtain the percentage of cells with aberrations (aberration index) after 72 h of incubation in MTX, 100 cells (stained using the Feulgen method) from each of the four root tips were analyzed. Three biological replicates were performed.

Nuclear DNA content was evaluated by means of microdensitometry using a Jenamed 2 microscope (Carl Zeiss, Jena, Germany) with the computer-aided Cytophotometer v1.2 (Forel, Lodz, Poland) for image analysis.

The ImageJ software ver. 1.54p (National Institutes of Health, Bethesda, MD, USA) program was used to analyze intensities of DAB staining. For each experimental series, the coloration intensity of 100 cells was measured.

All fluorescence assays were repeated at least three times, and the data set contained 100 cells per biological replicate.

Statistical analyses

The student’s t–test was used to determine the difference between the averages of two independent groups. The data obtained from all experiments were expressed as mean values ± standard deviation of the mean (± SD).

Results

DNA replication stress leads to chromosome aberrations

Current experiments show that continuous 72-h incubation of V. faba root meristems with 0.75 mM MTX caused approximately 31% of mitotic cells to exhibit changes in chromosome architecture (Fig. 2M). Compared to control seedlings (Fig. 2A–D), treatment of plants with MTX caused some mitotic cells to exhibit breaks in chromatin continuity (Fig. 2E, I), chromosome tangling (Fig. 2F), or abnormal formation of the metaphase plate in the equatorial plane of the cell (Fig. 2J). In the anaphase stage, lost chromosome fragments (Fig. 2G) and chromatin tangles (Fig. 2K) were observed. In the final mitotic stage, a few fragmented chromosomes (Fig. 2H) and telophase bridges (Fig. 2L) were revealed.

Fig. 2.

Feulgen DNA staining of V. faba root meristems incubated in water in a prophase (A), metaphase (B), anaphase (C), telophase (D), and following 72-h treatment with 0.75 mM MTX: prophase (E–I), metaphase (F, J), anaphase (G, K), and telophase (H, L). Scale bars = 10 μm. Mitotic indices (% ± SD; light blue diagrams) and percentage of aberrant mitotic (M) phase cells (% ± SD; dark blue diagrams) in root meristem cells from control and after 72-h incubation with MTX

Compared with the control, which had more condensed nuclei (Fig. 3A), in root meristems treated with MTX, some nuclei underwent enlargement, accompanied by a significant enlargement of the nuclear regions and the appearance of distinct and numerous chromocenters (shown by arrows; Fig. 3B–D). In addition, 3 days of incubation in MTX contributed to the appearance of micronuclei of various sizes (Fig. 3E, F).

Fig. 3.

Feulgen DNA staining of V. faba cell nuclei from the control (A), and MTX-treated seedlings (B–D). Post-telophase cell nuclei with micronuclei formed after treatment with MTX (E, F). Scale bar = 10 μm

Cytophotometric measurements of DNA content

Cytofluorometric measurements of Feulgen-stained nuclear DNA content revealed the presence of 2C, 2–4C, and 4C cells in both the control series (Fig. 4A; green histogram) and MTX-treated roots (Fig. 4B; blue histogram). Compared with the control, MTX treatment altered the proportions of cells in the G1, S, and G2 phases. Continuous incubation in MTX resulted in an approximately 10% increase in the G1 cell subpopulation, while a slight decrease in the number of G2 cells in MTX-treated roots was accompanied by a slight decrease in the number of S phase cells (Fig. 4B).

Fig. 4.

The effect of prolonged incubation with 0.75 mM MTX on cell populations in root meristems of V. faba. Frequency distributions (%) for nuclear DNA contents (Feulgen-staining; arbitrary units, a.u.) in the control (A) and following 72-h treatment with MTX (B). The table below the histograms (D) shows estimated frequencies (%) of G1 + 1/2 M phase cell nuclei (including of ana- and telophases), S phase cell nuclei, and G2 + 1/2 M phase cell nuclei (including pro- and metaphases) in the control and MTX-treated root meristems

