Next Article in Journal
Activation of the G Protein-Coupled Bile Acid Receptor TGR5 Modulates the HCP5/miR-139-5p/DDIT4 Axis to Antagonize Cervical Cancer Progression
Previous Article in Journal
Antifungal Synergy: Mechanistic Insights into the R-1-R Peptide and Bidens pilosa Extract as Potent Therapeutics against Candida spp. through Proteomics
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
IJMS
Volume 25
Issue 16
10.3390/ijms25168940
ijms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Arabidopsis Histone Variant H2A.X Functions in the DNA Damage-Coupling Abscisic Acid Signaling Pathway

by
Peng Guo
1,
Tian-Jing Wang
1,
Shuang Wang
1,
Xiaoyuan Peng
1,
Dae Heon Kim
2,* and
Yutong Liu
1,*
1
Key Laboratory of Molecular Epigenetics of the Ministry of Education (MOE), Northeast Normal University, Changchun 130024, China
2
Department of Biomedical Science, Sunchon National University, Suncheon 57922, Republic of Korea
*
Authors to whom correspondence should be addressed.
Int. J. Mol. Sci. 2024, 25(16), 8940; https://doi.org/10.3390/ijms25168940
Submission received: 27 June 2024 / Revised: 14 August 2024 / Accepted: 15 August 2024 / Published: 16 August 2024
(This article belongs to the Section Molecular Plant Sciences)
Browse Figures
Versions Notes

Abstract

:
Environmental variations initiate chromatin modifications, leading to the exchange of histone subunits or the repositioning of nucleosomes. The phosphorylated histone variant H2A.X (γH2A.X) is recognized for the formation of foci that serve as established markers of DNA double-strand breaks (DSBs). Nevertheless, the precise roles of H2A.X in the cellular response to genotoxic stress and the impact of the plant hormone abscisic acid (ABA) remain incompletely understood. In this investigation, we implemented CRISPR/Cas9 technology to produce loss-of-function mutants of AtHTA3 and AtHTA5 in Arabidopsis. The phenotypes of the athta3 and athta5 single mutants were nearly identical to those of the wild-type Col-0. Nevertheless, the athta3 athta5 double mutants exhibited aberrant embryonic development, increased sensitivity to DNA damage, and higher sensitivity to ABA. The RT-qPCR analysis indicates that AtHTA3 and AtHTA5 negatively regulate the expression of AtABI3, a fundamental regulator in the ABA signaling pathway. Subsequent investigation demonstrated that AtABI3 participates in the genotoxic stress response by influencing the expression of DNA damage response genes, such as AtBRCA1, AtRAD51, and AtWEE1. Our research offers new insights into the role of H2A.X in the genotoxic and ABA responses of Arabidopsis.

1. Introduction

Abiotic stress, which is the consequence of significant external environmental changes, can cause DNA damage, thereby jeopardizing the integrity and stability of the genome [1,2]. Various components of DNA, such as the purine and pyrimidine bases, sugar residues, and phosphodiester linkages, are susceptible to damage from both environmental and endogenous genotoxic agents [3]. A multitude of signaling pathways that are essential for DNA repair, cell cycle regulation, or cell death are initiated by exposure to genotoxins, DNA-damaging agents [4]. The coordination of repair processes at the loci of DNA damage, which are complex assemblies formed in the vicinity of the damaged site, is of the utmost importance [5,6]. In order to fit within the restricted nuclear space, the genomes of eukaryotic cells are condensed into chromatin fibers. The nucleosome is the fundamental component of chromatin. Each nucleosome is composed of approximately 150 base pairs of DNA that are wrapped around a histone octamer. This octamer is composed of two copies of the core histones H2A, H2B, H3, and H4 [7,8,9]. Distinctive histone variants have evolved in both animals and plants, playing critical roles in chromatin dynamics. The replication-independent incorporation of these variants can profoundly alter chromatin structure, influencing various biological processes such as transcriptional regulation and genome stability [10,11,12,13,14]. In mice, the essential role of histone variants in maintaining genomic stability is highlighted by the association of H2A.X loss with radiation sensitivity and growth retardation, both of which stem from compromised genomic stability [15]. In humans, H2A.X also participates in degradosome formation combined with endonuclease cyclophilin A and the apoptosis-inducing factor (AIF) enzyme to favor chromatinolysis processes under certain redox conditions within the cell [16,17]. The early cellular response to DNA damage is significantly influenced by the rapid phosphorylation of H2A.X at a conserved serine/glutamine (SQ) motif in its carboxyl terminus, which leads to the formation of γH2A.X [18,19,20,21,22,23]. Although the precise function of γH2A.X foci in yeast and mammals is a subject of debate, they are considered markers of DNA double-strand breaks (DSBs). The kinase activity of ataxia telangiectasia mutated (ATM) and Rad-3-related (ATR) kinase is the primary factor determining the rapid formation of γH2A.X foci following the induction of DSBs [24,25]. In Arabidopsis, there are two H2A.X isoforms, AtHTA3 and AtHTA5, which are distinguished by a mere two amino acids in their primary protein structure. Previous research has demonstrated that H2A.X-silenced lines, which employ RNA interference (RNAi) technology and retain residual gene expression, exhibit mild hypersensitivity to genotoxic agents, including bleomycin and camptothecin [26]. T-DNA insertional mutants for H2A.X demonstrate altered patterns of DSB repair, indicating that the H2A.X isoforms play both redundant and divergent roles in the modulation of Non-Homologous End Joining (NHEJ) and Homologous Recombination (HR) repair pathways [27]. It is important to note that the double mutants of h2axa-2 (athta5) and h2axb (athta3) on a T-DNA background exhibited delayed radicle emergence in comparison to the wild-type Col-0. The h2axb mutants did not demonstrate a substantial difference in seed aging sensitivity compared to Col-0, whereas the h2axa-2 mutants exhibited delayed germination [28].
Abscisic acid (ABA) is a critical plant hormone that controls a variety of biological processes, such as seed germination, plant growth, and responses to abiotic stress [29,30]. Studies have demonstrated that elevated levels of ABA inhibit DNA replication and cell division, which leads to growth retardation [31,32]. Major loss-of-function mutants of genes implicated in DSB repair, particularly in the NHEJ and HR pathways, demonstrate increased sensitivity to ABA treatment. An additional investigation of the phosphorylation status of H2A.X subsequent to ABA treatment has demonstrated that ABA induces γH2A.X and initiates a DNA damage response [33]. ABA activates HR-related genes during post-germination periods and stimulates the expression of DSB repair genes via HR during seed germination [33,34]. Abscisic acid insensitive 3 (ABI3), a transcription factor that is specific to plants and is a member of the B3 superfamily, is essential for the ABA signaling pathway and the transition from seed to seedling stages [35,36]. During seed maturation, the abi3 mutants demonstrate severe defects, including an inability to degrade chlorophyll in dried seeds and an intolerance to desiccation [37,38,39]. The Sph/RY element CATGCA in the promoter region enables ABI3 to activate its own expression at the transcriptional level [40]. It is induced by LEAFY COTYLEDON1 (LEC1), LEAFY COTYLEDON 2 (LEC2), and FUSCA3 (FUS3) [41,42,43]. Moreover, the transcriptional regulator DESPIERTO (DEP) and the chromatin-remodeling enzyme PICKLE (PLK) substantially reduce ABI3 expression [44,45].
In this investigation, we effectively generated independent loss-of-function single mutants for AtHTA3 and AtHTA5 by utilizing clustered regularly interspaced short palindromic repeats/CRISPR-associated nuclease 9 (CRISPR/Cas9) technology, as well as a double mutant called athta3 athta5 through crossing. Notably, the phenotypes of the athta3 and athta5 single mutants were indistinguishable from those of the wild-type Col-0. In contrast, the athta3 athta5 double mutants exhibited a significant degree of ABA hypersensitivity, as well as apparent DNA damage and aberrant embryonic development. Our investigation demonstrated that the histone variant genes AtHTA3 and AtHTA5 were situated downstream of the ABA signaling regulatory gene AtABI3. Our results offer new perspectives on the function of AtH2A.X in the DNA damage-coupling ABA signaling pathway by influencing the expression of AtABI3.