Changes in DNA replication after treatment with MTX

The effect of 72-h treatment with 0.75 mM MTX on DNA replication was revealed by combining microfluorimetric quantification of DNA content in DAPI-stained nuclei (Fig. 5A′–C′) with EdU labeling analysis (Fig. 5A–C). The number of cells in the early, middle, and late S phases was calculated as the percentage of cell nuclei exhibiting weak (spotty, characteristic of euchromatin), strong (nearly homogeneous, characteristic of euchromatin), and punctate (characteristic of heterochromatin) fluorescent staining patterns, respectively. Microscopic analyses showed that the highest percentage of untreated cells was observed in the early S phase (approximately 48%), slightly lower in the middle phase (approximately 37%), and lowest in the late S phase (approximately 14%; Fig. 5D). After MTX treatment, changes in the proportions of cells in each phase were observed. Compared with the control series, the highest decrease in the number of nuclei with the weakest fluorescence was accompanied by an increase in the number of cells in the middle and late S phases (Fig. 5D).

Fig. 5.

DNA replication in cell populations evidenced by EdU incorporation. Control cell population: early S phase (A), middle S phase (B), and late S phase (C); cell population stained with DAPI corresponds with the population of EDU stained cells (A′–C′). Scale bar = 10 µm. Fractions [(%) ± SD] of early, mid, and late S phase cells, calculated by combining microfluorimetric quantitation of DNA contents in DAPI-stained cell nuclei and visual analysis of EdU fluorescence patterns. Statistically significant change in MI values is marked by asterisks: ** indicates p < 0.01 and *** indicates p < 0.001. When compared with the control, statistically significant changes in mean values (± SD) are marked by asterisks: **p < 0.01, *** indicates p < 0.001. Each mean value is based on the analysis of ten root meristems (N = 100 cells/meristem)

Induction of hydrogen peroxide

Microcolorimetric assessment of hydrogen peroxide (H₂O₂) levels was also used to evaluate the harmful effects of MTX (Fig. 6). H₂O₂, a type of ROS, was detected in the control (Fig. 6A) and mock buffer (Fig. 6B) as well as in MTX-treated roots (Fig. 6C). The Thordal–Christensen et al. (1997) method was used to visualize H₂O₂ using DAB. Microcolorimetric analysis showed that the average intracellular H₂O₂ level was almost three times higher in MTX-treated cells than in the control and mock buffer (Fig. 6D).

Fig. 6.

DAB-stained cells in the control cells (A), after treatment with mock buffer (B), and after 72-h treatment with MTX (C). Scale bar = 10 μm. Microcolorimetric evaluation of the mean DAB staining intensities [arbitrary units (a.u.)] (D). Compared with the control, statistically significant changes at p < 0.001 are marked by asterisks. When compared with the control, statistically significant changes in mean values (± SD) are marked by asterisks: *** indicates p < 0.001. Each mean value is based on the analysis of ten root meristems (N = 100 cells/meristem)

Identification of dead and living cells

The use of Evans blue revealed blue-stained nuclei (dead cells); live nuclei remained unstained (Fig. 7A). Statistical analyses of the cell viability results are shown in Fig. 7B. After 72 h of exposure to 0.75 mM MTX, the percentage of dead cells increased to approximately 6%, compared with the control series. The positive control consisted of plants incubated in 10% H₂O₂. After 3 h of H₂O₂ treatment, the number of dead cells was found to be 72% (Fig. 7B).

Fig. 7.

Images of the Evans blue stained nuclei of MTX (A; blue nuclei—dead, unstained—alive). Percentage of dead cells (% ± SD) in root meristem cells from control after incubation with MTX and H₂O₂. Scale bar = 10 μm

Acetylation of histone H3 on lysine 56 (H3K56Ac)

Acetylation of histone H3 at lysine 56 (H3K56Ac) is a key epigenetic modification that occurs in response to DNA damage. We observed that in both control and MTX-treated roots, cell nuclei containing 2, 2–4, and 4C DNA were characterized by the presence of fluorescent foci (spots) of H3K56Ac located in the nuclear chromatin region (Fig. 8A–F). The analysis included the mean total number of intranuclear H3K56Ac fluorescent foci during the G1, S, and G2 phases (Fig. 8G). Data calculated for cells in all analyzed phases revealed that 72 h of MTX treatment caused an increase in the average number of H3K56Ac foci; the highest, almost threefold increase in foci was observed in the G1 and G2 phases (Fig. 8G). Additionally, interactive 3D surface plots confirmed, in comparison with control cells (Fig. 8A′–C′), a higher number of so-called peaks (corresponding to H3K56Ac foci) in the nuclei of meristematic cells incubated in MTX (Fig. 8D′–F′).