2. Results

2.1. Phenotypes of athta3 and athta5 Loss-of-Function Mutants under Different DNA Damage Reagents

To investigate the function of H2A.X in Arabidopsis, we utilized CRISPR/Cas9 technology to generate independent mutants. Single-guide RNA (sgRNA) target sites for AtHTA3 and AtHTA5 were cloned into the CRISPR/Cas9 system driven by the YAO promoter [46]. In the athta3 single mutants, a 1 bp insertion of either T or A at the position of 33 bp downstream of the start codon (ATG) resulted in a premature stop codon (Figure S1A). In the athta5-1 mutants, a 1 bp insertion occurred at the position of 18 bp downstream of the ATG, while in the athta5-2 mutants, a 7 bp deletion coupled with a 2 bp substitution was found at the position of 11 bp downstream of the ATG; both induced a frameshift mutation and consequently a premature stop codon (Figure S1B). We obtained Cas9 gene-free plants by evaluating their resistance to hygromycin (Figure S1C). Subsequently, we crossed the athta3-1 plants with the athta5-1 plants to produce the athta3-1 athta5-1 double mutants. We then established two complementation lines (Com lines) by expressing 2 × FLAG epitope-tagged AtHTA3 or AtHTA5 genomic DNA under the regulation of their native promoters into the athta3-1 athta5-1 double mutant background. The expressions of the constructs AtHTA3pro:gAtHTA3-2 × FLAG and AtHTA5pro:gAtHTA5-2 × FLAG in the Com lines, designated as the athta3-1 athta5-1/Com3 and the athta3-1 athta5-1/Com5, were confirmed using an anti-FLAG antibody (Figure S1D). During the vegetative stage, we observed that the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the two Com lines exhibited no abnormal phenotype and developed similar biomass (also defined as fresh weight) to the wild-type Col-0 (Figure 1A,B). Notably, in the athta3-1 athta5-1 double mutants, we observed that silique sizes were smaller, and these siliques contained approximately 35% aborted seeds, contrasting sharply with the less than 2% seed abortion rate observed in the wild-type Col-0, the athta3 and athta5 single mutants, and the two Com lines (Figure 1C,D). Collectively, these findings suggested that AtH2A.X may have redundant functions in silique development and embryogenesis during the reproductive stages.
To evaluate the function of AtH2A.X in the response to DNA damage, reagents with different mechanisms were used: the DSB inducer Zeocin and Phleomycin (PM), the DNA intra-strand crosslinker mitomycin C (MMC), the DNA synthesis inhibitor hydroxyurea (HU), and the alkylating agent methyl methanesulfonate (MMS) [47]. DNA damage is recognized as triggering cell division arrest in the meristem [48]. Consequently, we measured the relative root length of the wild-type Col-0, the athta3 single mutants, the athta5 single mutants, and the athta3-1 athta5-1 double mutants, as well as the athta3-1 athta5-1/Com3 and the athta3-1 athta5-1/Com5 lines (Figure 2). The atatr and atfen1 mutants, known to be susceptible to DNA damage, were used as positive controls [49,50]. The athta3-1 athta5-1 double mutants showed a much-decreased root development phenotype in the presence of the five chemicals mentioned above compared to the wild-type Col-0 and the Com lines, as shown in Figure 2. By contrast, there were no discernible phenotypes of root development in the athta3 and athta5 single mutants. These results suggested that AtH2A.X plays a major and redundant role in the DNA damage response.
To further substantiate the impact of AtH2A.X loss on the DNA damage response, we evaluated the expression profiles of critical DNA damage response genes in both the Col-0 and mutant lines. Our analysis encompassed genes that are recognized for their involvement in DNA damage repair, including breast cancer susceptibility gene 1 (AtBRCA1) and RAS associated with diabetes protein 51 (AtRAD51), which are genes involved in the HR repair pathway [51,52]. We also examined the expression of TSO MEANING ‘UGLY’ IN CHINESE 2 (AtTSO2), which encodes a subunit of ribonucleotide reductase (RNR). This gene demonstrates transcriptional upregulation to mitigate the depletion of the nucleotide pool as a result of HU treatments [53]. Subsequently, we investigated the expression of WEE1-like kinase (AtWEE1). In response to replication stress induced by HU, AtWEE1 induces cell cycle arrest [6,54]. As shown in Figure S2, we found that the expression levels of these DNA damage response genes were increased in the athta3 athta5 double mutants, with the effect being particularly pronounced after exposure to genotoxic agents. These findings indicated that the loss of AtH2A.X function may reduce the efficacy of DNA damage repair, prolong the duration of cell cycle arrest, and elicit an enhanced DNA damage response.

2.2. Tissue-Specific Gene Expression Patterns and Subcellular Localizations of AtHTA3 and AtHTA5

To investigate the expression patterns of AtHTA3 and AtHTA5, whose N-terminal tails differ by only two amino acids, we generated transgenic plants that expressed the β-glucuronidase (GUS) reporter gene under the control of the native AtHTA3 and AtHTA5 promoters (denoted by AtHTA3pro:GUS and AtHTA5pro:GUS). Various tissues, such as leaves, flowers, stems, lateral roots, root apexes, vascular tissues, guard cells, major veins of the cotyledons, styles, valves, septums, ovule funiculi and abscission zones in mature siliques, and germinated seeds, exhibited largely overlapping expression patterns for the two AtH2A.X isoforms (Figure 3A). Nevertheless, the AtHTA5pro:GUS construct exhibited substantially higher expression levels in roots and flowers than the AtHTA3pro:GUS construct. Furthermore, the stele, columella, and lateral root cap exhibited distinct AtHTA5pro:GUS activity, whereas AtHTA3pro:GUS exhibited only mild activity in these tissues (Figure 3A). Our findings were subsequently verified through reverse transcription-quantitative polymerase chain reaction (RT-qPCR) analysis. The GUS expression patterns were shown to be correlated with the transcript levels of AtHTA3 and AtHTA5, as evidenced by the intensity of GUS staining in a variety of tissues (Figure 3B). To further examine the subcellular localization of AtHTA3 and AtHTA5, we generated fusion proteins by affixing green fluorescence protein (GFP) to the C-termini of AtHTA3 and AtHTA5, respectively. This process resulted in the recombinant proteins AtHTA3-GFP and AtHTA5-GFP. These constructs were subsequently introduced to the Arabidopsis protoplasts together with nuclear localization signal (NLS)-RFP (marker for labeling nucleus). Both AtHTA3-GFP and AtHTA5-GFP were primarily localized to the nucleus, as illustrated in Figure 3C. A minor fraction was also observed in the cytosol. Collectively, these findings suggested that AtHTA3 and AtHTA5 exhibited comparable spatial and temporal expression patterns.

2.3. AtHTA3 and AtHTA5 Function in the ABA Response

Prior research demonstrated that mutants associated with the HR pathway and DSB sensor genes exhibited relatively increased ABA sensitivities [33]. Therefore, we conducted a more thorough investigation of the ABA sensitivity of the athta3 single mutants, the athta5 single mutants, and the athta3 athta5 double mutants in the presence of ABA during the seed germination and seedling growth stages. The phenotypic alterations were not detectable under normal conditions; however, the athta3-1 athta5-1 double mutants exhibited an ABA hypersensitivity phenotype with reduced cotyledon greening rates (percentage of greening seedlings over all germinated seedlings) after germination on day 7 at varying concentrations of ABA (Figure 4A,B). Of note, the phenotypes of the athta3-1 athta5-1/Com3 and the athta3-1 athta5-1/Com5 lines were comparable to those of the athta3 and athta5 single mutants, as well as the wild-type Col-0 (Figure 4A,B). A high concentration of exogenous ABA inhibits root elongation [55,56], and to further investigate whether AtHTA3 and AtHTA5 are involved in the inhibition of root growth by ABA, the wild-type Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the complementation lines were grown in solid medium containing 0 and 5 μM ABA. The primary root growth of the athta3 and athta5 single mutants, the athta3-1 athta5-1 double mutants, the complementary lines and the wild-type Col-0 did not exhibit any phenotypic changes following ABA application during post-germination developmental phases (Figure 4C,D). These findings suggested that AtHTA3 and AtHTA5 function redundantly in response to ABA during the seed germination stage, but they are not engaged in the response to ABA in terms of primary root growth.
To examine the possible role of AtHTA3 and AtHTA5 in response to ABA, we examine the expression of several important marker genes in ABA signaling using RT-qPCR analysis. It is intriguing that the expression levels of Abscisic acid insensitive 1 (AtABI1) and AtABI3 were the only ones that were substantially altered in the athta3-1 athta5-1 double mutants in comparison to the wild-type Col-0 with or without ABA treatment (Figure 5). Previous research indicated that the cell cycle is a critical target for ABI3, and it is conceivable that the abi3 mutation could result in ineffective ABA signaling in the cell cycle, thereby reducing seed quiescence [35]. Therefore, we proposed that the loss of function of AtH2A.X results in an abnormal embryo and an ABA-hypersensitive phenotype by excessively activating the expression of AtABI3 under normal conditions and in response to ABA.

2.4. AtABI3 Functions Genetically Downstream of AtHTA3 and AtHTA5

Since AtHTA3 and AtHTA5 negatively impact the expression of AtABI3, we sought to examine the genetic interaction among AtHTA3, AtHTA5, and AtABI3. In order to produce the athta3-1 atabi3-8 double mutants, the athta5-1 atabi3-8 double mutants, and the athta3-1 athta5-1 atabi3-8 triple mutants, we crossed the athta3-1, the athta5-1, and the athta3-1 athta5-1 with atabi3-8, a loss-of-function mutant line that results in a critical amino acid conversion of leucine 298 to phenylalanine within the B1 domain [57,58]. In our assessment of ABA sensitivity following germination, the double mutants athta3-1 atabi3-8 and athta5-1 atabi3-8, as well as the triple mutants athta3-1 athta5-1 atabi3-8, exhibited a hyposensitivity to ABA that was consistent with the atabi3-8 single mutants during the germination phase (Figure 6A,B). Nevertheless, the athta3-1 atabi3-8 double mutants, the athta5-1 atabi3-8 double mutants, and the athta3-1 athta5-1 atabi3-8 triple mutants did not exhibit phenotypes in terms of primary root growth after ABA treatment during seedling developmental phases (Figure 6C,D). This finding suggested that AtABI3 functions genetically downstream of AtHTA3 and AtHTA5 during the germination stages of seeds that are inhibited by ABA.