Fig. 8.

Immunofluorescence of H3K56Ac in cell nuclei from: the control (A–C) and MTX-treated root meristems (D–F) in the G1 (A, D), S (B, E), and G2 phases (C, F); scale bar = 10 μm. Below each micrograph are corresponding interactive 3D surface plots (A′–F′). Mean number of H3K56Ac foci in G1, S, and G2 phase nuclei in the control and MTX-treated root meristem cells (G). Error bars represent standard deviation (SD). Statistical significance between mean values (at ***, p < 0.001, indicated by a black asterisks) was assessed with student’s t test. When compared with the control, statistically significant changes in mean values (± SD) are marked by asterisks: *** indicates p < 0.001. Each mean value is based on the analysis of ten root meristems (N = 100 cells/meristem)

Microscopic images also revealed that the nucleoli were often surrounded by a ring or wreath of fluorescent spots (Fig. 9). Compared with control cells (Fig. 9A, B), where there were fewer fluorescent foci in the perinucleolar chromatin (average number approximately 7), MTX-treated cells revealed a higher number of dots around the nucleolus (approximately 14; Fig. 9C, D).

Fig. 9.

Immunofluorescence of H3K56Ac in cell nuclei from: the control (A, B) and after treatment with MTX (C, D). Nuclei stained with DAPI (A′–D′) and merged images (A″–G″); fluorescent foci around the nuclei marked with yellow circles. Scale bar = 10 μm

Acetylation of histone H4 on lysine 5 (H4K5Ac)

Acetylation of lysine 5 in histone H4 (H4K5Ac) is primarily associated with DNA replication (Jasencakova et al. 2000, 2001). Immunodetection of H4K5Ac in V. faba interphase nuclei revealed three types of cell nucleus labeling patterns: (1) euchromatic type (Fig. 10A, B), associated with euchromatin labeling, (2) heterochromatic type (Fig. 10C, D), associated with strongly labeled heterochromatin domains, sometimes characterized by a typical Rabla configuration, and (3) nucleolar type, showing the strongest, heterogeneous signals within the nucleolus (Fig. 10E–I; Fig. 11A, B, A″, B″, C–E). Our microscopic analyses revealed that the variable relationships between H4K5 acetylation and successive stages of the cell cycle (Fig. 12) can be strongly modified by MTX-induced stress conditions. This manifested itself in a change in the proportion of cell nuclei characterized by fluorescence of either decondensed chromatin, condensed chromatin and/or nucleolar domains.

Fig. 10.

Different labeling patterns observed in meristematic cell nuclei in the control (A, C, E, G) and after MTX treatment (B, D, F, H, I) after immunostaining with antibodies against K5-acetylated H4 histones: nucleolar staining in cell nuclei of euchromatic type (A, B), cell nuclei with immunostained heterochromatin areas (C, D), and different labeling intensities of nucleoli = different nucleolar staining (E–I). Nuclei stained with DAPI (A′–I′) and merged images (A″–I″). Scale bar = 10 μm

Fig. 11.

Confocal immunolocalization of H4K5Ac (green) in isolated cell nuclei counterstained with DAPI (blue) from control (A, A′, A″) and MTX-treated onion seedling (B, B′, B″, C, D, E). Overlaying the images (green and blue) and 3D confocal analysis indicating that, both in the controls (A, A′, A″), and after MTX-treatment (B, B′, B″, C, D, E), that the fluorescent signals are located within the nucleolus. Scale bar = 10 μm

Fig. 12.