2.5. AtABI3 Plays a Role in Genotoxic Stress Response

Given that the cell cycle is a likely target for ABI3, as previously reported [35], and considering that AtABI3 expression is down-regulated by AtHTA3 and AtHTA5, we conducted further research to determine if AtABI3 is implicated in the DNA damage response. The atabi3-8 mutants were subjected to various DNA-damaging agents, including Zeocin, PM, MMC, HU, and MMS, with the atatr and atfen1 single mutants serving as positive controls. As illustrated in Figure 7A–D, the atabi3-8 mutants displayed hypersensitivity to MMC and HU, as evidenced by stunted root growth and diminished leaf size relative to the wild-type Col-0. In contrast, other DNA-damaging agents had no significant impact on the growth of the atabi3-8 mutants. These findings indicated that AtABI3 plays a role in the response to genotoxic stress, particularly in the context of inter-strand crosslinks (ICLs) and replication fork stalling-induced DNA damage. To substantiate the impact of AtABI3 loss on the DNA damage response, we assessed the expression levels of key DNA damage response genes, namely AtBRCA1, AtRAD51, and AtWEE1. The results demonstrated that the absence of AtABI3 function resulted in a notable upregulation of these genes following MMC and HU treatments in comparison to the wild-type Col-0 (Figure 7E,F). This finding highlighted a disrupted DNA damage response. Moreover, we investigated the expression of AtCYCD1;1, which encodes a D-type cyclin that plays a crucial role in cell cycle re-entry during the activation of the meristem, thereby facilitating germination and early seedling growth [54,59]. In the atabi3-8 mutants, AtCYCD1;1 expression demonstrated fluctuating patterns under HU treatment in comparison to the wild-type Col-0 (Figure 7F). This variability may be attributed to a synchronized progression through mitosis induced by HU, which ultimately results in the observed root growth retardation [53].

3. Discussion

The previously published H2A.X-silenced line (miH2AX) retained 15% of H2AXa (AtHTA5) and 52% of H2AXb (AtHTA3) expression levels compared to the wild type. This partial retention of expression is likely the reason why the miH2AX line shows minimal phenotypic abnormalities and only a modest growth delay when exposed to agents that induce DSBs [26]. Notably, the H2AXa;H2AXb double mutants, with T-DNA insertion in the promoter region of H2AXa and intron region of H2AXb, also failed to completely abolish H2A.X. The H2AXa;H2AXb double mutants show no significant effect on fertility, only a marginal effect on root growth, even under conditions of ionizing radiation [60]. Previous studies have implicated H2AXb in both NHEJ and HR DNA repair mechanisms, whereas H2AXa appears to have a more pronounced influence on HR. This differential involvement suggests that the two isoforms may modulate different DSB repair pathways [27]. Therefore, establishing and studying the complete knockout (KO) lines is crucial for thoroughly elucidating the function of AtH2A.X.
In this study, leveraging the capabilities of CRISPR/Cas9 technology, we successfully generated several independent AtHTA3 and AtHTA5 loss-of-function mutants that differ from those in previous reports. Notably, we found that neither the athta3 nor the athta5 single mutants exhibited growth retardation under normal conditions or when treated with genotoxic agents. However, the athta3 athta5 double mutants displayed heightened sensitivity to genotoxic stress induced by Zeocin, PM, MMC, HU, and MMS, as compared to the single mutants, the wild-type control (Col-0), and the complementation lines. The h2axa-2 h2axb double mutants exhibited substantial hypersensitivity to MMS, X-rays, and MMC, as previously noted by Wanda et al. [28]. Of note, they also observed that the spatial and temporal expression patterns of the two isoforms are primarily overlapping. This observation is comparable to ours. Previous studies have shown that Zeocin and PM, members of the bleomycin family, are known to induce DSBs [61]. MMC is recognized for causing inter-strand crosslinks, which are addressed through the γH2A.X-mediated Fanconi anemia (FA) repair pathway [62]. HU leads to replication fork stalling due to the depletion of the deoxyribonucleotide (dNTP) pool, thereby inhibiting DNA replication [63]. MMS is known for causing DNA base damage, which could be repaired through the ADP-ribosylation of H2A.X [64]. According to our findings, AtH2A.X is a critical component of numerous DNA repair pathways in Arabidopsis, and these two isoforms could complement each other’s functions.
We also identified that the ABA-hypersensitive phenotypes of athta3 athta5 double mutants were observed during seed germination, as opposed to under seedling growth conditions. New insights into the interaction between the DNA repair pathway and ABA signaling are provided by the ABA overly sensitive mutant excessively ABA-sensitive mutant abo4-1, where ABO4 encodes DNA pol ε, as evidenced by previous research. In abo4-1, ABA exposure results in an elevation of DSBs, genome instability, and an increase in HR, marked by the upregulation of DSB-responsive genes such as MEIOTIC RECOMBINATION 11 (MRE11) and GAMMA RESPONSE 1 (GR1), and the downregulation of others like RAD51, BRCA1, and KU70 [34]. Arabidopsis DNA replication factor C1 (RFC1), a principal component of the RFC complex, is implicated in somatic HR and is subject to ABA modulation. ABA intervention leads to a decline in HR and the expression of genes pertinent to DNA damage response in rfc1 mutants, underscoring RFC1’s role in ABA-mediated HR [65]. Furthermore, the Arabidopsis loss-of-function mutants for genes involved in the DSB sensing and signaling (atatm-2, atatr, atmre11, atrad50 and atnbs1), NHEJ (atku80, atku70, atpolε-1, atlig4 and atxrcc4), and HR (atrad51, atrad52, atrad54 and atbrca1) pathways have all exhibited ABA hypersensitivity [33]. These results emphasize the interaction between ABA responses and genotoxic stress. Intriguingly, we discovered that the expression of AtABI3 is negatively regulated by AtHTA3 and AtHTA5 in response to ABA. The evidence that AtABI3 expression is significantly induced in athta3 athta5 mutants under ABA treatment corroborates this outcome.
The transition from a germinating embryo to an autotrophic seedling is hampered by dehydration and duress. A critical stress hormone, ABA, regulates the cell developmental checkpoints in plants that are experiencing water deficiency. AtABI3, a critical transcription factor, is responsible for the sustained but reversible growth arrest of germinated embryos in response to stress. As previously reported, the abi3-1 mutant is incapable of executing the cell developmental checkpoints [66]. In our study, we unexpectedly found that another AtABI3 loss-of-function mutant, atabi3-8, showed retarded growth under the treatment of MMC and HU, during seedling growth stages. We also found that, by RT-qPCR analysis, the expression patterns of a number of significant DNA damage response genes, including AtBRCA1, AtRAD51, and AtWEE1, were different from the wild-type Col-0. These data suggested that ABA may affect the genotoxic stress response through AtABI3 and that AtABI3 may be involved in the regulation of these DNA damage-responsive genes.

4. Materials and Methods

4.1. Plant Materials and Growth Conditions

This study utilized Arabidopsis thaliana (L.) Heynh. ecotype Columbia-0 (Col-0) as the model plant system. Seedlings intended for crosses, developmental phenotype observations, and propagation were grown in soil under greenhouse conditions. In contrast, seedlings designated for stress treatments were germinated and cultivated on Murashige and Skoog (MS) medium [67] containing 1% sucrose (adjusted to pH 5.7 with KOH) within growth chambers. The plants were subjected to a 16 h light/8 h dark photoperiod at temperatures of 23 °C during the light phase with 160 μmol m−2 s−1 intensity and 21 °C during the dark phase, with a maintained relative humidity of 60%. The atabi3-8 and atatr (SALK_032841) mutants were sourced from the Nottingham Arabidopsis Stock Centre (NASC), with homozygosity confirmed through PCR-based genotyping employing LB/RP or LP/RP primer sets (see Table S1 for details). The atfen1 mutants were generously provided by Zhizhong Gong [50].

4.2. Plasmid Construction

CRISPR/Cas9 vectors were generated by designing guide targets for AtHTA3 and AtHTA5 using the online tool available at http://www.rgenome.net [68]. Subsequently, the single-guide RNA cassette was cloned into the pYAO:hSpCas9 binary vector using the restriction enzyme SpeI [46]. To generate AtHTA3pro:GUS and AtHTA5pro:GUS constructs, a 2094 bp fragment upstream of the AtHTA3 start codon and a 2200 bp fragment upstream of the AtHTA5 start codon were individually amplified and cloned into the pCAMBIA3301 binary vector using EcoRI and NcoI sites. The cDNA fragments of AtHTA3 and AtHTA5, excluding stop codons, were amplified from the Arabidopsis leaf cDNA library and then individually inserted into the 326-sGFP plasmid using the restriction sites XbaI and BamHI sites. To construct AtHTA3pro:AtHTA3-2 × FLAG and AtHTA5pro:AtHTA5-2 × FLAG complementation plasmids, the promoter region along with genomic sequences lacking stop codons of AtHTA3 and AtHTA5 were inserted into the pCAMBIA1302 binary vector using EcoRI and BamHI sites through the In-Fusion HD Cloning Kit (Takara, 639650, Kusatsu, Japan). All primers used are listed in Table S1.

4.3. Mutant Isolation and Generation of Transgenic Plants

Transgenic Arabidopsis plants were generated using the Agrobacterium tumefaciens-mediated transformation method, employing the floral dip technique in the presence of 0.05% Silwet L-77 (Coolaber, CS9791, Shanghai, China) [69]. Agrobacterium tumefaciens GV3101 strain was cultivated on Luria–Bertani (LB) broth medium containing 50 mg/L rifampicin (Sangon Biotech, A430385, Shanghai, China). For the isolation of CRISPR/Cas9 mutants, T1 progeny seeds were selected on a B5 medium containing 50 mg/L hygromycin (Sangon Biotech, A600230, Shanghai, China). The presence of mutations in AtHTA3 and AtHTA5 was confirmed by Sanger sequencing. Homozygous T3 lines, which no longer contained the Cas9 construct, were chosen for further experimentation. For the construction of athta3 athta5 double mutants by crossing, based on Mendel’s genetic law, F2 populations of plants (an average of 40 per F2 population) were screened for homozygosity or heterozygosity for the mutation of AtHTA3 or AtHTA5 locus genotyped using HTA3-CR-Check and HTA5-CR-Check primer sets, and separated homozygous F3 seeds were used for further experiments. For complementation lines, T1 generation seeds were screened on B5 medium containing 50 mg/L hygromycin, and resistant seedlings were planted in soil, T2 populations with approximately 75% resistant seedlings, indicating single-locus T-DNA insertion, were considered for further analyses with confirmation using anti-FLAG antibody (Sigma, F3165, Setagaya, Japan), and homozygous T3 seeds were used for further experiments. For promoter-GUS transgenic plants, T1 seeds were selected on B5 medium containing 0.2‰ Basta (Sangon Biotech, A614229, Shanghai, China), and resistant seedlings were planted in soil with confirmation using GUS staining. T2 populations with single-locus T-DNA insertion were considered for further analyses.

4.4. DNA Damage Agents and ABA Treatments

For DNA damage sensitivity assays, sterilized seeds were cultured in liquid MS medium with various concentrations of DNA damage agents: 25 μM Zeocin (Invitrogen, R250-01, Waltham, MA, USA); 75 p.p.m. Phleomycin (Sangon Biotech, A620211-0100, Shanghai, China); 10 µM MMC (Selleck, S8146, Houston, TX, USA); 1 mM HU (Sigma, H8627, Burlington, MA, USA); 100 p.p.m. MMS (Sigma, 129925, Burlington, MA, USA). The medium was refreshed every 2 days to ensure chemical efficacy. After 2 weeks, seedlings were transferred to a solid MS medium for measurement and imaging. For ABA sensitivity assays for monitoring cotyledon greening, sterilized seeds were sown in liquid ½ MS medium with 0, 0.25, or 0.5 µM ABA (Coolaber, CA1011, Beijing, China). The percentage of greening was determined after 7 days, with images taken for documentation. To evaluate root growth, 4-day-old seedlings with similar initial root lengths were transplanted to solid ½ MS medium with 0 or 5 µM ABA and allowed to grow vertically. Subsequently, the root lengths were reassessed after a further 2 weeks of growth.

4.5. GUS Staining and Subcellular Localization

For the GUS staining experiment, transgenic plants Arabidopsis harboring either the AtHTA3pro:GUS or AtHTA5pro:GUS constructs were infiltrated with a GUS staining solution (Clontech, 631721, Kusatsu, Japan) and incubated at 37 °C. Following the staining procedure, the plants were subjected to an overnight clearing process. To observe the subcellular localization of AtHTA3 and AtHTA5, mesophyll protoplasts isolated from 3-week-old seedlings were co-transfected with AtHTA3-GFP or AtHTA5-GFP along with NLS-RFP constructs using polyethylene glycol (PEG)-mediated transfection [70]. Once the transfection was complete, the protoplasts (2 × 105 mL−1) were incubated in 1 mL W5 buffer (pH 5.7) containing 2 mM MES, 5 mM KCl, 154 mM NaCl, and 125 mM CaCl2 at 22 °C for 12 to 16 h in a growth chamber and then visualized using the fluorescence microscope (Olympus, BX53, Tokyo, Japan).

4.6. RT-qPCR Analysis

Briefly, total RNA was isolated using TRIzol reagent (Invitrogen, 15596026CN, Waltham, MA, USA), and RNA quality was assessed by detecting the A260/A280 absorption ratio using a spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA), followed by formaldehyde agarose gel electrophoresis to evaluate RNA integrity. cDNA was synthesized from 2 μg total RNA using the One-Step gDNA Removal and cDNA Synthesis SuperMix Kit (TransGen Biotech, AT311-03, Beijing, China). Each reaction included 25 ng cDNA with PowerUp SYBR Green Master Mix (Thermo Fisher Scientific, A25742). The length of the qPCR products is 80-200 bp. The cycling conditions were set in the QuantStudio™ 5 Real-Time PCR System (Applied Biosystems, Carlsbad, CA, USA) according to the manual, with an initial hold stage at 50 °C 2 min and 95 °C 2 min, followed by 95 °C 15 sec for denaturation and 60 °C 1 min for annealing/extension with 40 cycles in the PCR stage. Melt curve analysis was performed to ensure specific amplification. QuantStudio™ Design and Analysis Software v1.2.x (Applied Biosystems, Carlsbad, CA, USA) was used to acquire raw qPCR data. The 2−ΔΔCt method was used for data analysis [71], and the expression stability of GAPDH as an internal control was assessed using BestKeeper software (https://www.gene-quantification.de/bestkeeper.html) [72]. All reactions were performed using samples from three independent biological replicates. All primers used for RT-qPCR are listed in Table S1.

4.7. Statistical Analysis

All experimental data were expressed as the means ± standard errors (SD). Normality of the data was assessed using the Shapiro–Wilk normality test with R v.4.0.1 software. One- or two-way ANOVA statistical analyses were performed using GraphPad Prism v.9.0 software.

5. Conclusions

In conclusion, our investigation demonstrates that the histone variants AtHTA3 and AtHTA5 play crucial roles in DNA damage repair and ABA-mediated seed germination, possibly by regulating the ABA signaling gene AtABI3. This study lays a foundation for further research on the role of AtH2A.X in the DNA damage-coupling ABA signaling pathway by influencing the expression of AtABI3 in Arabidopsis. Further investigations are necessary to ascertain if AtABI3 can directly interact with the signaling molecules involved in DNA damage response.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/ijms25168940/s1.

Author Contributions

Y.L. and D.H.K. conceptualized and supervised the research. P.G. performed the physiological analyses of plants. T.-J.W. performed molecular cloning. S.W. performed RT-qPCR. X.P. generated the mutant lines. Y.L. wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This study was supported by the Natural Science Foundation of Jilin Province (20240101274JC) and the National Research Foundation of Korea (NRF) grant funded by the Korean government (MSIT, 2023R1A2C1007284).

Institutional Review Board Statement

Not applicable.

Data Availability Statement

All data and materials presented in this paper are available on request from the corresponding author.