Frequencies (% ± SD) of nuclei with H4K5Ac immunolabeling localized in: euchromatic regions (A), heterochromatic regions (B), and nucleolar regions (C), discriminated with respect to successive stages of the cell cycle (G1 phase, early S phase, mid S phase, late S phase, and G2 phase) in root meristems; control and MTX-treated seedlings. Compared with the control, statistically significant changes at p < 0.001, p < 0.01 and p < 0.05 are marked by asterisks. When compared with the control, statistically significant changes in mean values (± SD) are marked by asterisks: ** p < 0.01, *** indicates p < 0.001. Each mean value is based on the analysis of ten root meristems (N = 100 cells/meristem)

The data presented in Fig. 12A revealed that the early stages of the cell cycle, including cells in mid S phase, in root meristems not treated with MTX were mainly represented by cells with nuclei with stained, euchromatic areas of nucleoplasm. It was observed that MTX caused a significant reduction (especially in the G1 phase and mid S phase) in the frequency of cell populations characterized by immunofluorescence of decondensed chromatin. In cells with the second type of chromatin staining, an increased population of control cell nuclei with strong heterochromatin immunofluorescence occurred mainly in the mid and late S phase, and also in the G2 phase (Fig. 12B). The analyses showed that MTX almost completely reduced the occurrence of heterochromatin-type nuclei; approximately 1% of cells were observed in mid S phase (Fig. 10B).

Interesting results were obtained for cells with fluorescently stained nuclear domains (Fig. 12C). In the control meristematic cell population, especially at the beginning and end of interphase, the level of stained nucleoli fluctuated at around 1.5%. By contrast, in cells treated with MTX, in addition to the G1 phase, a significant increase in the number of nuclei with labelled nucleoli was observed, reaching approximately 15% in the late S phase and G2 phase (Fig. 12C).

In mitotic cells, strong immunofluorescence appears to be localized in nuclear organizing regions (NORs) (Fig. 13). Considering the number of visible fluorescent spots, two in prophase and metaphase (Fig. 13A–C) and four in anaphase (Fig. 13D), our earlier experiments have demonstrated (Żabka et al. 2021) that they topologically correspond to brown chromosomal NOR regions revealed by the silver staining procedure (AgNOR).

Fig. 13.

Immunofluorescence labeling with antibodies against K5-acetylated H4 histones of the chromosomal areas: prophase (A, B), metaphase (C), and anaphase (D) in the control cells. Nuclei stained with DAPI (A′–D′) and merged images (A″–D″); fluorescent spots corresponding to NOR regions are marked with red arrows. Scale bar = 10 μm

Discussion

DNA repair mechanisms are tightly regulated in time and space, which is crucial for maintaining genome integrity. The cell coordinates this process using a complex arsenal of enzymatic tools capable of remodeling and repairing DNA, which adapt their response to the type of damage and the phase of the cell cycle. Eukaryotic cells protect their genetic material through a complex signaling pathway called the DNA damage response (DDR), which detects DNA breaks and responds to replication stress (RS), then initiates a response to protect the cell and mitigate the threat to the organism (Harper and Elledge 2007; Jackson and Bartek 2009; Ciccia and Elledge 2010). The final outcome of the DDR depends on the number and type of damages, as well as the effectiveness of repair processes, and can lead to either complete DNA repair, cell cycle arrest, or apoptosis (Adachi et al. 2011; Nisa et al. 2019).

Research by Nihal et al. (2014) showed that, in melanoma cell lines, the effects of a 72-h treatment with 1 μM MTX were associated with increased apoptosis and the activation of caspase–8 and caspase–9. Melanoma cells exhibited growth inhibition and reduced cell viability (Nihal et al. 2014). In our studies, after 72 h of exposure to 0.75 mM MTX, the percentage of dead cells was approximately 6%. It was found that V. faba cells exhibited greater tolerance to MTX-induced oxidative stress, characterized by less damage to cell membranes and higher viability.