Acknowledgments

We would like to thank Zhizhong Gong for kindly providing atfen1 mutants.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Waterworth, W.M.; Drury, G.E.; Bray, C.M.; West, C.E. Repairing breaks in the plant genome: The importance of keeping it together. New Phytol. 2011, 192, 805–822. [Google Scholar] [CrossRef] [PubMed]
  2. Balestrazzi, A.; Confalonieri, M.; Macovei, A.; Donà, M.; Carbonera, D. Genotoxic stress and DNA repair in plants: Emerging functions and tools for improving crop productivity. Plant Cell Rep. 2011, 30, 287–295. [Google Scholar] [CrossRef] [PubMed]
  3. Bray, C.M.; West, C.E. DNA repair mechanisms in plants: Crucial sensors and effectors for the maintenance of genome integrity. New Phytol. 2005, 168, 511–528. [Google Scholar] [CrossRef] [PubMed]
  4. Jackson, S.P.; Bartek, J. The DNA-damage response in human biology and disease. Nature 2009, 461, 1071–1078. [Google Scholar] [CrossRef] [PubMed]
  5. Huang, R.X.; Zhou, P.K. DNA damage response signaling pathways and targets for radiotherapy sensitization in cancer. Signal Transduct Target Ther. 2020, 5, 60. [Google Scholar] [CrossRef] [PubMed]
  6. Herbst, J.; Li, Q.Q.; De Veylder, L. Mechanistic insights into DNA damage recognition and checkpoint control in plants. Nat. Plants 2024, 10, 539–550. [Google Scholar] [CrossRef]
  7. Kornberg, R.D. Chromatin structure: A repeating unit of histones and DNA. Science 1974, 184, 868–871. [Google Scholar] [CrossRef]
  8. Luger, K.; Mäder, A.W.; Richmond, R.K.; Sargent, D.F.; Richmond, T.J. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 1997, 389, 251–260. [Google Scholar] [CrossRef]
  9. Richmond, T.J.; Davey, C.A. The structure of DNA in the nucleosome core. Nature 2003, 423, 145–150. [Google Scholar] [CrossRef]
  10. Maze, I.; Noh, K.M.; Soshnev, A.A.; Allis, C.D. Every amino acid matters: Essential contributions of histone variants to mammalian development and disease. Nat. Rev. Genet. 2014, 15, 259–271. [Google Scholar] [CrossRef]
  11. Sura, W.; Kabza, M.; Karlowski, W.M.; Bieluszewski, T.; Kus-Slowinska, M.; Pawełoszek, Ł.; Sadowski, J.; Ziolkowski, P.A. Dual Role of the Histone Variant H2A.Z in Transcriptional Regulation of Stress-Response Genes. Plant Cell 2017, 29, 791–807. [Google Scholar] [CrossRef] [PubMed]
  12. Venkatesh, S.; Workman, J.L. Histone exchange, chromatin structure and the regulation of transcription. Nat. Rev. Mol. Cell Biol. 2015, 16, 178–189. [Google Scholar] [CrossRef]
  13. Probst, A.V. Deposition and eviction of histone variants define functional chromatin states in plants. Curr. Opin. Plant Biol. 2022, 69, 102266. [Google Scholar] [CrossRef] [PubMed]
  14. Misteli, T.; Soutoglou, E. The emerging role of nuclear architecture in DNA repair and genome maintenance. Nat. Rev. Mol. Cell Biol. 2009, 10, 243–254. [Google Scholar] [CrossRef] [PubMed]
  15. Celeste, A.; Petersen, S.; Romanienko, P.J.; Fernandez-Capetillo, O.; Chen, H.T.; Sedelnikova, O.A.; Reina-San-Martin, B.; Coppola, V.; Meffre, E.; Difilippantonio, M.J.; et al. Genomic instability in mice lacking histone H2AX. Science 2002, 296, 922–927. [Google Scholar] [CrossRef] [PubMed]
  16. Novo, N.; Romero-Tamayo, S.; Marcuello, C.; Boneta, S.; Blasco-Machin, I.; Velázquez-Campoy, A.; Villanueva, R.; Moreno-Loshuertos, R.; Lostao, A.; Medina, M.; et al. Beyond a platform protein for the degradosome assembly: The Apoptosis-Inducing Factor as an efficient nuclease involved in chromatinolysis. PNAS Nexus 2023, 2, 312. [Google Scholar] [CrossRef] [PubMed]
  17. Bano, D.; Prehn, J.H.M. Apoptosis-Inducing Factor (AIF) in Physiology and Disease: The Tale of a Repented Natural Born Killer. EBioMedicine 2018, 30, 29–37. [Google Scholar] [CrossRef] [PubMed]
  18. Rogakou, E.P.; Pilch, D.R.; Orr, A.H.; Ivanova, V.S.; Bonner, W.M. DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J. Biol. Chem. 1998, 273, 5858–5868. [Google Scholar] [CrossRef] [PubMed]
  19. Rogakou, E.P.; Boon, C.; Redon, C.; Bonner, W.M. Megabase chromatin domains involved in DNA double-strand breaks in vivo. J. Cell Biol. 1999, 146, 905–916. [Google Scholar] [CrossRef]
  20. Bonner, W.M.; Redon, C.E.; Dickey, J.S.; Nakamura, A.J.; Sedelnikova, O.A.; Solier, S.; Pommier, Y. GammaH2AX and cancer. Nat. Rev. Cancer 2008, 8, 957–967. [Google Scholar] [CrossRef]
  21. Kinner, A.; Wu, W.; Staudt, C.; Iliakis, G. Gamma-H2AX in recognition and signaling of DNA double-strand breaks in the context of chromatin. Nucleic Acids Res. 2008, 36, 5678–5694. [Google Scholar] [CrossRef] [PubMed]
  22. Yuan, J.; Adamski, R.; Chen, J. Focus on histone variant H2AX: To be or not to be. FEBS Lett. 2010, 584, 3717–3724. [Google Scholar] [CrossRef] [PubMed]
  23. Pinto, D.M.; Flaus, A. Structure and function of histone H2AX. Subcell Biochem. 2010, 50, 55–78. [Google Scholar] [CrossRef] [PubMed]
  24. Burma, S.; Chen, B.P.; Murphy, M.; Kurimasa, A.; Chen, D.J. ATM phosphorylates histone H2AX in response to DNA double-strand breaks. J. Biol. Chem. 2001, 276, 42462–42467. [Google Scholar] [CrossRef]
  25. Ward, I.M.; Chen, J. Histone H2AX is phosphorylated in an ATR-dependent manner in response to replicational stress. J. Biol. Chem. 2001, 276, 47759–47762. [Google Scholar] [CrossRef]
  26. Lang, J.; Smetana, O.; Sanchez-Calderon, L.; Lincker, F.; Genestier, J.; Schmit, A.C.; Houlné, G.; Chabouté, M.E. Plant γH2AX foci are required for proper DNA DSB repair responses and colocalize with E2F factors. New Phytol. 2012, 194, 353–363. [Google Scholar] [CrossRef]
  27. Qi, Y.; Zhang, Y.; Baller, J.A.; Voytas, D.F. Histone H2AX and the small RNA pathway modulate both non-homologous end-joining and homologous recombination in plants. Mutat. Res. 2016, 783, 9–14. [Google Scholar] [CrossRef]
  28. Waterworth, W.M.; Wilson, M.; Wang, D.; Nuhse, T.; Warward, S.; Selley, J.; West, C.E. Phosphoproteomic analysis reveals plant DNA damage signalling pathways with a functional role for histone H2AX phosphorylation in plant growth under genotoxic stress. Plant J. 2019, 100, 1007–1021. [Google Scholar] [CrossRef]
  29. Chen, K.; Li, G.J.; Bressan, R.A.; Song, C.P.; Zhu, J.K.; Zhao, Y. Abscisic acid dynamics, signaling, and functions in plants. J. Integr. Plant Biol. 2020, 62, 25–54. [Google Scholar] [CrossRef] [PubMed]
  30. Muhammad Aslam, M.; Waseem, M.; Jakada, B.H.; Okal, E.J.; Lei, Z.; Saqib, H.S.A.; Yuan, W.; Xu, W.; Zhang, Q. Mechanisms of Abscisic Acid-Mediated Drought Stress Responses in Plants. Int. J. Mol. Sci. 2022, 23, 1084. [Google Scholar] [CrossRef] [PubMed]
  31. Finkelstein, R.R.; Gampala, S.S.; Rock, C.D. Abscisic acid signaling in seeds and seedlings. Plant Cell 2002, 14 (Suppl. S1), S15–S45. [Google Scholar] [CrossRef] [PubMed]
  32. Swiatek, A.; Lenjou, M.; Van Bockstaele, D.; Inzé, D.; Van Onckelen, H. Differential effect of jasmonic acid and abscisic acid on cell cycle progression in tobacco BY-2 cells. Plant Physiol. 2002, 128, 201–211. [Google Scholar] [CrossRef] [PubMed]
  33. Roy, S.; Das, K.P. Homologous Recombination Defective Arabidopsis Mutants Exhibit Enhanced Sensitivity to Abscisic Acid. PLoS ONE 2017, 12, e0169294. [Google Scholar] [CrossRef] [PubMed]
  34. Yin, H.; Zhang, X.; Liu, J.; Wang, Y.; He, J.; Yang, T.; Hong, X.; Yang, Q.; Gong, Z. Epigenetic regulation, somatic homologous recombination, and abscisic acid signaling are influenced by DNA polymerase epsilon mutation in Arabidopsis. Plant Cell 2009, 21, 386–402. [Google Scholar] [CrossRef] [PubMed]
  35. Rohde, A.; Kurup, S.; Holdsworth, M. ABI3 emerges from the seed. Trends Plant Sci. 2000, 5, 418–419. [Google Scholar] [CrossRef] [PubMed]
  36. Gazzarrini, S.; Song, L. LAFL Factors in Seed Development and Phase Transitions. Annu. Rev. Plant Biol. 2024. [Google Scholar] [CrossRef] [PubMed]
  37. Parcy, F.; Valon, C.; Raynal, M.; Gaubier-Comella, P.; Delseny, M.; Giraudat, J. Regulation of gene expression programs during Arabidopsis seed development: Roles of the ABI3 locus and of endogenous abscisic acid. Plant Cell 1994, 6, 1567–1582. [Google Scholar] [CrossRef]
  38. Nambara, E.; McCourt, P.; Naito, S. A regulatory role for the ABI3 gene in the establishment of embryo maturation in Arabidopsis thaliana. Development 1995, 121, 629–636. [Google Scholar] [CrossRef]
  39. Parcy, F.; Valon, C.; Kohara, A.; Miséra, S.; Giraudat, J. The ABSCISIC ACID-INSENSITIVE3, FUSCA3, and LEAFY COTYLEDON1 loci act in concert to control multiple aspects of Arabidopsis seed development. Plant Cell 1997, 9, 1265–1277. [Google Scholar] [CrossRef]
  40. Bedi, S.; Nag Chaudhuri, R. Transcription factor ABI3 auto-activates its own expression during dehydration stress response. FEBS Lett. 2018, 592, 2594–2611. [Google Scholar] [CrossRef]
  41. To, A.; Valon, C.; Savino, G.; Guilleminot, J.; Devic, M.; Giraudat, J.; Parcy, F. A network of local and redundant gene regulation governs Arabidopsis seed maturation. Plant Cell 2006, 18, 1642–1651. [Google Scholar] [CrossRef]
  42. Tian, R.; Wang, F.; Zheng, Q.; Niza, V.; Downie, A.B.; Perry, S.E. Direct and indirect targets of the arabidopsis seed transcription factor ABSCISIC ACID INSENSITIVE3. Plant J. 2020, 103, 1679–1694. [Google Scholar] [CrossRef]
  43. Song, J.; Xie, X.; Chen, C.; Shu, J.; Thapa, R.K.; Nguyen, V.; Bian, S.; Kohalmi, S.E.; Marsolais, F.; Zou, J.; et al. LEAFY COTYLEDON1 expression in the endosperm enables embryo maturation in Arabidopsis. Nat. Commun. 2021, 12, 3963. [Google Scholar] [CrossRef]
  44. Perruc, E.; Kinoshita, N.; Lopez-Molina, L. The role of chromatin-remodeling factor PKL in balancing osmotic stress responses during Arabidopsis seed germination. Plant J. 2007, 52, 927–936. [Google Scholar] [CrossRef] [PubMed]
  45. Barrero, J.M.; Millar, A.A.; Griffiths, J.; Czechowski, T.; Scheible, W.R.; Udvardi, M.; Reid, J.B.; Ross, J.J.; Jacobsen, J.V.; Gubler, F. Gene expression profiling identifies two regulatory genes controlling dormancy and ABA sensitivity in Arabidopsis seeds. Plant J. 2010, 61, 611–622. [Google Scholar] [CrossRef]
  46. Yan, L.; Wei, S.; Wu, Y.; Hu, R.; Li, H.; Yang, W.; Xie, Q. High-Efficiency Genome Editing in Arabidopsis Using YAO Promoter-Driven CRISPR/Cas9 System. Mol. Plant 2015, 8, 1820–1823. [Google Scholar] [CrossRef] [PubMed]
  47. Pecinka, A.; Liu, C.H. Drugs for plant chromosome and chromatin research. Cytogenet. Genome Res. 2014, 143, 51–59. [Google Scholar] [CrossRef]
  48. Maloof, J.N. Plant development: Slowing root growth naturally. Curr. Biol. 2004, 14, R395–R396. [Google Scholar] [CrossRef]
  49. Díaz, M.; Pečinková, P.; Nowicka, A.; Baroux, C.; Sakamoto, T.; Gandha, P.Y.; Jeřábková, H.; Matsunaga, S.; Grossniklaus, U.; Pecinka, A. The SMC5/6 Complex Subunit NSE4A Is Involved in DNA Damage Repair and Seed Development. Plant Cell 2019, 31, 1579–1597. [Google Scholar] [CrossRef] [PubMed]
  50. Zhang, J.; Xie, S.; Zhu, J.K.; Gong, Z. Requirement for flap endonuclease 1 (FEN1) to maintain genomic stability and transcriptional gene silencing in Arabidopsis. Plant J. 2016, 87, 629–640. [Google Scholar] [CrossRef]
  51. Nisa, M.U.; Huang, Y.; Benhamed, M.; Raynaud, C. The Plant DNA Damage Response: Signaling Pathways Leading to Growth Inhibition and Putative Role in Response to Stress Conditions. Front. Plant Sci. 2019, 10, 653. [Google Scholar] [CrossRef] [PubMed]
  52. Szurman-Zubrzycka, M.; Jędrzejek, P.; Szarejko, I. How Do Plants Cope with DNA Damage? A Concise Review on the DDR Pathway in Plants. Int. J. Mol. Sci. 2023, 24, 2404. [Google Scholar] [CrossRef] [PubMed]
  53. Cools, T.; Iantcheva, A.; Maes, S.; Van den Daele, H.; De Veylder, L. A replication stress-induced synchronization method for Arabidopsis thaliana root meristems. Plant J. 2010, 64, 705–714. [Google Scholar] [CrossRef]
  54. Wang, L.; Zhan, L.; Zhao, Y.; Huang, Y.; Wu, C.; Pan, T.; Qin, Q.; Xu, Y.; Deng, Z.; Li, J.; et al. The ATR-WEE1 kinase module inhibits the MAC complex to regulate replication stress response. Nucleic Acids Res. 2021, 49, 1411–1425. [Google Scholar] [CrossRef] [PubMed]
  55. Miao, R.; Yuan, W.; Wang, Y.; Garcia-Maquilon, I.; Dang, X.; Li, Y.; Zhang, J.; Zhu, Y.; Rodriguez, P.L.; Xu, W. Low ABA concentration promotes root growth and hydrotropism through relief of ABA INSENSITIVE 1-mediated inhibition of plasma membrane H(+)-ATPase 2. Sci. Adv. 2021, 7. [Google Scholar] [CrossRef]
  56. Waadt, R.; Seller, C.A.; Hsu, P.-K.; Takahashi, Y.; Munemasa, S.; Schroeder, J.I. Plant hormone regulation of abiotic stress responses. Nat. Rev. Mol. Cell Biol. 2022, 23, 680–694. [Google Scholar] [CrossRef] [PubMed]
  57. Nambara, E.; Suzuki, M.; Abrams, S.; McCarty, D.R.; Kamiya, Y.; McCourt, P. A screen for genes that function in abscisic acid signaling in Arabidopsis thaliana. Genetics 2002, 161, 1247–1255. [Google Scholar] [CrossRef] [PubMed]
  58. Delmas, F.; Sankaranarayanan, S.; Deb, S.; Widdup, E.; Bournonville, C.; Bollier, N.; Northey, J.G.; McCourt, P.; Samuel, M.A. ABI3 controls embryo degreening through Mendel’s I locus. Proc. Natl. Acad. Sci. USA 2013, 110, E3888–E3894. [Google Scholar] [CrossRef]
  59. Masubelele, N.H.; Dewitte, W.; Menges, M.; Maughan, S.; Collins, C.; Huntley, R.; Nieuwland, J.; Scofield, S.; Murray, J.A. D-type cyclins activate division in the root apex to promote seed germination in Arabidopsis. Proc. Natl. Acad. Sci. USA 2005, 102, 15694–15699. [Google Scholar] [CrossRef]
  60. Huefner, N.D.; Friesner, J.; Britt, A.B. Characterization of Two H2AX Homologues in Arabidopsis thaliana and their Response to Ionizing Radiation. In Induced Plant Mutations in the Genomics Era; FAO: Rome, Italy, 2009. [Google Scholar]
  61. Fulcher, N.; Sablowski, R. Hypersensitivity to DNA damage in plant stem cell niches. Proc. Natl. Acad. Sci. USA 2009, 106, 20984–20988. [Google Scholar] [CrossRef]
  62. Bogliolo, M.; Lyakhovich, A.; Callén, E.; Castellà, M.; Cappelli, E.; Ramírez, M.J.; Creus, A.; Marcos, R.; Kalb, R.; Neveling, K.; et al. Histone H2AX and Fanconi anemia FANCD2 function in the same pathway to maintain chromosome stability. EMBO J. 2007, 26, 1340–1351. [Google Scholar] [CrossRef]
  63. Culligan, K.; Tissier, A.; Britt, A. ATR regulates a G2-phase cell-cycle checkpoint in Arabidopsis thaliana. Plant Cell 2004, 16, 1091–1104. [Google Scholar] [CrossRef] [PubMed]
  64. Chen, Q.; Bian, C.; Wang, X.; Liu, X.; Ahmad Kassab, M.; Yu, Y.; Yu, X. ADP-ribosylation of histone variant H2AX promotes base excision repair. EMBO J. 2021, 40, e104542. [Google Scholar] [CrossRef] [PubMed]
  65. Yao, T.; Jin, D.; Liu, Q.; Gong, Z. Abscisic acid suppresses the highly occurred somatic homologous recombination in Arabidopsis rfc1 mutant. J. Genet. Genom. 2013, 40, 465–471. [Google Scholar] [CrossRef]
  66. Lopez-Molina, L.; Mongrand, S.; McLachlin, D.T.; Chait, B.T.; Chua, N.H. ABI5 acts downstream of ABI3 to execute an ABA-dependent growth arrest during germination. Plant J. 2002, 32, 317–328. [Google Scholar] [CrossRef]
  67. Murashige, T.; Skoog, F. A Revised Medium for Rapid Growth and Bio Assays with Tobacco Tissue Cultures. Physiol. Plant. 1962, 15, 473–497. [Google Scholar] [CrossRef]
  68. Bae, S.; Park, J.; Kim, J.S. Cas-OFFinder: A fast and versatile algorithm that searches for potential off-target sites of Cas9 RNA-guided endonucleases. Bioinformatics 2014, 30, 1473–1475. [Google Scholar] [CrossRef]
  69. Clough, S.J.; Bent, A.F. Floral dip: A simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 1998, 16, 735–743. [Google Scholar] [CrossRef]
  70. Yoo, S.D.; Cho, Y.H.; Sheen, J. Arabidopsis mesophyll protoplasts: A versatile cell system for transient gene expression analysis. Nat. Protoc. 2007, 2, 1565–1572. [Google Scholar] [CrossRef]
  71. Schmittgen, T.D.; Livak, K.J. Analyzing real-time PCR data by the comparative CT method. Nat. Protoc. 2008, 3, 1101–1108. [Google Scholar] [CrossRef]
  72. Pfaffl, M.W.; Tichopad, A.; Prgomet, C.; Neuvians, T.P. Determination of stable housekeeping genes, differentially regulated target genes and sample integrity: BestKeeper--Excel-based tool using pair-wise correlations. Biotechnol. Lett. 2004, 26, 509–515. [Google Scholar] [CrossRef] [PubMed]
Ijms 25 08940 g001
Figure 1. The phenotypes of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the Com lines grown under normal conditions. (A) Photographs of 15-day-old and 30-day-old plants grown under a long-day photoperiod (16 h light/8 h dark). White bars = 5 cm. (B) Measurement of fresh weight and flowering time for the indicated genotypes. The above-ground parts of 15-day-old plants were calculated as the fresh weight. The error bars represent the mean ± SD of three biological replicates, each with 15 plants. Lowercase letters denote the statistically significant differences among the groups (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test). (C) Photographs of longitudinal sections of siliques grown under a long-day photoperiod (16 h light/8 h dark). White bars = 1 mm. White asterisks indicate unfertilized ovules and red asterisks indicate aborted ovules. (D) Measurement of seed abortion rates for the indicated genotypes. Error bars represent the mean ± SD of three biological replicates, each comprising 15–20 siliques totally containing about 1000 seeds. Lowercase letters denote the significant differences among the groups (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test).