Our current experiments on V. faba root meristem cells subjected to prolonged exposure to 750 µM MTX focused mainly on modifications related to the DNA replication process. These included: histone H3 acetylation at lysine 56 (H3K56Ac) and acetylation of lysine 5 in nucleosomal histone H4 (H4K5Ac). The first of these modifications, H3K56Ac, is associated with newly synthesized histones and is involved in DNA biosynthesis (Masumoto et al. 2005; Zhou et al. 2006; Han et al. 2007). It has been observed that the elimination of regulator of Ty1 transposition 109 (Rtt109; histone acetyltransferase) or histone H3 mutations reducing the level of H3K56Ac lead to chromosome instability, manifested by spontaneous breaks mainly during the S, G2, and M phases (Driscoll et al. 2007; Han et al. 2007). The Rtt109 mutation causes synthetic lethality in combination with mutations in several key proteins involved in DNA replication, such as Proliferating Cell Nuclear Antigen (PCNA), DNA polymerase α and Cell Division Cycle 45 (Cdc45) (Han et al. 2007). Furthermore, research by Wurtele et al. (2012) shows that the lack of H3K56 acetylation causes a significant delay in the completion of replication after exposing cells to genotoxic agents such as methyl methanesulfonate (MMS) and camptothecin (CPT), and that the presence of H3K56ac in newly formed chromatin is essential for the proper completion of DNA damage repair during replication. Our studies have shown that in V. faba meristematic root cells, the average number of fluorescent intranuclear foci of histone H3K56Ac was low in all analyzed phases of the cell cycle (G1, S, and G2), while MTX-treated cells showed a significantly increased average number of fluorescent foci. This result, consistent with earlier observations obtained after exposure to hydroxyurea (HU) (Żabka et al. 2024), suggests that this phenomenon most likely correlates with the appearance of DNA damage. Therefore, the results obtained confirm the concept presented by Masumoto et al. (2005), according to which DNA damage initiates an increase in H3K56Ac levels through the activation of checkpoint proteins that play a key role in the repair of genetic material. Histone H3K56 acetylation thus creates a chromatin structure that is conducive to repair processes, making it a key element in the cellular response to DNA damage.

Our analyses indicate that the distribution of H4K5Ac in untreated nuclei of V. faba root meristematic cells depends on the cell cycle phase and on the intranuclear location, including euchromatin, heterochromatin, and nucleoli. Furthermore, the temporal dynamics of epigenetic marker spread from early G1 to late G2 suggest that H4K5 acetylation plays an important regulatory role not only in DNA replication but also in transcription regulation (Jasencakova et al. 2000; 2001). Our earlier work on cadmium stress on meristematic cells of field beans revealed that in the presence of Cd, epigenetic markers were eroded in the S phase (during DNA synthesis) and in the G1 and G2 phases of the cell cycle (when intensive transcription occurs after the M and S phases), and the process of H4K5 acetylation in nucleoli was much less susceptible to metal stress than eu- and heterochromatin areas in the nucleoplasm (Żabka et al. 2021). We obtained similar results in our current study using low concentrations of MTX. We observed that H4K5 acetylation in nucleoli was significantly less sensitive to RS than modifications in eu- and heterochromatin domains. As indicated by Jasencakova et al. (2000), the results obtained can be at least partially explained by the absence of nucleosomes in the nucleolus. The absence of nucleosomes in the nucleolus promotes greater DNA accessibility to transcription factors and regulatory complexes, which may affect both transcription dynamics and the observed epigenetic properties. Due to the fact that histone acetylation is stably maintained during mitosis, daughter cells inherit the epigenetic pattern of H4K5 acetylation present on parental chromosomes before division. Histone acetylation may therefore function as a mechanism for preserving and transmitting "cellular memory," defined as the epigenetic maintenance of specific gene expression states (Jeppesen 1997).

Conclusions

Maintaining genome integrity requires proper DNA replication and even distribution of replicated genetic material during mitosis. Any disruption in these processes can lead to various genetic abnormalities. Cells are constantly exposed to various types of stress that can damage DNA. These stresses can be exogenous or endogenous. Endogenous stress is particularly important because it acts continuously throughout the life of the cell and is the main source of spontaneous genome damage. Among endogenous factors, two play a key role: replication stress, which results in delays or stoppages of replication forks, increasing the risk of double-strand breaks and other damage, and oxidative stress, which often cooccurs with replication stress because damaged or modified DNA hinders replication forks. Our experiments to date therefore open up new research opportunities for future studies on the mechanisms by which stress conditions affect DNA replication dynamics and the cell division cycle.

Author contributions

A.Ż. planned and carried out the experiments and prepared the final version of the article, M.W., K.C. and J.T. P. equally contributed to acquisition of the results, N.G.S. performed the statistical analyses. All authors have read and agreed to the published version of the manuscript.

Data availability

No datasets were generated or analyzed during the current study.

Declarations

Conflicts of interest

The authors declare no competing interests.

Consent for publication

Not applicable.