Figure 1. The phenotypes of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the Com lines grown under normal conditions. (A) Photographs of 15-day-old and 30-day-old plants grown under a long-day photoperiod (16 h light/8 h dark). White bars = 5 cm. (B) Measurement of fresh weight and flowering time for the indicated genotypes. The above-ground parts of 15-day-old plants were calculated as the fresh weight. The error bars represent the mean ± SD of three biological replicates, each with 15 plants. Lowercase letters denote the statistically significant differences among the groups (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test). (C) Photographs of longitudinal sections of siliques grown under a long-day photoperiod (16 h light/8 h dark). White bars = 1 mm. White asterisks indicate unfertilized ovules and red asterisks indicate aborted ovules. (D) Measurement of seed abortion rates for the indicated genotypes. Error bars represent the mean ± SD of three biological replicates, each comprising 15–20 siliques totally containing about 1000 seeds. Lowercase letters denote the significant differences among the groups (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test).
Ijms 25 08940 g001
Ijms 25 08940 g002
Figure 2. AtHTA3 and AtHTA5 play redundant roles in the response to DNA damage. (A,C) Sensitivity to genotoxic stress in the Col-0, the athta3 single mutants, the athta5 single mutants, the athta3-1 athta5-1 double mutants, and the Com lines. The plants were grown in a liquid MS medium, which was supplemented separately with 25 μM Zeocin, 75 p.p.m. Phleomycin (PM), 10 μM mitomycin C (MMC), 1 mM hydroxyurea (HU), or 100 p.p.m. methyl methanesulfonate (MMS) for 14 days. The dash lines were used to distinguish different genotypes. White bars = 5 mm. The atatr or atfen1 mutants were used as a control to assess DNA damage hypersensitivity. (B,D) Evaluation of the inhibitory effect of various DNA-damaging reagents on root growth. Relative root length was defined as the ratio of the root length under DNA damage conditions to the root length under control conditions. Error bars indicate the mean ± SD of three biological replicates, each with 20 seedlings. Lowercase letters represent the significant differences among groups (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test).
Figure 2. AtHTA3 and AtHTA5 play redundant roles in the response to DNA damage. (A,C) Sensitivity to genotoxic stress in the Col-0, the athta3 single mutants, the athta5 single mutants, the athta3-1 athta5-1 double mutants, and the Com lines. The plants were grown in a liquid MS medium, which was supplemented separately with 25 μM Zeocin, 75 p.p.m. Phleomycin (PM), 10 μM mitomycin C (MMC), 1 mM hydroxyurea (HU), or 100 p.p.m. methyl methanesulfonate (MMS) for 14 days. The dash lines were used to distinguish different genotypes. White bars = 5 mm. The atatr or atfen1 mutants were used as a control to assess DNA damage hypersensitivity. (B,D) Evaluation of the inhibitory effect of various DNA-damaging reagents on root growth. Relative root length was defined as the ratio of the root length under DNA damage conditions to the root length under control conditions. Error bars indicate the mean ± SD of three biological replicates, each with 20 seedlings. Lowercase letters represent the significant differences among groups (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test).
Ijms 25 08940 g002
Ijms 25 08940 g003
Figure 3. Tissue-specific expression patterns and subcellular localizations of AtHTA3 and AtHTA5. (A) Transgenic plants expressing AtHTA3pro:GUS and AtHTA5pro:GUS were stained with X-Gluc at different stages and tissues, including 15 d old plants (i), inflorescences and stems (ii), lateral roots (iii), guard cells (iv), 5 d old plants (v), root apexes (vi), styles, valves, septums, ovule funiculi, abscission zones in mature siliques (vii), and germinated seeds (viii), to measurement expression patterns of AtHTA3 and AtHTA5. Black bars = 3 cm (i); 2 cm (ii); 100 μm (iii); 5 mm (iv); 3 mm (v); 200 μm (vi); 1 cm (vii); 1 cm (viii). (B) Relative gene expression levels of AtHTA3 or AtHTA5 in the indicated tissues of Col-0 plants were determined by RT-qPCR, with GAPDH serving as the internal control. Data represent means ± SD of three biological replicates, each with indicated tissues from 10 to 20 plants for RNA extraction. Each circle on the plot is a replication of an independent experiment. Different colors indicate different tissues (C) Subcellular localizations of AtHTA3-GFP and AtHTA5-GFP. Protoplasts from Col-0 were co-transfected with constructs expressing AtHTA3-GFP or AtHTA5-GFP, as well as NLS-RFP as a nuclear marker. The fluorescence signals that resulted were analyzed with a confocal fluorescence microscope. GFP, green fluorescent protein; RFP, red fluorescent protein; Yellow signal: the overlap of green and red signals. White bars = 10 μm.
Figure 3. Tissue-specific expression patterns and subcellular localizations of AtHTA3 and AtHTA5. (A) Transgenic plants expressing AtHTA3pro:GUS and AtHTA5pro:GUS were stained with X-Gluc at different stages and tissues, including 15 d old plants (i), inflorescences and stems (ii), lateral roots (iii), guard cells (iv), 5 d old plants (v), root apexes (vi), styles, valves, septums, ovule funiculi, abscission zones in mature siliques (vii), and germinated seeds (viii), to measurement expression patterns of AtHTA3 and AtHTA5. Black bars = 3 cm (i); 2 cm (ii); 100 μm (iii); 5 mm (iv); 3 mm (v); 200 μm (vi); 1 cm (vii); 1 cm (viii). (B) Relative gene expression levels of AtHTA3 or AtHTA5 in the indicated tissues of Col-0 plants were determined by RT-qPCR, with GAPDH serving as the internal control. Data represent means ± SD of three biological replicates, each with indicated tissues from 10 to 20 plants for RNA extraction. Each circle on the plot is a replication of an independent experiment. Different colors indicate different tissues (C) Subcellular localizations of AtHTA3-GFP and AtHTA5-GFP. Protoplasts from Col-0 were co-transfected with constructs expressing AtHTA3-GFP or AtHTA5-GFP, as well as NLS-RFP as a nuclear marker. The fluorescence signals that resulted were analyzed with a confocal fluorescence microscope. GFP, green fluorescent protein; RFP, red fluorescent protein; Yellow signal: the overlap of green and red signals. White bars = 10 μm.
Ijms 25 08940 g003
Ijms 25 08940 g004
Figure 4. AtHTA3 and AtHTA5 function in the ABA response. (A,B) Measurement of cotyledon greening rates of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the Com lines. Plants were grown in liquid half-strength MS medium supplemented with DMSO, 0.25 μM or 0.5 μM ABA for 7 days. The cotyledon greening rate was the ratio of the number of seeds with cotyledon greening to the number of all cultivated seeds per genotype. Black bars = 5 mm. (C,D) Measurement of root length of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the Com lines. Plants were grown on half-strength MS medium for 4 days and then transferred to medium containing DMSO or 5 μM ABA for 14 days. The dash lines were used to distinguish different genotypes. The white lines at the bottom of each root show the approximate position of the root tip. White bars = 10 mm. Error bars indicate mean ± SD of three biological replicates, each with 50 seedlings in (A) and 12 seedlings in (C). Lowercase letters represent statistically significant differences (p < 0.05; two-way ANOVA, followed by Tukey’s multiple comparison test).
Figure 4. AtHTA3 and AtHTA5 function in the ABA response. (A,B) Measurement of cotyledon greening rates of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the Com lines. Plants were grown in liquid half-strength MS medium supplemented with DMSO, 0.25 μM or 0.5 μM ABA for 7 days. The cotyledon greening rate was the ratio of the number of seeds with cotyledon greening to the number of all cultivated seeds per genotype. Black bars = 5 mm. (C,D) Measurement of root length of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, and the Com lines. Plants were grown on half-strength MS medium for 4 days and then transferred to medium containing DMSO or 5 μM ABA for 14 days. The dash lines were used to distinguish different genotypes. The white lines at the bottom of each root show the approximate position of the root tip. White bars = 10 mm. Error bars indicate mean ± SD of three biological replicates, each with 50 seedlings in (A) and 12 seedlings in (C). Lowercase letters represent statistically significant differences (p < 0.05; two-way ANOVA, followed by Tukey’s multiple comparison test).
Ijms 25 08940 g004
Ijms 25 08940 g005
Figure 5. RT-qPCR validation analyses for validation of ABA-responsive gene expression. The 7-day-old Col-0 and athta3 athta5 double mutants were treated under 10 μM ABA treatment for 0 h and 3 h. GAPDH was used as an internal control. Error bars indicate mean ± SD of three biological replicates. Each point on the plot indicated one independent replicate. Statistical analyses were performed by Student’s test (ns indicates not significant; * indicates p < 0.05; ** indicates p < 0.01).
Figure 5. RT-qPCR validation analyses for validation of ABA-responsive gene expression. The 7-day-old Col-0 and athta3 athta5 double mutants were treated under 10 μM ABA treatment for 0 h and 3 h. GAPDH was used as an internal control. Error bars indicate mean ± SD of three biological replicates. Each point on the plot indicated one independent replicate. Statistical analyses were performed by Student’s test (ns indicates not significant; * indicates p < 0.05; ** indicates p < 0.01).
Ijms 25 08940 g005
Ijms 25 08940 g006
Figure 6. AtABI3 acts genetically downstream of AtHTA3 and AtHTA5. (A,B) Measurement of cotyledon greening rates of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, the athta3 atabi3-8 double mutants, the athta5 atabi3-8 double mutants, and the athta3 athta5 atabi3-8 triple mutants. Plants were cultivated in liquid half-strength MS medium, which was supplemented with DMSO as a solvent and either 0.25 µM or 0.5 µM ABA for 7 days. The cotyledon greening rate was the ratio of the number of seeds with cotyledon greening to the number of all cultivated seeds per genotype. Black bars = 5 mm. (C,D) Measurement of root length for indicated genotypes. Plants were cultivated on half-strength MS medium for 4 days and then transferred to a medium containing DMSO or 5 µM ABA for 14 days. The dash lines were used to distinguish different genotypes. The white lines at the bottom of each root show the approximate position of the root tip. White bars = 10 mm. The error bars indicate the mean ± SD of three biological replicates, each with 50 seedlings in (A) and 12 seedlings in (C). The letters represent statistically significant differences among groups (p < 0.05; two-way ANOVA, followed by Tukey’s multiple comparison test).
Figure 6. AtABI3 acts genetically downstream of AtHTA3 and AtHTA5. (A,B) Measurement of cotyledon greening rates of Col-0, the athta3 single mutants, the athta5 single mutants, the athta3 athta5 double mutants, the athta3 atabi3-8 double mutants, the athta5 atabi3-8 double mutants, and the athta3 athta5 atabi3-8 triple mutants. Plants were cultivated in liquid half-strength MS medium, which was supplemented with DMSO as a solvent and either 0.25 µM or 0.5 µM ABA for 7 days. The cotyledon greening rate was the ratio of the number of seeds with cotyledon greening to the number of all cultivated seeds per genotype. Black bars = 5 mm. (C,D) Measurement of root length for indicated genotypes. Plants were cultivated on half-strength MS medium for 4 days and then transferred to a medium containing DMSO or 5 µM ABA for 14 days. The dash lines were used to distinguish different genotypes. The white lines at the bottom of each root show the approximate position of the root tip. White bars = 10 mm. The error bars indicate the mean ± SD of three biological replicates, each with 50 seedlings in (A) and 12 seedlings in (C). The letters represent statistically significant differences among groups (p < 0.05; two-way ANOVA, followed by Tukey’s multiple comparison test).
Ijms 25 08940 g006
Ijms 25 08940 g007
Figure 7. The involvement of AtABI3 in the DNA damage response. (A,B) Sensitivity to genotoxic stress of Col-0 and atabi3-8. For a period of 14 days, the plants were cultivated in liquid MS medium and supplemented with variable concentrations of genotoxic reagents: 25 µM Zeocin, 75 p.p.m. Phleomycin (PM), 10 µM mitomycin C (MMC), 1 mM hydroxyurea (HU), or 100 p.p.m. methyl methanesulfonate (MMS) separately. The atatr and atfen1 mutants were employed as controls for DNA damage hypersensitivity. The dash lines were used to distinguish different genotypes. White bars = 5 mm. (C,D) Root length measurement. Relative root length was defined as the ratio of the root length under DNA damage conditions to the root length under control conditions. The mean ± SD of three biological replicates with 18 seedlings each is denoted by the error bars. Significant differences among groups are indicated by lowercase letters (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test). (E,F) RT-qPCR validation analyses of DNA damage response-related marker genes. Col-0 and atabi3-8 were treated under 10 μM MMC or 1 mM HU for 0 h, 6 h and 12 h. GAPDH was used as an internal control. Error bars indicate mean ± SD of three biological replicates. Each point on the plot indicated one independent replicate. Statistical analyses were performed by Student’s test (ns indicates not significant; * indicates p < 0.05; ** indicates p < 0.01; *** indicates p < 0.001; **** indicates p < 0.0001).
Figure 7. The involvement of AtABI3 in the DNA damage response. (A,B) Sensitivity to genotoxic stress of Col-0 and atabi3-8. For a period of 14 days, the plants were cultivated in liquid MS medium and supplemented with variable concentrations of genotoxic reagents: 25 µM Zeocin, 75 p.p.m. Phleomycin (PM), 10 µM mitomycin C (MMC), 1 mM hydroxyurea (HU), or 100 p.p.m. methyl methanesulfonate (MMS) separately. The atatr and atfen1 mutants were employed as controls for DNA damage hypersensitivity. The dash lines were used to distinguish different genotypes. White bars = 5 mm. (C,D) Root length measurement. Relative root length was defined as the ratio of the root length under DNA damage conditions to the root length under control conditions. The mean ± SD of three biological replicates with 18 seedlings each is denoted by the error bars. Significant differences among groups are indicated by lowercase letters (p < 0.05; one-way ANOVA, followed by Tukey’s multiple comparison test). (E,F) RT-qPCR validation analyses of DNA damage response-related marker genes. Col-0 and atabi3-8 were treated under 10 μM MMC or 1 mM HU for 0 h, 6 h and 12 h. GAPDH was used as an internal control. Error bars indicate mean ± SD of three biological replicates. Each point on the plot indicated one independent replicate. Statistical analyses were performed by Student’s test (ns indicates not significant; * indicates p < 0.05; ** indicates p < 0.01; *** indicates p < 0.001; **** indicates p < 0.0001).
Ijms 25 08940 g007
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Guo, P.; Wang, T.-J.; Wang, S.; Peng, X.; Kim, D.H.; Liu, Y. Arabidopsis Histone Variant H2A.X Functions in the DNA Damage-Coupling Abscisic Acid Signaling Pathway. Int. J. Mol. Sci. 2024, 25, 8940. https://doi.org/10.3390/ijms25168940

AMA Style

Guo P, Wang T-J, Wang S, Peng X, Kim DH, Liu Y. Arabidopsis Histone Variant H2A.X Functions in the DNA Damage-Coupling Abscisic Acid Signaling Pathway. International Journal of Molecular Sciences. 2024; 25(16):8940. https://doi.org/10.3390/ijms25168940

Chicago/Turabian Style

Guo, Peng, Tian-Jing Wang, Shuang Wang, Xiaoyuan Peng, Dae Heon Kim, and Yutong Liu. 2024. "Arabidopsis Histone Variant H2A.X Functions in the DNA Damage-Coupling Abscisic Acid Signaling Pathway" International Journal of Molecular Sciences 25, no. 16: 8940. https://doi.org/10.3390/ijms25168940

APA Style

Guo, P., Wang, T. -J., Wang, S., Peng, X., Kim, D. H., & Liu, Y. (2024). Arabidopsis Histone Variant H2A.X Functions in the DNA Damage-Coupling Abscisic Acid Signaling Pathway. International Journal of Molecular Sciences, 25(16), 8940. https://doi.org/10.3390/ijms25168940

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop