# Differences in RAD51 transcriptional response and cell cycle dynamics reveal varying sensitivity to DNA damage among Arabidopsis thaliana root cell types

Konstantin Kutashev, Anis Meschichi, Svenja Reeck, Alejandro Fonseca, Kevin Sartori, Charles I. White, Adrien Sicard, Stefanie Rosa

<figure><img src="image_1.png" /></figure>

## To cite this version:

Konstantin Kutashev, Anis Meschichi, Svenja Reeck, Alejandro Fonseca, Kevin Sartori, et al.. Differences in RAD51 transcriptional response and cell cycle dynamics reveal varying sensitivity to DNA damage among Arabidopsis thaliana root cell types. New Phytologist, In press, 10.1111/nph.19875. hal-04595318

HAL Id: hal-04595318

https://hal.science/hal-04595318

Submitted on 31 May 2024

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<figure><img src="image_2.png" /></figure>
# Differences in RAD51 transcriptional response and cell cycle dynamics reveal varying sensitivity to DNA damage among Arabidopsis thaliana root cell types

Konstantin Kutashev, Anis Meschichi, Svenja Reeck, Alejandro Fonseca, Kevin Sartori, Charles I. White, Adrien Sicard, Stefanie Rosa

► To cite this version:

Konstantin Kutashev, Anis Meschichi, Svenja Reeck, Alejandro Fonseca, Kevin Sartori, et al. Differences in RAD51 transcriptional response and cell cycle dynamics reveal varying sensitivity to DNA damage among Arabidopsis thaliana root cell types. New Phytologist, In press, 10.1111/nph.19875. hal-04595318

HAL Id: hal-04595318

https://hal.science/hal-04595318v1

Submitted on 31 May 2024

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>
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# Differences in RAD51 transcriptional response and cell cycle dynamics reveal varying sensitivity to DNA damage among *Arabidopsis thaliana* root cell types

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Konstantin Kutashev<fnref n="1" />, Anis Meschichi<fnref n="2" />, Svenja Reeck<fnref n="3" />, Alejandro Fonseca<fnref n="1" />, Kevin Sartori<fnref n="1" />, Charles White<fnref n="4" />, Adrien Sicard<fnref n="1" />, Stefanie Rosa<fnref n="1" />#

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<footnote n="1">¹ Swedish University of Agricultural Sciences, Plant Biology Department, Uppsala, Sweden.</footnote>

<footnote n="2">² Institute of Molecular Plant Biology, Department of Biology, Swiss Federal Institute of Technology Zürich, 8092 Zürich, Switzerland.</footnote>

<footnote n="3">³Department of Cell and Developmental Biology, John Innes Centre, Norwich Research Park NR4 7UH, UK</footnote>

<footnote n="4">⁴ Institut Génétique Reproduction et Développement (iGReD), Université Clermont Auvergne, UMR 6293, CNRS, U1103 INSERM, Clermont-Ferrand, France.</footnote>

<footnote>#corresponding author: stefanie.rosa@slu.se; Tel. +46 18 67 3324.</footnote>
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# Differences in RAD51 transcriptional response and cell cycle dynamics reveal varying sensitivity to DNA damage among *Arabidopsis thaliana* root cell types

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Konstantin Kutashev<fnref n="1" />, Anis Meschichi<fnref n="2" />, Svenja Reeck<fnref n="3" />, Alejandro Fonseca<fnref n="1" />, Kevin Sartori<fnref n="1" />, Charles White<fnref n="4" />, Adrien Sicard<fnref n="1" />, Stefanie Rosa<fnref n="1" />#

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<footnote n="1">¹ Swedish University of Agricultural Sciences, Plant Biology Department, Uppsala, Sweden.</footnote>

<footnote n="2">² Institute of Molecular Plant Biology, Department of Biology, Swiss Federal Institute of Technology Zürich, 8092 Zürich, Switzerland.</footnote>

<footnote n="3">³Department of Cell and Developmental Biology, John Innes Centre, Norwich Research Park NR4 7UH, UK</footnote>

<footnote n="4">⁴ Institut Génétique Reproduction et Développement (iGReD), Université Clermont Auvergne, UMR 6293, CNRS, U1103 INSERM, Clermont-Ferrand, France.</footnote>

<footnote>#corresponding author: stefanie.rosa@slu.se; Tel. +46 18 67 3324.</footnote>
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# SUMMARY

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* Throughout their lifecycle, plants are subjected to DNA damage from various sources, both environmental and endogenous. Investigating the mechanisms of the DNA damage response (DDR) is essential to unravel how plants adjust to the changing environment that can elicit varying amounts of DNA damage.

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* Using a combination of state-of-the-art cell biology methods including whole-mount single-molecule RNA fluorescence *in situ* hybridization (WM-smFISH), allowing detection of individual mRNA molecules in intact plant tissue and plant cell cycle reporter lines we investigated how the transcriptional activation of a key homologous recombination (HR) gene, *RAD51*, occurs in response to increasing amounts of DNA damage in *Arabidopsis thaliana* roots.

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* The results uncover consistent variations in *RAD51* transcriptional response and cell cycle arrest among distinct cell types and developmental zones. Furthermore, we demonstrate that DNA damage induced by genotoxic stress results in *RAD51* transcription throughout the whole cell cycle, dissociating its traditional link with S/G2 phases.

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* This work advances the current comprehension of DNA damage response in plants showing quantitative differences in DDR activation. In addition, it reveals new associations with the cell cycle and cell types, providing crucial insights for further studies of the broader response mechanisms in plants.

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**Keywords:** DNA damage, Double Strand Breaks; Homologous Recombination, *RAD51* transcription, Cell cycle arrest, Cell cycle checkpoints, *Arabidopsis thaliana*.

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# INTRODUCTION

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Plants, due to their sessile nature, are constantly exposed to various DNA damaging agents from both the environment and endogenous processes. One of the most dangerous lesions that can occur on the DNA are double-stranded breaks (DSBs) (Vítor *et al.*, 2020). The occurrence of this type of lesions requires immediate repair, resulting in DNA damage response (DDR) activation, recruitment of the DNA repair machinery to the lesion site and cell cycle arrest until the repair is complete (Preuss & Britt, 2003; Cools *et al.*, 2011). In most cases, DSBs are repaired by one of two mechanisms: non-homologous end joining (NHEJ) or homologous
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# SUMMARY

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* Throughout their lifecycle, plants are subjected to DNA damage from various sources, both environmental and endogenous. Investigating the mechanisms of the DNA damage response (DDR) is essential to unravel how plants adjust to the changing environment that can elicit varying amounts of DNA damage.

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* Using a combination of state-of-the-art cell biology methods including whole-mount single-molecule RNA fluorescence *in situ* hybridization (WM-smFISH), allowing detection of individual mRNA molecules in intact plant tissue and plant cell cycle reporter lines we investigated how the transcriptional activation of a key homologous recombination (HR) gene, *RAD51*, occurs in response to increasing amounts of DNA damage in *Arabidopsis thaliana* roots.

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* The results uncover consistent variations in *RAD51* transcriptional response and cell cycle arrest among distinct cell types and developmental zones. Furthermore, we demonstrate that DNA damage induced by genotoxic stress results in *RAD51* transcription throughout the whole cell cycle, dissociating its traditional link with S/G2 phases.

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* This work advances the current comprehension of DNA damage response in plants showing quantitative differences in DDR activation. In addition, it reveals new associations with the cell cycle and cell types, providing crucial insights for further studies of the broader response mechanisms in plants.

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**Keywords:** DNA damage, Double Strand Breaks; Homologous Recombination, *RAD51* transcription, Cell cycle arrest, Cell cycle checkpoints, *Arabidopsis thaliana*.

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# INTRODUCTION

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Plants, due to their sessile nature, are constantly exposed to various DNA damaging agents from both the environment and endogenous processes. One of the most dangerous lesions that can occur on the DNA are double-stranded breaks (DSBs) (Vítor *et al.*, 2020). The occurrence of this type of lesions requires immediate repair, resulting in DNA damage response (DDR) activation, recruitment of the DNA repair machinery to the lesion site and cell cycle arrest until the repair is complete (Preuss & Britt, 2003; Cools *et al.*, 2011). In most cases, DSBs are repaired by one of two mechanisms: non-homologous end joining (NHEJ) or homologous
recombination (HR) (West et al., 2004). HR is typically viewed as an error-free repair system that relies on an intact DNA strand acting as a template for reconstruction of the broken DNA strand (Schuermann et al., 2005).

Soon after the occurrence of DSB thousands of kilobases around the newly formed DSB are labeled by phosphorylated form of H2A.X histone variant (γ-H2AX) that participates in the early signaling of the lesion and recruitment of DNA repair machinery proteins (Rogakou et al., 1999; Stewart et al., 2003; Lang et al., 2012; Fan et al., 2022). Histone γ-H2AX levels were shown to correlate with DNA damage amounts (Friesner et al., 2005; Redon et al., 2009; Lee et al., 2019), and its dynamics of recruitment and loss are employed to measure DSB repair dynamics (Löbrich et al., 2010; Lee et al., 2019).

HR pathway is intricately connected to the S and G2 phases of the cell cycle, owing to its inherent need for an intact repair template, with the sister chromatid predominantly serving this function (Johnson, 2000; Saleh-Gohari, 2004; Saintigny et al., 2007; Goldfarb & Lichten, 2010; Bee et al., 2013). This regulation is accomplished through cell-cycle-linked transcriptional control of HR proteins and post-translational modifications of proteins during the S and G2 phases (Yata et al., 2012; Weimer et al., 2016; Lim et al., 2020). Cell cycle arrest is an integral part of the DDR, providing the necessary time for repair to take place thus ensuring the integrity of genetic material (Muschel et al., 1991; Raleigh & O'Connell, 2000; Chen et al., 2017). In A. thaliana, cell cycle arrest in response to DNA damage occurs mostly at G2/M and G1/S phase checkpoints (De Schutter et al., 2007; Cui et al., 2017; Cabral et al., 2020). Different sources of DNA damage arrest the cell cycle at different stages. For example, hydroxyurea or cadmium-induced DNA damage arrests the cell cycle at G1/S transition (Culligan et al., 2004; Cui et al., 2017), while γ-irradiation and aphidicolin arrest the cell cycle at G2/M transition (Culligan et al., 2004).

RAD51 protein plays a key role in repair via HR. RAD51 promotes essential strand-invasion step where resected 3' single-stranded DNA end aligns with a homologous template, ensuring proper placement of broken DNA strand overhangs (Shinohara et al., 1992; Abe et al., 2005; Li et al., 2004; Su et al., 2017; Wang et al., 2014; Yu et al., 2023; Banerjee & Roy, 2021). The widespread presence of RAD51 homologs across various species underscores its fundamental functional significance (Bonilla et al., 2020). Mutation in RAD51 gene is lethal in animals but
recombination (HR) (West et al., 2004). HR is typically viewed as an error-free repair system that relies on an intact DNA strand acting as a template for reconstruction of the broken DNA strand (Schuermann et al., 2005).

Soon after the occurrence of DSB thousands of kilobases around the newly formed DSB are labeled by phosphorylated form of H2A.X histone variant (γ-H2AX) that participates in the early signaling of the lesion and recruitment of DNA repair machinery proteins (Rogakou et al., 1999; Stewart et al., 2003; Lang et al., 2012; Fan et al., 2022). Histone γ-H2AX levels were shown to correlate with DNA damage amounts (Friesner et al., 2005; Redon et al., 2009; Lee et al., 2019), and its dynamics of recruitment and loss are employed to measure DSB repair dynamics (Löbrich et al., 2010; Lee et al., 2019).

HR pathway is intricately connected to the S and G2 phases of the cell cycle, owing to its inherent need for an intact repair template, with the sister chromatid predominantly serving this function (Johnson, 2000; Saleh-Gohari, 2004; Saintigny et al., 2007; Goldfarb & Lichten, 2010; Bee et al., 2013). This regulation is accomplished through cell-cycle-linked transcriptional control of HR proteins and post-translational modifications of proteins during the S and G2 phases (Yata et al., 2012; Weimer et al., 2016; Lim et al., 2020). Cell cycle arrest is an integral part of the DDR, providing the necessary time for repair to take place thus ensuring the integrity of genetic material (Muschel et al., 1991; Raleigh & O'Connell, 2000; Chen et al., 2017). In A. thaliana, cell cycle arrest in response to DNA damage occurs mostly at G2/M and G1/S phase checkpoints (De Schutter et al., 2007; Cui et al., 2017; Cabral et al., 2020). Different sources of DNA damage arrest the cell cycle at different stages. For example, hydroxyurea or cadmium-induced DNA damage arrests the cell cycle at G1/S transition (Culligan et al., 2004; Cui et al., 2017), while γ-irradiation and aphidicolin arrest the cell cycle at G2/M transition (Culligan et al., 2004).

RAD51 protein plays a key role in repair via HR. RAD51 promotes essential strand-invasion step where resected 3' single-stranded DNA end aligns with a homologous template, ensuring proper placement of broken DNA strand overhangs (Shinohara et al., 1992; Abe et al., 2005; Li et al., 2004; Su et al., 2017; Wang et al., 2014; Yu et al., 2023; Banerjee & Roy, 2021). The widespread presence of RAD51 homologs across various species underscores its fundamental functional significance (Bonilla et al., 2020). Mutation in RAD51 gene is lethal in animals but
dispensable for vegetative development in *A. thaliana* (Lim & Hasty, 1996; Tsuzuki *et al.*, 1996; Li *et al.*, 2004). Upon DNA damage, *RAD51* transcription is activated (Wang *et al.*, 2014; Feng *et al.*, 2017; Ryu *et al.*, 2019; Da Ines *et al.*, 2022) in a dose-dependent manner (Osakabe *et al.*, 2005; De Schutter *et al.*, 2007). RAD51 protein is subsequently loaded onto the lesion site by BRCA2 or CX3 complex (Wang *et al.*, 2010; Su *et al.*, 2017). This accumulation of RAD51 at the broken strand overhang (Flott *et al.*, 2011; Biedermann *et al.*, 2017; Da Ines *et al.*, 2022) facilitates the search for a homologous donor template (Hicks *et al.*, 2011; Coïc *et al.*, 2011; Meschichi *et al.*, 2022). The activity of RAD51 is tightly regulated at the post-translational level, serving as a substrate of multiple kinases in human (Sørensen *et al.*, 2005; Chabot *et al.*, 2019; Woo *et al.*, 2021) and budding yeasts (Flott *et al.*, 2011; Woo *et al.*, 2020, 2021). Although RAD51 phosphorylation by the cyclin kinase CDKB1-CYCB1 complex was reported *in vitro* for *A. thaliana*, its exact function remains elusive. Nevertheless, it is highly likely that this process is linked to RAD51 activation and recruitment to the DNA double-strand break sites, as evidenced by compromised RAD51 foci formation in *cycb1;1* mutants (Weimer *et al.*, 2016).

Most plant studies have analyzed *RAD51* expression using bulk measurements, combining material from multiple plants (Wang *et al.*, 2014; Ryu *et al.*, 2019). Although this approach is suitable for many purposes, it does not allow determining how gene expression is tuned at the level of individual plants, tissues, or cell types. In this study, we used single-molecule RNA FISH (smFISH) (Duncan *et al.*, 2017; Zhao *et al.*, 2023) to quantify the transcriptional response of *RAD51* at the cellular level to increasing amounts of DNA damage induced by DNA damaging agent zeocin (Adachi *et al.*, 2011). Our findings show a positive correlation between *RAD51* transcription and increasing amounts of damage, and we demonstrate that *RAD51* mRNA output reaches a maximum at the cellular level upon surpassing a certain damage threshold. Notably, *RAD51* transcriptional response was different between root cell types and developmental zones. Our data also shows prominent *RAD51* transcription outside S/G2 cell cycle phases under DNA damage, challenging the proposed strict association between HR and S/G2 phases.

## 129 MATERIALS AND METHODS

### 130 Plant material
dispensable for vegetative development in *A. thaliana* (Lim & Hasty, 1996; Tsuzuki *et al.*, 1996; Li *et al.*, 2004). Upon DNA damage, *RAD51* transcription is activated (Wang *et al.*, 2014; Feng *et al.*, 2017; Ryu *et al.*, 2019; Da Ines *et al.*, 2022) in a dose-dependent manner (Osakabe *et al.*, 2005; De Schutter *et al.*, 2007). RAD51 protein is subsequently loaded onto the lesion site by BRCA2 or CX3 complex (Wang *et al.*, 2010; Su *et al.*, 2017). This accumulation of RAD51 at the broken strand overhang (Flott *et al.*, 2011; Biedermann *et al.*, 2017; Da Ines *et al.*, 2022) facilitates the search for a homologous donor template (Hicks *et al.*, 2011; Coïc *et al.*, 2011; Meschichi *et al.*, 2022). The activity of RAD51 is tightly regulated at the post-translational level, serving as a substrate of multiple kinases in human (Sørensen *et al.*, 2005; Chabot *et al.*, 2019; Woo *et al.*, 2021) and budding yeasts (Flott *et al.*, 2011; Woo *et al.*, 2020, 2021). Although RAD51 phosphorylation by the cyclin kinase CDKB1-CYCB1 complex was reported *in vitro* for *A. thaliana*, its exact function remains elusive. Nevertheless, it is highly likely that this process is linked to RAD51 activation and recruitment to the DNA double-strand break sites, as evidenced by compromised RAD51 foci formation in *cycb1;1* mutants (Weimer *et al.*, 2016).

Most plant studies have analyzed *RAD51* expression using bulk measurements, combining material from multiple plants (Wang *et al.*, 2014; Ryu *et al.*, 2019). Although this approach is suitable for many purposes, it does not allow determining how gene expression is tuned at the level of individual plants, tissues, or cell types. In this study, we used single-molecule RNA FISH (smFISH) (Duncan *et al.*, 2017; Zhao *et al.*, 2023) to quantify the transcriptional response of *RAD51* at the cellular level to increasing amounts of DNA damage induced by DNA damaging agent zeocin (Adachi *et al.*, 2011). Our findings show a positive correlation between *RAD51* transcription and increasing amounts of damage, and we demonstrate that *RAD51* mRNA output reaches a maximum at the cellular level upon surpassing a certain damage threshold. Notably, *RAD51* transcriptional response was different between root cell types and developmental zones. Our data also shows prominent *RAD51* transcription outside S/G2 cell cycle phases under DNA damage, challenging the proposed strict association between HR and S/G2 phases.

## 129 MATERIALS AND METHODS

### 130 Plant material
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All *Arabidopsis thaliana* lines used in this study were derived from Columbia (Col-0) ecotype.

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Transgenic lines used in this study come from the following sources: RAD51-GFP line (Da

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Ines et al., 2013), Cytrap (Aki & Umeda, 2016), PlaCCI and CDT1-CFP lines (Desvoyes et al., 2020).

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## Plant growth

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Arabidopsis seeds were surface sterilized in 5% (v/v) sodium hypochlorite for 5 min and rinsed three times in sterile distilled water. Seeds were then stratified for 2 days at 4°C before germination in a growth chamber in a vertically oriented Petri dish containing 1% plant agar (Duchefa Biochemie, P1001.1000) MS medium plate, pH 5.7 (Gamborg et al., 1976). Plants were grown under a photoperiod of 16 hours day and 8 hours night and a temperature cycle of 22°C during the day and 20°C during the night.

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## Expression analysis using real-time RT-PCR (qPCR)

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For total RAD51 transcript analysis, 10-day-old *A. thaliana* (Col-0) seedlings were measured by qPCR. 9-day-old seedlings were transferred onto 1% plant agar MS medium plates containing different concentrations of zeocin (0 µM, 10 µM, 50 µM, 170 µM) (Gibco, 10072492). Seedling roots were cut with a razor blade and collected after overnight zeocin exposure. A total of 0.1g of roots per zeocin concentration was used. RNA was isolated using Quiagen RNeasy Plant Mini kit (Quiagen, 74904). RNA concentration was measured using Nanodrop ND-1000 spectrophotometer. A total of 1 µg of RNA was treated with DNase (Thermo Fisher, EN0521) and reverse transcribed with Reverse Transcriptase (Thermo Fisher, EP0441). This template was then used to quantify relative mRNA abundance using the SensiMix SYBR Low-ROX kit (Bioline), a LightCycler® 480 (Roche) and the primers described below. RAD51 expression was analyzed using normalization to PP2A gene using following primers: Rad51 forward GCGCAAGTAGATGGTTCAGC, Rad51 reverse TTCCTCAACGCCAACCTTGT, PP2A forward TAACGTGGCCAAAATGATGC, PP2A reverse GTTCTCCACAACCGCTTGGT. Reactions were performed in triplicate, results were calculated using the 2⁻ΔΔCT method, standard deviation values shown on a graph.

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## Single molecule fluorescence in situ hybridization (smFISH) on root squashes

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All *Arabidopsis thaliana* lines used in this study were derived from Columbia (Col-0) ecotype.

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Transgenic lines used in this study come from the following sources: RAD51-GFP line (Da Ines et al., 2013), Cytrap (Aki & Umeda, 2016), PlaCCI and CDT1-CFP lines (Desvoyes et al., 2020).

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## Plant growth

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Arabidopsis seeds were surface sterilized in 5% (v/v) sodium hypochlorite for 5 min and rinsed three times in sterile distilled water. Seeds were then stratified for 2 days at 4°C before germination in a growth chamber in a vertically oriented Petri dish containing 1% plant agar (Duchefa Biochemie, P1001.1000) MS medium plate, pH 5.7 (Gamborg et al., 1976). Plants were grown under a photoperiod of 16 hours day and 8 hours night and a temperature cycle of 22°C during the day and 20°C during the night.

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## Expression analysis using real-time RT-PCR (qPCR)

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For total RAD51 transcript analysis, 10-day-old *A. thaliana* (Col-0) seedlings were measured by qPCR. 9-day-old seedlings were transferred onto 1% plant agar MS medium plates containing different concentrations of zeocin (0 µM, 10 µM, 50 µM, 170 µM) (Gibco, 10072492). Seedling roots were cut with a razor blade and collected after overnight zeocin exposure. A total of 0.1g of roots per zeocin concentration was used. RNA was isolated using Quiagen RNeasy Plant Mini kit (Quiagen, 74904). RNA concentration was measured using Nanodrop ND-1000 spectrophotometer. A total of 1 µg of RNA was treated with DNase (Thermo Fisher, EN0521) and reverse transcribed with Reverse Transcriptase (Thermo Fisher, EP0441). This template was then used to quantify relative mRNA abundance using the SensiMix SYBR Low-ROX kit (Bioline), a LightCycler® 480 (Roche) and the primers described below. RAD51 expression was analyzed using normalization to PP2A gene using following primers: Rad51 forward GCGCAAGTAGATGGTTCAGC, Rad51 reverse TTCCTCAACGCCAACCTTGT, PP2A forward TAACGTGGCCAAAATGATGC, PP2A reverse GTTCTCCACAACCGCTTGGT. Reactions were performed in triplicate, results were calculated using the 2⁻ΔΔCT method, standard deviation values shown on a graph.

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## Single molecule fluorescence *in situ* hybridization (smFISH) on root squashes

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smFISH was performed on 5-6 days old seedlings according to previously published protocol (Duncan *et al.*, 2017) using probes designed against RAD51 and PP2A genes (listed in Supplementary Table 2). Seedlings were transferred onto MS medium plates containing selected concentrations of zeocin (Gibco, 10072492) or no zeocin for control sample. Seedlings were collected after overnight zeocin exposure and treated further according to protocol.

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## 173 Immunodetection

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5-6 days old seedlings were transferred onto MS medium containing selected concentrations of zeocin overnight. Roots were then cut off from seedlings using a razor blade and fixed in 4% paraformaldehyde solution for 30 minutes in glass dishes. Roots were then washed twice with 1x PBS. 5 roots were then arranged on a slide in similar orientation, covered by a glass coverslip and squashed manually by applying pressure on coverslip. The slide was then submerged in liquid nitrogen until freezing and taken out, coverslip was then removed using a razor blade. Slides were left to dry at room temperature for 30 minutes. Samples were rinsed with 1x PBS three times and incubated with blocking buffer (0.5% BSA (Sigma-Aldrich A7030) in 1x PBS) in humid chamber at 37°C for 30 minutes. To ensure minimal disturbance of the sample we used small pieces of polypropylene waste bags instead of glass coverslips at all incubation stages of the protocol. Excess blocking buffer was removed, samples were incubated at 37°C overnight in a humid chamber with γH2AX primary antibody (Charbonnel *et al.*, 2010), provided by Charles White. Antibody was diluted 1:700 in 0.5% BSA. Slides were then rinsed with PBST buffer three times (1x PBS, 0,01% Tween20 (Sigma-Aldrich 8.22184)) and incubated with PBST buffer for 5 min. Secondary antibody (Agrisera, AS09633) diluted 1:200 in 0.5% BSA was then applied, and samples were incubated in a humid chamber at 37°C for 2 hours. Slides were rinsed three times with PBST buffer and incubated with 1x PBS buffer 2x 5 min. Excess buffer was removed, and samples were mounted in Vectashield medium (Vector laboratories, H-1000) containing DAPI diluted 1:1000 (Thermo Fisher, 62248).

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## 196 Sequential smFISH and immunodetection

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SmFISH and immunodetection protocols were performed sequentially in the described order. SmFISH in root squashes was performed first according to the protocol mentioned above. After imaging the coverslips were gently removed using additional volumes of 1x PBS. Samples

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smFISH was performed on 5-6 days old seedlings according to previously published protocol (Duncan *et al.*, 2017) using probes designed against RAD51 and PP2A genes (listed in Supplementary Table 2). Seedlings were transferred onto MS medium plates containing selected concentrations of zeocin (Gibco, 10072492) or no zeocin for control sample. Seedlings were collected after overnight zeocin exposure and treated further according to protocol.

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## 173 Immunodetection

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5-6 days old seedlings were transferred onto MS medium containing selected concentrations of zeocin overnight. Roots were then cut off from seedlings using a razor blade and fixed in 4% paraformaldehyde solution for 30 minutes in glass dishes. Roots were then washed twice with 1x PBS. 5 roots were then arranged on a slide in similar orientation, covered by a glass coverslip and squashed manually by applying pressure on coverslip. The slide was then submerged in liquid nitrogen until freezing and taken out, coverslip was then removed using a razor blade. Slides were left to dry at room temperature for 30 minutes. Samples were rinsed with 1x PBS three times and incubated with blocking buffer (0.5% BSA (Sigma-Aldrich A7030) in 1x PBS) in humid chamber at 37°C for 30 minutes. To ensure minimal disturbance of the sample we used small pieces of polypropylene waste bags instead of glass coverslips at all incubation stages of the protocol. Excess blocking buffer was removed, samples were incubated at 37°C overnight in a humid chamber with γH2AX primary antibody (Charbonnel *et al.*, 2010), provided by Charles White. Antibody was diluted 1:700 in 0.5% BSA. Slides were then rinsed with PBST buffer three times (1x PBS, 0,01% Tween20 (Sigma-Aldrich 8.22184)) and incubated with PBST buffer for 5 min. Secondary antibody (Agrisera, AS09633) diluted 1:200 in 0.5% BSA was then applied, and samples were incubated in a humid chamber at 37°C for 2 hours. Slides were rinsed three times with PBST buffer and incubated with 1x PBS buffer 2x 5 min. Excess buffer was removed, and samples were mounted in Vectashield medium (Vector laboratories, H-1000) containing DAPI diluted 1:1000 (Thermo Fisher, 62248).

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## 196 Sequential smFISH and immunodetection

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SmFISH and immunodetection protocols were performed sequentially in the described order. SmFISH in root squashes was performed first according to the protocol mentioned above. After imaging the coverslips were gently removed using additional volumes of 1x PBS. Samples

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201 were rinsed with 1x PBS three times and samples were processed according to
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204 **Whole-mount smFISH (WM-smFISH)**

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206 WM-smFISH was performed on 5-6 days old seedlings according to previously published
207 protocol (Zhao et al., 2023) using probes designed against Rad51 gene (listed in Supplementary
208 Table 1). Seedlings were transferred onto MS medium plates containing selected
209 concentrations of zeocin (Gibco, 10072492) or no zeocin for control sample. Seedlings were
210 collected after overnight zeocin exposure and treated further according to protocol.

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212 **Sequential WM-smFISH and 5-ethynyl-2'-deoxyuridine (EdU) labeling**
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214 5-6 days old seedling were first transferred onto the MS medium containing zeocin for 10
215 hours. Seedlings were then transferred onto MS medium containing same concentration of
216 zeocin and 20 µM EdU (Invitrogen, A10044) for two hours. WM-smFISH was performed first
217 according to the described protocol. After imaging, coverslips were gently removed from the
218 samples using additional volumes of 2x SSC buffer. Samples were then rinsed with 2x SSC
219 buffer three times and incubated with 3% BSA in 1x PBS solution at 37°C in a humid chamber
220 for 15 minutes. Samples were incubated with Click-iT reaction cocktail (Invitrogen C10269)
221 mixed according to the manufacturer's instructions with addition of Alexa Fluor 488 azide
222 (Thermo Fisher, A10266), 500x dilution. Samples were then rinsed and incubated with a wash
223 buffer (10% formamide (Thermo Scientific, 17899) and 2xSSC) for 5 minutes. Samples were
224 incubated with SCRI Renaissance 2200 solution (Musielak et al., 2015) for 15 minutes at 37°C
225 in a humid chamber. Slides were rinsed and incubated for 5 min in the wash buffer. Samples
226 were then mounted in a drop of Vectashield medium.

227
228 **RAD51 mRNA half-life quantification**
229

230 5-6 days old seedling were transferred onto MS medium containing 10µM zeocin for selected
time periods: 12, 10, 8, 6 hours. Seedlings exposed to zeocin for 10, 8 and 6 hours were then
231 transferred to MS medium containing zeocin with Actinomycin D (Thermo Fisher,
232 J60148.LB0) or zeocin with DMSO (Sigma-Aldrich, D4540) for 2, 4 and 6 hours accordingly.
233 Seedlings were then collected and processed according to smFISH protocol for root squashes
234
201 were rinsed with 1x PBS three times and samples were processed according to
202 immunodetection protocol above.

203
204 Whole-mount smFISH (WM-smFISH)

205
206 WM-smFISH was performed on 5-6 days old seedlings according to previously published
207 protocol (Zhao et al., 2023) using probes designed against Rad51 gene (listed in Supplementary
208 Table 1). Seedlings were transferred onto MS medium plates containing selected
209 concentrations of zeocin (Gibco, 10072492) or no zeocin for control sample. Seedlings were
210 collected after overnight zeocin exposure and treated further according to protocol.

211
212 Sequential WM-smFISH and 5-ethynyl-2'-deoxyuridine (EdU) labeling

213
214 5-6 days old seedling were first transferred onto the MS medium containing zeocin for 10
215 hours. Seedlings were then transferred onto MS medium containing same concentration of
216 zeocin and 20 µM EdU (Invitrogen, A10044) for two hours. WM-smFISH was performed first
217 according to the described protocol. After imaging, coverslips were gently removed from the
218 samples using additional volumes of 2x SSC buffer. Samples were then rinsed with 2x SSC
219 buffer three times and incubated with 3% BSA in 1x PBS solution at 37°C in a humid chamber
220 for 15 minutes. Samples were incubated with Click-iT reaction cocktail (Invitrogen C10269)
221 mixed according to the manufacturer's instructions with addition of Alexa Fluor 488 azide
222 (Thermo Fisher, A10266), 500x dilution. Samples were then rinsed and incubated with a wash
223 buffer (10% formamide (Thermo Scientific, 17899) and 2xSSC) for 5 minutes. Samples were
224 incubated with SCRI Renaissance 2200 solution (Musielak et al., 2015) for 15 minutes at 37°C
225 in a humid chamber. Slides were rinsed and incubated for 5 min in the wash buffer. Samples
226 were then mounted in a drop of Vectashield medium.

227
228 RAD51 mRNA half-life quantification

229
230 5-6 days old seedling were transferred onto MS medium containing 10µM zeocin for selected
time periods: 12, 10, 8, 6 hours. Seedlings exposed to zeocin for 10, 8 and 6 hours were then
231 transferred to MS medium containing zeocin with Actinomycin D (Thermo Fisher,
232 J60148.LB0) or zeocin with DMSO (Sigma-Aldrich, D4540) for 2, 4 and 6 hours accordingly.
233 Seedlings were then collected and processed according to smFISH protocol for root squashes
234
235 using probes for RAD51 gene. The decay rate ($k_{decay}$) for RAD51 and then its half-life ($t_{1/2}$) 236 were calculated by adjusting the number of molecules per cell (N) counted in the smFISH 237 images as an exponential function of time (t). The mathematical adjustment for N(t) was 238 developed in R assuming a constant decay rate, according to the function: $N(t) = e^{-k_{decay} * t}$, 239 then the half-life was calculated using the formula: $\ln(2)/k_{decay}$ (Narsai et al., 2007; Sorenson 240 et al., 2018). 241

241

## 242 Image acquisition

243

244 Samples were imaged on Zeiss LSM780 and LSM800 inverted confocal microscopes (Zen 245 Black Software) using a 63X water-immersion objective (1.20 NA). smFISH on root squashes 246 imaging was performed using widefield mode, we used a cooled quad-port CCD (charge- 247 coupled device) ZEISS Axiocam 503 mono camera. A series of optical sections with z-steps of 248 0.22 µm were collected throughout the whole cell volume. For DAPI imaging an excitation 249 filter of 335-383 nm was used and emission was collected at 420 - 470 nm. Quasar570 250 fluorescent probes were imaged using 533-558 nm excitation filter and 570-640 nm signal 251 detection range. For immunostaining experiments were did not use widefield mode, for DAPI 252 signals excitation line of 405 nm was used with emission detection at 410-600 nm. For GFP 253 signals of labelled histone γH2AX excitation line of 488 nm and emission at 490 - 540 nm 254 settings were used. Imaging was performed in a manually adjusted single plain selected to 255 have a maximal number of nuclei in focus. 256

256 WM smFISH imaging was performed in confocal mode using a 63X water-immersion 257 objective (1.20 NA). For SCRI Renaissance 2200 imaging we used a 405 nm laser line and and 258 emission was collected at 410-600 nm. Quasar570 probe signals were captured with 561 nm 259 excitation line and emission collection at 565-700 nm. CFP signals were imaged using 455 nm 260 excitation line and emission detection at 460-600 nm. 261

## 261 Image analysis

262

263 smFISH

264 Nuclei and cellular outlines in smFISH were defined using CellProfiler software (Stirling et al., 265 2021). RNA foci were detected and counted using FISH-quant-v3 (Mueller et al., 2013) in 266 Matlab. First, the “cell segmentation” tool was used to generate text files with the outline
235 using probes for RAD51 gene. The decay rate ($k_{decay}$) for RAD51 and then its half-life ($t_{1/2}$) 236 were calculated by adjusting the number of molecules per cell (N) counted in the smFISH 237 images as an exponential function of time (t). The mathematical adjustment for N(t) was 238 developed in R assuming a constant decay rate, according to the function: $N(t) = e^{-k_{decay} * t}$, 239 then the half-life was calculated using the formula: $\ln(2)/k_{decay}$ (Narsai et al., 2007; Sorenson 240 et al., 2018). 241

241

## 242 Image acquisition

243

244 Samples were imaged on Zeiss LSM780 and LSM800 inverted confocal microscopes (Zen 245 Black Software) using a 63X water-immersion objective (1.20 NA). smFISH on root squashes 246 imaging was performed using widefield mode, we used a cooled quad-port CCD (charge- 247 coupled device) ZEISS Axiocam 503 mono camera. A series of optical sections with z-steps of 248 0.22 µm were collected throughout the whole cell volume. For DAPI imaging an excitation 249 filter of 335-383 nm was used and emission was collected at 420 - 470 nm. Quasar570 250 fluorescent probes were imaged using 533-558 nm excitation filter and 570-640 nm signal 251 detection range. For immunostaining experiments were did not use widefield mode, for DAPI 252 signals excitation line of 405 nm was used with emission detection at 410-600 nm. For GFP 253 signals of labelled histone γH2AX excitation line of 488 nm and emission at 490 - 540 nm 254 settings were used. Imaging was performed in a manually adjusted single plain selected to 255 have a maximal number of nuclei in focus. 256

256 WM smFISH imaging was performed in confocal mode using a 63X water-immersion 257 objective (1.20 NA). For SCRI Renaissance 2200 imaging we used a 405 nm laser line and and 258 emission was collected at 410-600 nm. Quasar570 probe signals were captured with 561 nm 259 excitation line and emission collection at 565-700 nm. CFP signals were imaged using 455 nm 260 excitation line and emission detection at 460-600 nm. 261

## 261 Image analysis

262

263 smFISH

264 Nuclei and cellular outlines in smFISH were defined using CellProfiler software (Stirling et al., 265 2021). RNA foci were detected and counted using FISH-quant-v3 (Mueller et al., 2013) in 266 Matlab. First, the “cell segmentation” tool was used to generate text files with the outline
coordinates for the nuclei and cell masks. The outlines were uploaded, and images were pre-processed for increasing their signal-to-noise ratio though a dual-Gaussian filtering followed by a Gaussian Kernel. Dots were detected in the filtered image, first pre-detecting fluorescent foci with fluorescence over a threshold. Then, the pre-detected dots were fitted to a Gaussian fluorescence based on a point-spread function. Images were analyzed in the batch mode, and false positives were removed in the end by thresholding the Sigma-XY, amplitude, and pixel-intensity parameters to Gaussian distributions.

274

275 WM-smFISH

276 Cell segmentation was performed with Cellpose software (Stringer et al., 2021), using an 277 algorithm trained by us. RNA foci were detected and counted using FISH-quant-v3 (Mueller et 278 al., 2013) as described above. For RAD51-GFP line the signal intensities of both mRNA and 279 protein channels were quantified in CellProfiler software (Stirling et al., 2021). Colocalization 280 analysis and heatmap visualization was performed using CellProfiler software (Stirling et al., 281 2021).

282 Images from EdU staining, Cytrap and PlaCCI lines were analysed manually using ImageJ 283 software.

284

285 Correlation analysis of γH2AX signal and RAD51 transcription

286 Data on RAD51 transcription and H2AX levels were collected from the same cells for 287 correlation analysis. γH2AX integrated density was measured using ImageJ software and 288 normalized to DAPI integrated density. The number of detected RAD51 mRNA molecules was 289 normalized by the cell area to correct for cell size difference. Values obtained for both 290 parameters were log transformed. Data was visualized and correlation was evaluated using R 291 studio ggplot2 package.

292

293 RESULTS

294 RAD51 transcriptional response to increasing DNA damage levels

295 To elucidate RAD51 transcriptional response to DNA damage we first assessed RAD51 mRNA 296 levels on roots from Col-0 plants treated with increasing concentrations of the DSB-inducing 297 agent zeocin (0 µM, 10 µM, 50 µM, 170 µM) using real-time quantitative PCR (RT-qPCR). 298 The results demonstrated an increase in RAD51 mRNA quantities with increasing zeocin
coordinates for the nuclei and cell masks. The outlines were uploaded, and images were pre-processed for increasing their signal-to-noise ratio though a dual-Gaussian filtering followed by a Gaussian Kernel. Dots were detected in the filtered image, first pre-detecting fluorescent foci with fluorescence over a threshold. Then, the pre-detected dots were fitted to a Gaussian fluorescence based on a point-spread function. Images were analyzed in the batch mode, and false positives were removed in the end by thresholding the Sigma-XY, amplitude, and pixel-intensity parameters to Gaussian distributions.

274

275 *WM-smFISH*

276 Cell segmentation was performed with Cellpose software (Stringer *et al.*, 2021), using an algorithm trained by us. RNA foci were detected and counted using FISH-quant-v3 (Mueller *et al.*, 2013) as described above. For RAD51-GFP line the signal intensities of both mRNA and protein channels were quantified in CellProfiler software (Stirling *et al.*, 2021). Colocalization analysis and heatmap visualization was performed using CellProfiler software (Stirling *et al.*, 2021).

282 Images from EdU staining, Cytrap and PlaCCI lines were analysed manually using ImageJ software.

284

285 *Correlation analysis of γH2AX signal and RAD51 transcription*

286 Data on RAD51 transcription and H2AX levels were collected from the same cells for correlation analysis. γH2AX integrated density was measured using ImageJ software and normalized to DAPI integrated density. The number of detected RAD51 mRNA molecules was normalized by the cell area to correct for cell size difference. Values obtained for both parameters were log transformed. Data was visualized and correlation was evaluated using R studio ggplot2 package.

292

293 **RESULTS**

294 *RAD51 transcriptional response to increasing DNA damage levels*

295 To elucidate RAD51 transcriptional response to DNA damage we first assessed RAD51 mRNA levels on roots from Col-0 plants treated with increasing concentrations of the DSB-inducing agent zeocin (0 µM, 10 µM, 50 µM, 170 µM) using real-time quantitative PCR (RT-qPCR).
296
297
298 The results demonstrated an increase in RAD51 mRNA quantities with increasing zeocin
concentrations (Fig. 1a). To investigate RAD51 transcriptional upregulation as a function of DNA damage at the cellular and tissue level we employed smFISH (Duncan *et al.*, 2016), a method that allows absolute quantification of transcripts in individual cells (Fig. 1b, c(i)), using the same material preparation as for immunodetection. Consistent with the qPCR data (Fig. 1a) smFISH results revealed an increase in the total number of RAD51 mRNAs in root tissue with increasing zeocin concentrations (Fig. 1d). Of note, the total number of RAD51 mRNAs did not seem to increase in direct proportion to the concentration of zeocin. To evaluate the increase in DNA damage levels corresponding to increasing zeocin concentrations, we quantified γ-H2AX levels by immunodetection as a proxy marker for DSB levels in individual root cells. Single cell spreading achieved by root squashing facilitated antibody penetration required for immunodetection (Fig. 1c(ii)). The results showed an accumulation of γ-H2AX in the nuclei of root cells in response to growing zeocin concentrations (Fig. 1e), confirming the increase in DSBs. The observed increase in γ-H2AX accumulation was not directly proportional to the increase in zeocin concentration as the number of RAD51 mRNAs. To assess the direct relationship between RAD51 transcription and the extent of DNA damage within individual cell, we performed a sequential RAD51-smFISH/γH2AX-immunodetection protocol on cells obtained from root squashes and evaluated mRNA and DNA damage levels on the same cells (Fig. 1c). This analysis revealed a positive correlation between the number of RAD51 mRNA molecules per cell and the γ-H2AX levels with a correlation coefficient R=0.62 (p<1.4e-14) (Fig. 1f). Our analysis indicated that the interaction between the two variables is best described by a linear model with deviance of fit (DOF) value of 18.44615. DOF value of the exponential model, indicating a potential limit to the possible number of RAD51 mRNAs per cell, was however only slightly higher, 18.99684 (Fig. S1). Importantly, the mRNA counts for the house-keeping gene PP2A remained constant across zeocin concentrations (Fig. S1), confirming the specific RAD51 upregulation with increasing damage.

Cell-to-cell variability in RAD51 transcriptional response

To unravel potential differences in RAD51 transcriptional activation between different cell types and developmental zones of the root we performed recently developed whole-mount smFISH (WM-smFISH) protocol (Zhao *et al.*, 2023). This method overcomes the limitations of traditional root squash sample preparation, enabling the assessment of transcript numbers within intact tissues (Fig. 2a, Fig. S2). Heatmaps of RAD51 mRNA molecules per cell were
concentrations (Fig. 1a). To investigate RAD51 transcriptional upregulation as a function of DNA damage at the cellular and tissue level we employed smFISH (Duncan *et al.*, 2016), a method that allows absolute quantification of transcripts in individual cells (Fig. 1b, c(i)), using the same material preparation as for immunodetection. Consistent with the qPCR data (Fig. 1a) smFISH results revealed an increase in the total number of RAD51 mRNAs in root tissue with increasing zeocin concentrations (Fig. 1d). Of note, the total number of RAD51 mRNAs did not seem to increase in direct proportion to the concentration of zeocin. To evaluate the increase in DNA damage levels corresponding to increasing zeocin concentrations, we quantified γ-H2AX levels by immunodetection as a proxy marker for DSB levels in individual root cells. Single cell spreading achieved by root squashing facilitated antibody penetration required for immunodetection (Fig. 1c(ii)). The results showed an accumulation of γ-H2AX in the nuclei of root cells in response to growing zeocin concentrations (Fig. 1e), confirming the increase in DSBs. The observed increase in γ-H2AX accumulation was not directly proportional to the increase in zeocin concentration as the number of RAD51 mRNAs. To assess the direct relationship between RAD51 transcription and the extent of DNA damage within individual cell, we performed a sequential RAD51-smFISH/γH2AX-immunodetection protocol on cells obtained from root squashes and evaluated mRNA and DNA damage levels on the same cells (Fig. 1c). This analysis revealed a positive correlation between the number of RAD51 mRNA molecules per cell and the γ-H2AX levels with a correlation coefficient R=0.62 (p<1.4e-14) (Fig. 1f). Our analysis indicated that the interaction between the two variables is best described by a linear model with deviance of fit (DOF) value of 18.44615. DOF value of the exponential model, indicating a potential limit to the possible number of RAD51 mRNAs per cell, was however only slightly higher, 18.99684 (Fig. S1). Importantly, the mRNA counts for the house-keeping gene PP2A remained constant across zeocin concentrations (Fig. S1), confirming the specific RAD51 upregulation with increasing damage.

Cell-to-cell variability in RAD51 transcriptional response

To unravel potential differences in RAD51 transcriptional activation between different cell types and developmental zones of the root we performed recently developed whole-mount smFISH (WM-smFISH) protocol (Zhao *et al.*, 2023). This method overcomes the limitations of traditional root squash sample preparation, enabling the assessment of transcript numbers within intact tissues (Fig. 2a, Fig. S2). Heatmaps of RAD51 mRNA molecules per cell were
generated to visualize number of transcripts per cell across root tissues (Fig. 2b). The results
revealed that the number of transcribing cells as well as the number of RAD51 mRNA
molecules detected per cell increases in response to increasing zeocin concentrations. This
pattern is further evident in the histogram quantification (Fig. 2c), depicting a progressive rise
in the number of actively transcribing cells with increasing zeocin concentrations. This data
also indicates a possible upper boundary on the number of RAD51 mRNA molecules per cell.
Indeed, one cannot observe a large difference in mRNA numbers per cell between 50 µM and
170 µM zeocin concentrations despite the large increase in concentration, as visualized on
heatmaps (Fig. 2b) and in a graph form (Fig. 2c, d).

Importantly, our results indicate substantial variability among root cells in their sensitivity to DSBs induced by zeocin, as revealed by the non-uniform heatmaps. Some cells exhibited a strong transcriptional response even at a 10 µM zeocin concentration, with mRNA counts comparable to those induced by 50 µM and 170 µM (Fig. 2b, c). Conversely, certain cells displayed low mRNA counts even after exposure to 50 µM and 170 µM zeocin (Fig. 2b, c). To discern potential distinctions between cell types, we plotted the number of RAD51 mRNAs in different root cell types (Epidermis, Cortex, Endodermis, and Stele). The results demonstrated that RAD51 transcriptional response within stele cells was distinct from the other root cell types analyzed showing higher per cell mRNA output (Fig. 2d). The comparison also indicated no difference in mRNA counts per cell between 50 µM and 170 µM zeocin treated samples (Fig. 2d), potentially arguing in preference of previously proposed limit to per cell transcript output (Fig. S1a). Notably, across developmental regions RAD51 transcriptional output decreased in the elongation zone in comparison with the meristem region (Fig. S3) consistent with previous reports (Da Ines et al., 2013).

355

356 *Quantification of RAD51 protein levels per cell*

357

Next, we aimed to investigate the relationship between RAD51 mRNA and protein levels per cell to assess the extent to which the rise in mRNA numbers aligns with the resultant protein quantity. For that we performed WM-smFISH on RAD51-GFP line (Da Ines et al., 2013) and quantified mRNA and protein mean fluorescence intensity per cell as described previously (Fig. 3a) (Zhao et al., 2023). Similarly to Col-0 plants, RAD51 mRNA levels per cell exhibited an increase with increasing zeocin concentration in RAD51-GFP line (Fig. 3a(ii, iv); Fig. 3b). The results argue again in favor of linear growth of RAD51 transcriptional response. However,

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generated to visualize number of transcripts per cell across root tissues (Fig. 2b). The results
revealed that the number of transcribing cells as well as the number of RAD51 mRNA
molecules detected per cell increases in response to increasing zeocin concentrations. This
pattern is further evident in the histogram quantification (Fig. 2c), depicting a progressive rise
in the number of actively transcribing cells with increasing zeocin concentrations. This data
also indicates a possible upper boundary on the number of RAD51 mRNA molecules per cell.
Indeed, one cannot observe a large difference in mRNA numbers per cell between 50 µM and
170 µM zeocin concentrations despite the large increase in concentration, as visualized on
heatmaps (Fig. 2b) and in a graph form (Fig. 2c, d).

Importantly, our results indicate substantial variability among root cells in their sensitivity to DSBs induced by zeocin, as revealed by the non-uniform heatmaps. Some cells exhibited a strong transcriptional response even at a 10 µM zeocin concentration, with mRNA counts comparable to those induced by 50 µM and 170 µM (Fig. 2b, c). Conversely, certain cells displayed low mRNA counts even after exposure to 50 µM and 170 µM zeocin (Fig. 2b, c). To discern potential distinctions between cell types, we plotted the number of RAD51 mRNAs in different root cell types (Epidermis, Cortex, Endodermis, and Stele). The results demonstrated that RAD51 transcriptional response within stele cells was distinct from the other root cell types analyzed showing higher per cell mRNA output (Fig. 2d). The comparison also indicated no difference in mRNA counts per cell between 50 µM and 170 µM zeocin treated samples (Fig. 2d), potentially arguing in preference of previously proposed limit to per cell transcript output (Fig. S1a). Notably, across developmental regions RAD51 transcriptional output decreased in the elongation zone in comparison with the meristem region (Fig. S3) consistent with previous reports (Da Ines et al., 2013).

355

Quantification of RAD51 protein levels per cell
---

357

Next, we aimed to investigate the relationship between RAD51 mRNA and protein levels per cell to assess the extent to which the rise in mRNA numbers aligns with the resultant protein quantity. For that we performed WM-smFISH on RAD51-GFP line (Da Ines et al., 2013) and quantified mRNA and protein mean fluorescence intensity per cell as described previously (Fig. 3a) (Zhao et al., 2023). Similarly to Col-0 plants, RAD51 mRNA levels per cell exhibited an increase with increasing zeocin concentration in RAD51-GFP line (Fig. 3a(ii, iv); Fig. 3b). The results argue again in favor of linear growth of RAD51 transcriptional response. However,

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For that we performed WM-smFISH on RAD51-GFP line (Da Ines et al., 2013) and
quantified mRNA and protein mean fluorescence intensity per cell as described previously
(Fig. 3a) (Zhao et al., 2023). Similarly to Col-0 plants, RAD51 mRNA levels per cell exhibited
an increase with increasing zeocin concentration in RAD51-GFP line (Fig. 3a(ii, iv); Fig. 3b).
The results argue again in favor of linear growth of RAD51 transcriptional response. However,

365
we still observed that the increase in fluorescence was not isometric to the growth in zeocin concentration and substantial number of values between 50 µM and 170 µM zeocin treated samples were overlapping (Fig. 3b). The growth trend was similar for RAD51-GFP protein (Fig. 3a(iii, v); Fig. 3c). Of note, measured fluorescence signal intensity did not increase further after 50 µM zeocin concentration, suggesting a limit to RAD51 protein amount that can be present in the cell (Fig. 3c). Intriguingly, heatmaps evaluating ratio between mRNA and protein levels revealed slight differences between cells in terms of mRNA and protein accumulation (Fig. 3a(vi); Fig. S4-6). In line with our previous observations, mRNA molecules seem to have a higher abundance in stele (Fig. 3a(ii, iv), a(vi); Fig. S4-6). RAD51-GFP protein accumulation, on the other hand, was more prevalent in the cortex and epidermis of the root tip (Fig. 3a(iii, v), a(vi); Fig. S3-5). This differential accumulation between mRNA and protein among different cell types is intriguing and could suggest protein movement or differential degradation between cells but more investigation to validate these hypotheses would be required.

## RAD51 transcription through the cell cycle

RAD51 transcription is typically linked to the S and G2 phases of the cell cycle, motivated by the requirement of homologous DNA sequences during repair through HR (Schuermann *et al.*, 2005; Goldfarb & Lichten, 2010). Given the very high proportion of cells with RAD51 mRNAs signals in the zeocin-treated samples, we expected a considerable number of cells arrested at the S or G2/M checkpoints (Osakabe *et al.*, 2005; De Schutter *et al.*, 2007). To evaluate the cell cycle arrest in roots treated with zeocin, we conducted EdU staining to label cells that went through S-phase in a sequential smFISH/EdU protocol (Fig. 4a-c). Our results revealed a drastic decline in the number of EdU-positive cells with increasing zeocin concentration, with almost no labeled cells at 50 µM and 170 µM concentrations, indicating a strong cell cycle arrest in these samples (Fig. 4b, 4d, Fig. S7). EdU-positive cells tend to be most abundant in the root stele, possibly explaining higher RAD51 transcript output in this part of the root as it is usually associated with S/G2 phases of the cell cycle (Fig. S7a, S7b, S7c). Moreover, two-way ANOVA revealed that the observed variations in EdU-positive cell numbers can be explained by both zeocin concentration and the cell type with significant interaction between the two parameters ($p = 4.94 \times 10^{-15}$) (Fig. S7b). Subsequent pairwise comparisons revealed statistically significant changes in cell numbers between the concentrations only in the root stele. Importantly, comparing EdU labelling with RAD51 smFISH signals revealed EdU
we still observed that the increase in fluorescence was not isometric to the growth in zeocin concentration and substantial number of values between 50 µM and 170 µM zeocin treated samples were overlapping (Fig. 3b). The growth trend was similar for RAD51-GFP protein (Fig. 3a(iii, v); Fig. 3c). Of note, measured fluorescence signal intensity did not increase further after 50 µM zeocin concentration, suggesting a limit to RAD51 protein amount that can be present in the cell (Fig. 3c). Intriguingly, heatmaps evaluating ratio between mRNA and protein levels revealed slight differences between cells in terms of mRNA and protein accumulation (Fig. 3a(vi); Fig. S4-6). In line with our previous observations, mRNA molecules seem to have a higher abundance in stele (Fig. 3a(ii, iv), a(vi); Fig. S4-6). RAD51-GFP protein accumulation, on the other hand, was more prevalent in the cortex and epidermis of the root tip (Fig. 3a(iii, v), a(vi); Fig. S3-5). This differential accumulation between mRNA and protein among different cell types is intriguing and could suggest protein movement or differential degradation between cells but more investigation to validate these hypotheses would be required.

## RAD51 transcription through the cell cycle

RAD51 transcription is typically linked to the S and G2 phases of the cell cycle, motivated by the requirement of homologous DNA sequences during repair through HR (Schuermann et al., 2005; Goldfarb & Lichten, 2010). Given the very high proportion of cells with RAD51 mRNAs signals in the zeocin-treated samples, we expected a considerable number of cells arrested at the S or G2/M checkpoints (Osakabe et al., 2005; De Schutter et al., 2007). To evaluate the cell cycle arrest in roots treated with zeocin, we conducted EdU staining to label cells that went through S-phase in a sequential smFISH/EdU protocol (Fig. 4a-c). Our results revealed a drastic decline in the number of EdU-positive cells with increasing zeocin concentration, with almost no labeled cells at 50 µM and 170 µM concentrations, indicating a strong cell cycle arrest in these samples (Fig. 4b, 4d, Fig. S7). EdU-positive cells tend to be most abundant in the root stele, possibly explaining higher RAD51 transcript output in this part of the root as it is usually associated with S/G2 phases of the cell cycle (Fig. S7a, S7b, S7c). Moreover, two-way ANOVA revealed that the observed variations in EdU-positive cell numbers can be explained by both zeocin concentration and the cell type with significant interaction between the two parameters ($p = 4.94 \times 10^{-15}$) (Fig. S7b). Subsequent pairwise comparisons revealed statistically significant changes in cell numbers between the concentrations only in the root stele. Importantly, comparing EdU labelling with RAD51 smFISH signals revealed EdU
stained cells with modest RAD51 mRNA presence next to cells with no EdU signal and
abundant number of RAD51 mRNA molecules (Fig. 4c). This observation potentially
challenges the strict dependency of RAD51 transcription on the S/G2-phase of the cells. Of
note, EdU signals were observed in the elongation zone of the root at 50 µM and 170 µM
concentrations of zeocin, revealing distinct responses across the various root developmental
zones (Fig. S7c).

To further investigate the association between RAD51 transcription and S/G2 phases of the cell
cycle under damage, we used Cytrap (Yin et al., 2014), CDT1-CFP (Desvoyes et al., 2019)
and PlaCCI (Desvoyes et al., 2020) lines, which express fluorescent reporters specific to
individual cell cycle phases. Cytrap line allows visualization of S/G2 phase cells and G2/M
cells while PlaCCI line provides additional possibility of direct G1 phase cells visualization
using CDT1a-CFP construct which is also available as a separate line (Fig. 4e). Analysis of
Cytrap line revealed a decrease in S/G2-phase cells with increasing concentrations of zeocin,
consistent with EdU staining data (Fig. S8a), as well as an increase in the fraction of cells
expressing G2/M reporter, potentially corresponding to checkpoint arrest (Fig. S8b) (Preuss &
Britt, 2003). Statistical analysis revealed that the changes in S/G2 and G2/M phase cells can be
explained by both zeocin concentration and the cell type with significant interaction between
the two parameters (p = 1.08e-09, p = 6.56e-06 accordingly) (Fig. S8e-f). Further pairwise
comparisons revealed statistically significant changes between the concentrations only within
root stele group, an observation strikingly similar to the earlier reported RAD51 transcriptional
data. Of note, the combined percentage of S/G2 and G2/M cells at higher concentrations of
zeocin suggests that a large fraction of cells is not either in S phase or at G2/M checkpoint,
thus potentially residing in G1 phase (Fig. 4f). Considering the novelty of this finding we
decided to rely on direct visualization of G1 phase cells using recently developed PlaCCI and
CDT1-CFP lines. The results confirmed an increase in the amount of G1 cells in response to
increasing concentrations of zeocin in PlaCCI and CDT1-CFP lines indicating potential cell
cycle arrest at G1/S checkpoint (Fig. 4g, Fig. S9a). The proportion of cells with G2/M marker
also increased confirming the results obtained with Cytrap line (Fig. S9b, S9c). Analysis of the
results showed the changes in G1 cell numbers can be explained by both zeocin concentration
and the cell type with significant interaction between the two parameters (p = 0.0268) (Fig.
S9a, S9d). Further pairwise comparisons revealed statistically significant increase in G1 cells
only within root stele group.
stained cells with modest RAD51 mRNA presence next to cells with no EdU signal and
abundant number of RAD51 mRNA molecules (Fig. 4c). This observation potentially
challenges the strict dependency of RAD51 transcription on the S/G2-phase of the cells. Of
note, EdU signals were observed in the elongation zone of the root at 50 µM and 170 µM
concentrations of zeocin, revealing distinct responses across the various root developmental
zones (Fig. S7c).

To further investigate the association between RAD51 transcription and S/G2 phases of the cell
cycle under damage, we used Cytrap (Yin et al., 2014), CDT1-CFP (Desvoyes et al., 2019)
and PlaCCI (Desvoyes et al., 2020) lines, which express fluorescent reporters specific to
individual cell cycle phases. Cytrap line allows visualization of S/G2 phase cells and G2/M
cells while PlaCCI line provides additional possibility of direct G1 phase cells visualization
using CDT1a-CFP construct which is also available as a separate line (Fig. 4e). Analysis of
Cytrap line revealed a decrease in S/G2-phase cells with increasing concentrations of zeocin,
consistent with EdU staining data (Fig. S8a), as well as an increase in the fraction of cells
expressing G2/M reporter, potentially corresponding to checkpoint arrest (Fig. S8b) (Preuss &
Britt, 2003). Statistical analysis revealed that the changes in S/G2 and G2/M phase cells can be
explained by both zeocin concentration and the cell type with significant interaction between
the two parameters (p = 1.08e-09, p = 6.56e-06 accordingly) (Fig. S8e-f). Further pairwise
comparisons revealed statistically significant changes between the concentrations only within
root stele group, an observation strikingly similar to the earlier reported RAD51 transcriptional
data. Of note, the combined percentage of S/G2 and G2/M cells at higher concentrations of
zeocin suggests that a large fraction of cells is not either in S phase or at G2/M checkpoint,
thus potentially residing in G1 phase (Fig. 4f). Considering the novelty of this finding we
decided to rely on direct visualization of G1 phase cells using recently developed PlaCCI and
CDT1-CFP lines. The results confirmed an increase in the amount of G1 cells in response to
increasing concentrations of zeocin in PlaCCI and CDT1-CFP lines indicating potential cell
cycle arrest at G1/S checkpoint (Fig. 4g, Fig. S9a). The proportion of cells with G2/M marker
also increased confirming the results obtained with Cytrap line (Fig. S9b, S9c). Analysis of the
results showed the changes in G1 cell numbers can be explained by both zeocin concentration
and the cell type with significant interaction between the two parameters (p = 0.0268) (Fig.
S9a, S9d). Further pairwise comparisons revealed statistically significant increase in G1 cells
only within root stele group.
429
430

Intriguingly, we also observed a small fraction of cells without any fluorescent reporter presence in roots of PlaCCI line (Fig. S9c).

431
432
433
434
435
436
437
438
439
440
441

One possible explanation for *RAD51* transcripts observed in G1 cells is that they could be produced in S/G2 and carried over to G1 due to a potentially long half-life of transcripts. To evaluate the mRNA half-life, we treated seedlings with the transcription elongation inhibitor, actinomycin D (ActD), and conducted a time-series smFISH analysis (Fig. S10). The half-life of the *RAD51* mRNA was calculated from our data as 4.9 hours. Considering this measurement there is the possibility of *RAD51* mRNA persisting beyond the G2-phase of the cell cycle. Indeed, we detected mitotic cells, which normally do not actively transcribe genes, possessing *RAD51* mRNAs (Fig. S10c). However, this half-life (4.9h) is relatively short compared to the cell cycle duration (Rahni & Birnbaum, 2019) so while *RAD51* mRNA may be carried between cell divisions, its half-life alone seems unlikely to explain the high proportion of cells with *RAD51* mRNA signals in zeocin samples.

442
443
444
445
446
447
448
449
450
451
452
453
454
455
456

To show *RAD51* transcription in G1 arrested cells directly, we performed *RAD51* smFISH detection on CDT1-CFP line, expressing the same G1 marker as PlaCCI line alone (Fig. 4h). The results clearly show the presence of multiple *RAD51* mRNAs and most importantly active transcriptional sites, as judged by the presence of larger smFISH foci in the nucleus (Fig. 4h(ii)), in cells labeled with G1 phase reporter (Fig. 4h(iii)), thereby directly confirming predicted *RAD51* transcription during G1 phase under DNA damage. Upon further examination of transcription site numbers in G1 and non-G1 cells using the CDT1-CFP line, we observed a nearly equal partitioning (Supplementary table 1). This observation implies that *RAD51* transcription occurs with approximately equal probability during both G1 and other stages of the cell cycle under conditions of DNA damage. This result correlates with similar numbers of cells residing in G1 and other cell cycle phases (Fig. 4f, S9c). Intriguingly, some G1 *RAD51* transcription was possible even under control conditions.

454
455
456

DISCUSSION

457
458
459
460

This study describes the transcriptional activation of *RAD51* following increasing amounts of DNA damage. Our findings indicate a rise in total mRNA production that results from an increase in transcriptional output per cell as DNA damage increases. These results underscore the cell's capacity to sense the extent of damage and modulate *RAD51* transcription accordingly.
429
430

Intriguingly, we also observed a small fraction of cells without any fluorescent reporter presence in roots of PlaCCI line (Fig. S9c).

431
432
433
434
435
436
437
438
439
440
441

One possible explanation for *RAD51* transcripts observed in G1 cells is that they could be produced in S/G2 and carried over to G1 due to a potentially long half-life of transcripts. To evaluate the mRNA half-life, we treated seedlings with the transcription elongation inhibitor, actinomycin D (ActD), and conducted a time-series smFISH analysis (**Fig. S10**). The half-life of the *RAD51* mRNA was calculated from our data as 4.9 hours. Considering this measurement there is the possibility of *RAD51* mRNA persisting beyond the G2-phase of the cell cycle. Indeed, we detected mitotic cells, which normally do not actively transcribe genes, possessing *RAD51* mRNAs (**Fig. S10c**). However, this half-life (4.9h) is relatively short compared to the cell cycle duration (Rahni & Birnbaum, 2019) so while *RAD51* mRNA may be carried between cell divisions, its half-life alone seems unlikely to explain the high proportion of cells with *RAD51* mRNA signals in zeocin samples.

442
443
444
445
446
447
448
449
450
451
452
453
454
455
456

To show *RAD51* transcription in G1 arrested cells directly, we performed *RAD51* smFISH detection on CDT1-CFP line, expressing the same G1 marker as PlaCCI line alone (Fig. 4h). The results clearly show the presence of multiple *RAD51* mRNAs and most importantly active transcriptional sites, as judged by the presence of larger smFISH foci in the nucleus (Fig. 4h(ii)), in cells labeled with G1 phase reporter (Fig. 4h(iii)), thereby directly confirming predicted *RAD51* transcription during G1 phase under DNA damage. Upon further examination of transcription site numbers in G1 and non-G1 cells using the CDT1-CFP line, we observed a nearly equal partitioning (Supplementary table 1). This observation implies that *RAD51* transcription occurs with approximately equal probability during both G1 and other stages of the cell cycle under conditions of DNA damage. This result correlates with similar numbers of cells residing in G1 and other cell cycle phases (Fig. 4f, S9c). Intriguingly, some G1 *RAD51* transcription was possible even under control conditions.

454
455
456

DISCUSSION

457
458
459
460

This study describes the transcriptional activation of *RAD51* following increasing amounts of DNA damage. Our findings indicate a rise in total mRNA production that results from an increase in transcriptional output per cell as DNA damage increases. These results underscore the cell's capacity to sense the extent of damage and modulate *RAD51* transcription accordingly.
461 Using single-cell measurements by smFISH technique we obtained data showing differences in
462 DNA damage sensitivity between cells, manifested by varying RAD51 mRNA transcriptional
463 output in response to the same concentration of DNA damaging agent. Our results also revealed
464 dynamic changes in numbers of cells residing in different cell cycle phases in response to
465 increased DNA damage. Of note, these changes were not as evident between samples treated
466 with 50 µM and 170 µM zeocin concentrations, for which the number of RAD51 mRNAs per
467 cell did not differ substantially. One can therefore speculate that proportions of cell arrested at
468 different cell cycle phases achieved in 50 µM and 170 µM zeocin treated samples are the most
469 efficient for cells to cope with DNA damage.

470

471 Cell cycle arrest and upregulation of DDR genes are the two key elements of the DDR response.
472 Our data from several independent experiments showed that root stele cells consistently
differed from other cell types in both cell cycle changes and RAD51 transcriptional response
473 to growing amounts of DNA damage. Specifically, root stele cells exhibited a more extensive
474 RAD51 transcriptional activation as well as larger fluctuations in numbers of cells represented
475 at different cell cycle stages under the same zeocin concentrations. One potential explanation
476 for this observation could be linked to distinct cell cycle duration among the different cell types.
477 Live-imaging experiments indicate a shorter cell cycle duration for stele cells (~15 hours)
478 compared to other root cell types (~23 hours for cortex and 24h for the epidermis) (Rahni &
479 Birnbaum, 2019). Faster proliferation rates have been correlated with increased susceptibility
480 to DNA damage (Kiraly et al., 2015; Alhmoud et al., 2020). Notably, stele cells have
481 demonstrated higher sensitivity to cell death in response to zeocin (Yoshiyama et al., 2017;
482 Johnson et al., 2018; Ryu et al., 2019). Consequently, the greater accumulation of damage may
483 underscore the elevated transcriptional response of RAD51 in stele cells.
484

485

486 The observation of RAD51 transcription occurring outside S/G2 phases of the cell cycle is
487 another important finding of this study. DDR via HR and RAD51 gene expression has been
associated with S/G2-phase of the cell cycle in many organisms (Basile et al., 1992; Yamamoto
488 et al., 1996; Doutriaux et al., 1998). In A. thaliana RAD51 transcription in response to DNA
damage was coincident with the cell cycle arrest at G2/M checkpoint (Osakabe et al., 2005; De
489 Schutter et al., 2007). Later studies demonstrated that both G1/S and G2/M checkpoints can be
490 used to ensure cell cycle arrest in response to DNA damage. For example, hydroxyurea (HU)
491 treatment was shown to activate both G2/M cell cycle arrest (De Schutter et al., 2007) and G1/S
492 checkpoint (Saban & Bujak, 2009; Cabral et al., 2020), a phenomenon also observed in
493
494
Using single-cell measurements by smFISH technique we obtained data showing differences in DNA damage sensitivity between cells, manifested by varying RAD51 mRNA transcriptional output in response to the same concentration of DNA damaging agent. Our results also revealed dynamic changes in numbers of cells residing in different cell cycle phases in response to increased DNA damage. Of note, these changes were not as evident between samples treated with 50 µM and 170 µM zeocin concentrations, for which the number of RAD51 mRNAs per cell did not differ substantially. One can therefore speculate that proportions of cell arrested at different cell cycle phases achieved in 50 µM and 170 µM zeocin treated samples are the most efficient for cells to cope with DNA damage.

Cell cycle arrest and upregulation of DDR genes are the two key elements of the DDR response. Our data from several independent experiments showed that root stele cells consistently differed from other cell types in both cell cycle changes and RAD51 transcriptional response to growing amounts of DNA damage. Specifically, root stele cells exhibited a more extensive RAD51 transcriptional activation as well as larger fluctuations in numbers of cells represented at different cell cycle stages under the same zeocin concentrations. One potential explanation for this observation could be linked to distinct cell cycle duration among the different cell types. Live-imaging experiments indicate a shorter cell cycle duration for stele cells (~15 hours) compared to other root cell types (~23 hours for cortex and 24h for the epidermis) (Rahni & Birnbaum, 2019). Faster proliferation rates have been correlated with increased susceptibility to DNA damage (Kiraly et al., 2015; Alhmoud et al., 2020). Notably, stele cells have demonstrated higher sensitivity to cell death in response to zeocin (Yoshiyama et al., 2017; Johnson et al., 2018; Ryu et al., 2019). Consequently, the greater accumulation of damage may underscore the elevated transcriptional response of RAD51 in stele cells.

The observation of RAD51 transcription occurring outside S/G2 phases of the cell cycle is another important finding of this study. DDR via HR and RAD51 gene expression has been associated with S/G2-phase of the cell cycle in many organisms (Basile et al., 1992; Yamamoto et al., 1996; Doutriaux et al., 1998). In A. thaliana RAD51 transcription in response to DNA damage was coincident with the cell cycle arrest at G2/M checkpoint (Osakabe et al., 2005; De Schutter et al., 2007). Later studies demonstrated that both G1/S and G2/M checkpoints can be used to ensure cell cycle arrest in response to DNA damage. For example, hydroxyurea (HU) treatment was shown to activate both G2/M cell cycle arrest (De Schutter et al., 2007) and G1/S checkpoint (Saban & Bujak, 2009; Cabral et al., 2020), a phenomenon also observed in
response to gamma irradiation (Hefner, 2003; Hefner et al., 2006; Ricaud et al., 2007). Zeocin, the radiomimetic drug used in this study to induce DSBs, was so far reported to promote arrest at the G2/M checkpoint (De Schutter et al., 2007; Chen et al., 2017). Our findings challenge this view by demonstrating that a considerable number of cells undergo arrest in the G1 phase while still exhibiting RAD51 transcription. This observation does not mean that RAD51 is not transcribed in S/G2. Indeed, our data indicated equal representation of transcription site signals in G1 cells and cells in the rest of the cell cycle, potentially indicating absence of preference for RAD51 transcription between the cell cycle phases under varying amounts of DNA damage. Our results therefore suggest RAD51 transcription being more widespread across the cell cycle than initially anticipated.

Previous studies suggest a potential reason and implication behind the release of the S/G2 restriction of RAD51 expression. For instance, it was shown that repetitive sequences can be repaired via HR during G1 phase, proven by the recruitment of RAD51 to centromeric break sites in mouse and human cells (Yilmaz et al., 2021). The HR machinery is also involved in G1 repair of ribosomal DNA, another type of repetitive sequence in human cell cultures (van Sluis & McStay, 2015). Moreover, non-recombinogenic functions in DNA reparation were suggested for RAD51 and some HR proteins (Cano-Linares et al., 2021; Prado, 2021). We suggest this as one of the potential reasons behind our observation of active RAD51 transcription in G1 after DNA damage exposure. Also, in A. thaliana, RAD54 foci were shown to emerge with high frequency in both in G1 and G2 cells after gamma irradiation (Hirakawa & Matsunaga, 2019). The necessity of prior RAD51 foci formation for the formation of RAD54 foci points to the possibility of RAD51 foci presence in G1 phase A. thaliana cells (Hirakawa et al., 2017).

Altogether, the results of this article shed new light on the DNA damage response in plants, uncovering distinctions in the transcriptional response of RAD51 across various cell types. Moreover, it highlights the noteworthy occurrence of transcription during the G1 phase of the cell cycle.

## ACKNOWLEDGEMENTS

We would like to acknowledge Lihua Zhao for advice on whole mount smFISH protocol performance. We are grateful to Benedicte Desvoyes and Crisanto Gutierrez for providing seeds of PlaCCI and CDT1-CFP lines and critical reading of the manuscript. This work was
response to gamma irradiation (Hefner, 2003; Hefner et al., 2006; Ricaud et al., 2007). Zeocin, the radiomimetic drug used in this study to induce DSBs, was so far reported to promote arrest at the G2/M checkpoint (De Schutter et al., 2007; Chen et al., 2017). Our findings challenge this view by demonstrating that a considerable number of cells undergo arrest in the G1 phase while still exhibiting RAD51 transcription. This observation does not mean that RAD51 is not transcribed in S/G2. Indeed, our data indicated equal representation of transcription site signals in G1 cells and cells in the rest of the cell cycle, potentially indicating absence of preference for RAD51 transcription between the cell cycle phases under varying amounts of DNA damage. Our results therefore suggest RAD51 transcription being more widespread across the cell cycle than initially anticipated.

Previous studies suggest a potential reason and implication behind the release of the S/G2 restriction of RAD51 expression. For instance, it was shown that repetitive sequences can be repaired via HR during G1 phase, proven by the recruitment of RAD51 to centromeric break sites in mouse and human cells (Yilmaz et al., 2021). The HR machinery is also involved in G1 repair of ribosomal DNA, another type of repetitive sequence in human cell cultures (van Sluis & McStay, 2015). Moreover, non-recombinogenic functions in DNA reparation were suggested for RAD51 and some HR proteins (Cano-Linares et al., 2021; Prado, 2021). We suggest this as one of the potential reasons behind our observation of active RAD51 transcription in G1 after DNA damage exposure. Also, in A. thaliana, RAD54 foci were shown to emerge with high frequency in both in G1 and G2 cells after gamma irradiation (Hirakawa & Matsunaga, 2019). The necessity of prior RAD51 foci formation for the formation of RAD54 foci points to the possibility of RAD51 foci presence in G1 phase A. thaliana cells (Hirakawa et al., 2017).

Altogether, the results of this article shed new light on the DNA damage response in plants, uncovering distinctions in the transcriptional response of RAD51 across various cell types. Moreover, it highlights the noteworthy occurrence of transcription during the G1 phase of the cell cycle.

## ACKNOWLEDGEMENTS

We would like to acknowledge Lihua Zhao for advice on whole mount smFISH protocol performance. We are grateful to Benedicte Desvoyes and Crisanto Gutierrez for providing seeds of PlaCCI and CDT1-CFP lines and critical reading of the manuscript. This work was
528

supported by Swedish Research Council (Vetenskapsrådet) grant number 2018-04101 to SR;

529

Knut and Alice Wallenberg Foundation (KAW 2019-0062).

530

COMPETING INTERESTS

531

532

None declared.

533

534

AUTHOR CONTRIBUTIONS

535

536

KK, SvR, AM, AS and SR designed the research. KK performed the research and data analysis.

537

AF assisted data analysis and created software pipelines for image analysis. KS assisted

538

statistical analysis of the data. KK and SR wrote the manuscript. CW provided material for the

539

study. CW and AS critically read the manuscript.

540

541

DATA AVAILABILITY

542

543

The data from this study are not currently deposited in external repositories. However, they can

544

be requested directly from the corresponding author. Upon acceptance, the data will be made

545

available in repositories.

546

547

REFERENCES

548

549

Abe K, Osakabe K, Nakayama S, Endo M, Tagiri A, Todoriki S, Ichikawa H, Toki S. 2005.

550

Arabidopsis RAD51C gene is important for homologous recombination in meiosis and mitosis.

551

*Plant Physiology* **139**: 896–908.

552

Adachi S, Minamisawa K, Okushima Y, Inagaki S, Yoshiyama K, Kondou Y, Kaminuma

553

E, Kawashima M, Toyoda T, Matsui M, et al. 2011. Programmed induction of

554

endoreduplication by DNA double-strand breaks in *Arabidopsis*. *Proceedings of the National*

555

*Academy of Sciences* **108**: 10004–10009.

556

Aki SS, Umeda M. 2016. Cytrap Marker Systems for In Vivo Visualization of Cell Cycle

557

Progression in Arabidopsis. In: Caillaud M-C, ed. Methods in Molecular Biology. Plant Cell

558

Division. New York, NY: Springer New York, 51–57.
528

supported by Swedish Research Council (Vetenskapsrådet) grant number 2018-04101 to SR;

529

Knut and Alice Wallenberg Foundation (KAW 2019-0062).

530

COMPETING INTERESTS

531

532

None declared.

533

534

AUTHOR CONTRIBUTIONS

535

536

KK, SvR, AM, AS and SR designed the research. KK performed the research and data analysis.

537

AF assisted data analysis and created software pipelines for image analysis. KS assisted

538

statistical analysis of the data. KK and SR wrote the manuscript. CW provided material for the

539

study. CW and AS critically read the manuscript.

540

541

DATA AVAILABILITY

542

543

The data from this study are not currently deposited in external repositories. However, they can

544

be requested directly from the corresponding author. Upon acceptance, the data will be made

545

available in repositories.

546

547

REFERENCES

548

549

Abe K, Osakabe K, Nakayama S, Endo M, Tagiri A, Todoriki S, Ichikawa H, Toki S. 2005.

550

Arabidopsis RAD51C gene is important for homologous recombination in meiosis and mitosis.

551

*Plant Physiology* **139**: 896–908.

552

Adachi S, Minamisawa K, Okushima Y, Inagaki S, Yoshiyama K, Kondou Y, Kaminuma

553

E, Kawashima M, Toyoda T, Matsui M, et al. 2011. Programmed induction of

554

endoreduplication by DNA double-strand breaks in *Arabidopsis*. *Proceedings of the National*

555

*Academy of Sciences* **108**: 10004–10009.

556

Aki SS, Umeda M. 2016. Cytrap Marker Systems for In Vivo Visualization of Cell Cycle

557

Progression in Arabidopsis. In: Caillaud M-C, ed. Methods in Molecular Biology. Plant Cell

558

Division. New York, NY: Springer New York, 51–57.
559

Alhmoud JF, Woolley JF, Al Moustafa A-E, Malki MI. 2020. DNA Damage/Repair Management in Cancers. *Cancers* 12: 1050.

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Banerjee S, Roy S. 2021. An insight into understanding the coupling between homologous recombination mediated DNA repair and chromatin remodeling mechanisms in plant genome: an update. *Cell Cycle* (Georgetown, Tex.) 20: 1760–1784.

563

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Basile G, Aker M, Mortimer RK. 1992. Nucleotide sequence and transcriptional regulation of the yeast recombinational repair gene RAD51. *Molecular and Cellular Biology* 12: 3235–3246.

565

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567

Bee L, Fabris S, Cherubini R, Mognato M, Celotti L. 2013. The Efficiency of Homologous Recombination and Non-Homologous End Joining Systems in Repairing Double-Strand Breaks during Cell Cycle Progression (S Cotterill, Ed.). *PLoS ONE* 8: e69061.

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Biedermann S, Harashima H, Chen P, Heese M, Bouyer D, Sofroni K, Schnittger A. 2017. The retinoblastoma homolog RBR 1 mediates localization of the repair protein RAD 51 to DNA lesions in *Arabidopsis*. *The EMBO Journal* 36: 1279–1297.

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Bonilla B, Hengel SR, Grundy MK, Bernstein KA. 2020. RAD51 Gene Family Structure and Function. *Annual Review of Genetics* 54: 25–46.

574

575

Cabral D, Banora MY, Antonino JD, Rodiuc N, Vieira P, Coelho RR, Chevalier C, Eekhout T, Engler G, De Veylder L, et al. 2020. The plant WEE1 kinase is involved in checkpoint control activation in nematode-induced galls. *New Phytologist* 225: 430–447.

576

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Cano-Linares MI, Yáñez-Vilches A, García-Rodríguez N, Barrientos-Moreno M, González-Prieto R, San-Segundo P, Ulrich HD, Prado F. 2021. Non-recombinogenic roles for Rad52 in translesion synthesis during DNA damage tolerance. *EMBO reports* 22: e50410.

579

580

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Chabot T, Defontaine A, Marquis D, Renodon-Corniere A, Courtois E, Fleury F, Cheraud Y. 2019. New Phosphorylation Sites of Rad51 by c-Met Modulates Presynaptic Filament Stability. *Cancers* 11: 413.

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Charbonnel C, Gallego ME, White CI. 2010. Xrcc1-dependent and Ku-dependent DNA double-strand break repair kinetics in Arabidopsis plants: Double-strand break repair kinetics in Arabidopsis. *The Plant Journal* 64: 280–290.

585

586
559

Alhmoud JF, Woolley JF, Al Moustafa A-E, Malki MI. 2020. DNA Damage/Repair Management in Cancers. *Cancers* 12: 1050.

560

561

Banerjee S, Roy S. 2021. An insight into understanding the coupling between homologous recombination mediated DNA repair and chromatin remodeling mechanisms in plant genome: an update. *Cell Cycle* (Georgetown, Tex.) 20: 1760–1784.

563

564

Basile G, Aker M, Mortimer RK. 1992. Nucleotide sequence and transcriptional regulation of the yeast recombinational repair gene RAD51. *Molecular and Cellular Biology* 12: 3235–3246.

565

566

567

Bee L, Fabris S, Cherubini R, Mognato M, Celotti L. 2013. The Efficiency of Homologous Recombination and Non-Homologous End Joining Systems in Repairing Double-Strand Breaks during Cell Cycle Progression (S Cotterill, Ed.). *PLoS ONE* 8: e69061.

568

569

570

Biedermann S, Harashima H, Chen P, Heese M, Bouyer D, Sofroni K, Schnittger A. 2017. The retinoblastoma homolog RBR 1 mediates localization of the repair protein RAD 51 to DNA lesions in *Arabidopsis*. *The EMBO Journal* 36: 1279–1297.

571

572

573

Bonilla B, Hengel SR, Grundy MK, Bernstein KA. 2020. RAD51 Gene Family Structure and Function. *Annual Review of Genetics* 54: 25–46.

574

575

Cabral D, Banora MY, Antonino JD, Rodiuc N, Vieira P, Coelho RR, Chevalier C, Eekhout T, Engler G, De Veylder L, et al. 2020. The plant WEE1 kinase is involved in checkpoint control activation in nematode-induced galls. *New Phytologist* 225: 430–447.

576

577

578

Cano-Linares MI, Yáñez-Vilches A, García-Rodríguez N, Barrientos-Moreno M, González-Prieto R, San-Segundo P, Ulrich HD, Prado F. 2021. Non-recombinogenic roles for Rad52 in translesion synthesis during DNA damage tolerance. *EMBO reports* 22: e50410.

579

580

581

Chabot T, Defontaine A, Marquis D, Renodon-Corniere A, Courtois E, Fleury F, Cheraud Y. 2019. New Phosphorylation Sites of Rad51 by c-Met Modulates Presynaptic Filament Stability. *Cancers* 11: 413.

582

583

584

Charbonnel C, Gallego ME, White CI. 2010. Xrcc1-dependent and Ku-dependent DNA double-strand break repair kinetics in Arabidopsis plants: Double-strand break repair kinetics in Arabidopsis. *The Plant Journal* 64: 280–290.

585

586
587 Chen P, Takatsuka H, Takahashi N, Kurata R, Fukao Y, Kobayashi K, Ito M, Umeda M.
588 2017. Arabidopsis R1R2R3-Myb proteins are essential for inhibiting cell division in response
589 to DNA damage. *Nature Communications* **8**: 635.

590 Coïc E, Martin J, Ryu T, Tay SY, Kondev J, Haber JE. 2011. Dynamics of Homology
591 Searching During Gene Conversion in *Saccharomyces cerevisiae* Revealed by Donor
592 Competition. *Genetics* **189**: 1225-1233.

593 Cools T, Iantcheva A, Weimer AK, Boens S, Takahashi N, Maes S, Van den Daele H, Van
594 Isterdael G, Schnittger A, De Veylder L. 2011. The *Arabidopsis thaliana* Checkpoint Kinase
595 WEE1 Protects against Premature Vascular Differentiation during Replication Stress. *The Plant
596 Cell* **23**: 1435-1448.

597 Cui W, Wang H, Song J, Cao X, Rogers HJ, Francis D, Jia C, Sun L, Hou M, Yang Y, et
598 al. 2017. Cell cycle arrest mediated by Cd-induced DNA damage in Arabidopsis root tips.
599 *Ecotoxicology and Environmental Safety* **145**: 569-574.

600 Culligan K, Tissier A, Britt A. 2004. ATR Regulates a G2-Phase Cell-Cycle Checkpoint in
601 *Arabidopsis thaliana*. *The Plant Cell* **16**: 1091-1104.

602 Da Ines O, Bazile J, Gallego ME, White CI. 2022. DMC1 attenuates RAD51-mediated
603 recombination in Arabidopsis (IR Henderson, Ed.). *PLOS Genetics* **18**: e1010322.

604 Da Ines O, Degroote F, Goubely C, Amiard S, Gallego ME, White CI. 2013. Meiotic
605 Recombination in Arabidopsis Is Catalysed by DMC1, with RAD51 Playing a Supporting Role
606 (FCH Franklin, Ed.). *PLoS Genetics* **9**: e1003787.

607 De Schutter K, Joubès J, Cools T, Verkest A, Corellou F, Babiychuk E, Van Der Schueren
608 E, Beeckman T, Kushnir S, Inzé D, et al. 2007. Arabidopsis WEE1 kinase controls cell cycle
609 arrest in response to activation of the DNA integrity checkpoint. *The Plant Cell* **19**: 211-225.

610 Desvoyes B, Arana-Echarri A, Barea MD, Gutierrez C. 2020. A comprehensive fluorescent
611 sensor for spatiotemporal cell cycle analysis in Arabidopsis. *Nature Plants* **6**: 1330-1334.

612 Desvoyes B, Noir S, Masoud K, López MI, Genschik P, Gutierrez C. 2019. *FBL17 targets*
613 *CDT1a* for degradation in early S-phase to prevent *Arabidopsis* genome instability. *Plant
614 Biology.*
587 Chen P, Takatsuka H, Takahashi N, Kurata R, Fukao Y, Kobayashi K, Ito M, Umeda M.
588 2017. Arabidopsis R1R2R3-Myb proteins are essential for inhibiting cell division in response
589 to DNA damage. *Nature Communications* **8**: 635.

590 Coïc E, Martin J, Ryu T, Tay SY, Kondev J, Haber JE. 2011. Dynamics of Homology
591 Searching During Gene Conversion in *Saccharomyces cerevisiae* Revealed by Donor
592 Competition. *Genetics* **189**: 1225-1233.

593 Cools T, Iantcheva A, Weimer AK, Boens S, Takahashi N, Maes S, Van den Daele H, Van
594 Isterdael G, Schnittger A, De Veylder L. 2011. The *Arabidopsis thaliana* Checkpoint Kinase
595 WEE1 Protects against Premature Vascular Differentiation during Replication Stress. *The Plant
596 Cell* **23**: 1435-1448.

597 Cui W, Wang H, Song J, Cao X, Rogers HJ, Francis D, Jia C, Sun L, Hou M, Yang Y, et
598 al. 2017. Cell cycle arrest mediated by Cd-induced DNA damage in Arabidopsis root tips.
599 *Ecotoxicology and Environmental Safety* **145**: 569-574.

600 Culligan K, Tissier A, Britt A. 2004. ATR Regulates a G2-Phase Cell-Cycle Checkpoint in
601 *Arabidopsis thaliana*. *The Plant Cell* **16**: 1091-1104.

602 Da Ines O, Bazile J, Gallego ME, White CI. 2022. DMC1 attenuates RAD51-mediated
603 recombination in Arabidopsis (IR Henderson, Ed.). *PLOS Genetics* **18**: e1010322.

604 Da Ines O, Degroote F, Goubely C, Amiard S, Gallego ME, White CI. 2013. Meiotic
605 Recombination in Arabidopsis Is Catalysed by DMC1, with RAD51 Playing a Supporting Role
606 (FCH Franklin, Ed.). *PLoS Genetics* **9**: e1003787.

607 De Schutter K, Joubès J, Cools T, Verkest A, Corellou F, Babiychuk E, Van Der Schueren
608 E, Beeckman T, Kushnir S, Inzé D, et al. 2007. Arabidopsis WEE1 kinase controls cell cycle
609 arrest in response to activation of the DNA integrity checkpoint. *The Plant Cell* **19**: 211-225.

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Yu C, Hou L, Huang Y, Cui X, Xu S, Wang L, Yan S. 2023. The MULTI-BRCT domain protein DDRM2 promotes the recruitment of RAD51 to DNA damage sites to facilitate homologous recombination. *New Phytologist* **238**: 1073–1084.

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SUPPORTING INFORMATION

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Additional supporting information may be found in the online version of this article.

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**Fig. S1** Evaluation of *PP2A* transcription in response to growing amounts of DNA damage in s-quashed roots of *Arabidopsis thaliana*.

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**Fig. S2** Quantification of RAD51 mRNA molecules in Col-0 *Arabidopsis thaliana* roots exposed to growing amounts of DNA damage using whole-mount smFISH.

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**Fig. S3** Evaluation of RAD51 transcriptional response in meristematic and elongation zones of the *Arabidopsis thaliana* root using whole mount smFISH.

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**Fig. S4** Quantification of *RAD51* mRNA and RAD51-GFP protein signals in *Arabidopsis thaliana* RAD51-GFP line roots treated with 0 µM and 10 µM concentrations of zeocin.

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**Fig. S5** Quantification of *RAD51* mRNA and RAD51-GFP protein signals in *Arabidopsis thaliana* RAD51-GFP line roots treated with 50 µM concentration of zeocin.

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776

Yin K, Ueda M, Takagi H, Kajihara T, Sugamata Aki S, Nobusawa T, Umeda-Hara C,
Umeda M. 2014. A dual-color marker system for *in vivo* visualization of cell cycle progression
in Arabidopsis. *The Plant Journal* **80**: 541–552.

777

778

779

Yoshiyama KO, Kaminoyama K, Sakamoto T, Kimura S. 2017. Increased Phosphorylation
of Ser-Gln Sites on SUPPRESSOR OF GAMMA RESPONSE1 Strengthens the DNA Damage
Response in *Arabidopsis thaliana*. *The Plant Cell* **29**: 3255–3268.

780

781

782

Yu C, Hou L, Huang Y, Cui X, Xu S, Wang L, Yan S. 2023. The MULTI-BRCT domain protein DDRM2 promotes the recruitment of RAD51 to DNA damage sites to facilitate homologous recombination. *New Phytologist* **238**: 1073–1084.

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Zhao L, Fonseca A, Meschichi A, Sicard A, Rosa S. 2023. Whole-mount smFISH allows combining RNA and protein quantification at cellular and subcellular resolution. *Nature Plants* **9**: 1094–1102.

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SUPPORTING INFORMATION

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Additional supporting information may be found in the online version of this article.

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**Fig. S1** Evaluation of *PP2A* transcription in response to growing amounts of DNA damage in s-quashed roots of *Arabidopsis thaliana*.

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**Fig. S2** Quantification of RAD51 mRNA molecules in Col-0 *Arabidopsis thaliana* roots exposed to growing amounts of DNA damage using whole-mount smFISH.

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**Fig. S3** Evaluation of RAD51 transcriptional response in meristematic and elongation zones of the *Arabidopsis thaliana* root using whole mount smFISH.

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**Fig. S4** Quantification of *RAD51* mRNA and RAD51-GFP protein signals in *Arabidopsis thaliana* RAD51-GFP line roots treated with 0 µM and 10 µM concentrations of zeocin.

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**Fig. S5** Quantification of *RAD51* mRNA and RAD51-GFP protein signals in *Arabidopsis thaliana* RAD51-GFP line roots treated with 50 µM concentration of zeocin.

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**Fig. S6** Quantification of RAD51 mRNA and RAD51-GFP protein signals in *Arabidopsis thaliana* RAD51-GFP line roots treated with 170 µM concentration of zeocin.

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**Fig. S7** Evaluation of cell cycle arrest in *Arabidopsis thaliana* roots after exposure to growing amounts of DNA damage using EdU staining.

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**Fig. S8** Evaluation of cell cycle changes in *Arabidopsis thaliana* Cytrap line roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin.

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**Fig. S9** Evaluation of cell cycle changes in *Arabidopsis thaliana* PlaCCI line roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin.

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**Fig. S10** RAD51 mRNA molecule half-life evaluation under DNA damage in squashed roots of *Arabidopsis thaliana*.

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**Table S1** RAD51 transcription site (TS) representation in cells residing at G1 and other phases of the cell cycle in *Arabidopsis thaliana* CDT1-CFP line roots.

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**Fig. S6** Quantification of RAD51 mRNA and RAD51-GFP protein signals in *Arabidopsis thaliana* RAD51-GFP line roots treated with 170 µM concentration of zeocin.

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**Fig. S7** Evaluation of cell cycle arrest in *Arabidopsis thaliana* roots after exposure to growing amounts of DNA damage using EdU staining.

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**Fig. S8** Evaluation of cell cycle changes in *Arabidopsis thaliana* Cytrap line roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin.

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**Fig. S9** Evaluation of cell cycle changes in *Arabidopsis thaliana* PlaCCI line roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin.

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820

**Fig. S10** RAD51 mRNA molecule half-life evaluation under DNA damage in squashed roots of *Arabidopsis thaliana*.

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**Table S1** RAD51 transcription site (TS) representation in cells residing at G1 and other phases of the cell cycle in *Arabidopsis thaliana* CDT1-CFP line roots.

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828
a

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**Fig. 1 RAD51 transcriptional response to γ-H2AX accumulation in response to increasing DNA damage in *Arabidopsis thaliana* root squashes.** (a) qPCR quantification of RAD51 transcriptional response in roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. RAD51 expression measured relative to PP2A gene as a control. ANOVA revealed statistically significant difference in RAD51 expression by zeocin concentration (F(3)=78.05, p=9.59e-06)). Letters indicate results of TukeyHSD test with 95% confidence level. Error bars indicate standard deviation. (b) Schematic image of single mRNA molecule labelling using multiple fluorescent probes by smFISH protocol. (c) Images acquired using sequential smFISH and immunodetection protocol in squashed root cells after 50 µM zeocin exposure. (i) RAD51 mRNA detection via smFISH, nuclei counterstained with DAPI (blue). (ii) γ-H2AX immunodetection (red) and DAPI (blue) signals. Scale bars, 5 µm. (d) Total number of RAD51 mRNA molecules detected in 30 randomly selected cells after zeocin exposure from dataset containing (n=160, n=116, n=60, n=96 cells for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively), subset was selected 30 times. ANOVA revealed statistically significant difference in RAD51 mRNA molecule number by zeocin concentration (F(3)=1497, p<2e-16)). Letters indicate results of TukeyHSD test with 95% confidence level. (e) γ-H2AX fluorescence signal intensities measured after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin in squashed roots. Values represent nuclear signal fluorescence intensity measured as lg (Integrated Density). ANOVA revealed statistically significant difference in γ-H2AX fluorescence by zeocin concentration (F(3)=175.4, p<2e-08)) in our measurements (n=236, n=145, n=226, n=182 cells for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively). Letters indicate results of TukeyHSD test with 95% confidence level. (f) Correlation analysis between the number of RAD51 mRNA molecules and γ-H2AX signal intensity in individual cells of squashed roots with linear model fit. Number of RAD51 transcripts normalized by corresponding cell area, log10 of this value used for the corresponding axis. γ-H2AX fluorescence intensity measured as lg(Integrated Density) with prior normalization to DAPI Integrated density. Correlation coefficient (R) and p-value shown on a graph. DOF indicates deviance of fit calculated for the model. Dataset contains n=40, n=40, n=25, n=22 measurements for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively.
**Fig. 1 RAD51 transcriptional response to γ-H2AX accumulation in response to increasing DNA damage in *Arabidopsis thaliana* root squashes.** (a) qPCR quantification of RAD51 transcriptional response in roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. RAD51 expression measured relative to PP2A gene as a control. ANOVA revealed statistically significant difference in RAD51 expression by zeocin concentration (F(3)=78.05, p=9.59e-06)). Letters indicate results of TukeyHSD test with 95% confidence level. Error bars indicate standard deviation. (b) Schematic image of single mRNA molecule labelling using multiple fluorescent probes by smFISH protocol. (c) Images acquired using sequential smFISH and immunodetection protocol in squashed root cells after 50 µM zeocin exposure. (i) RAD51 mRNA detection via smFISH, nuclei counterstained with DAPI (blue). (ii) γ-H2AX immunodetection (red) and DAPI (blue) signals. Scale bars, 5 µm. (d) Total number of RAD51 mRNA molecules detected in 30 randomly selected cells after zeocin exposure from dataset containing (n=160, n=116, n=60, n=96 cells for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively), subset was selected 30 times. ANOVA revealed statistically significant difference in RAD51 mRNA molecule number by zeocin concentration (F(3)=1497, p<2e-16)). Letters indicate results of TukeyHSD test with 95% confidence level. (e) γ-H2AX fluorescence signal intensities measured after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin in squashed roots. Values represent nuclear signal fluorescence intensity measured as lg (Integrated Density). ANOVA revealed statistically significant difference in γ-H2AX fluorescence by zeocin concentration (F(3)=175.4, p<2e-08)) in our measurements (n=236, n=145, n=226, n=182 cells for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively). Letters indicate results of TukeyHSD test with 95% confidence level. (f) Correlation analysis between the number of RAD51 mRNA molecules and γ-H2AX signal intensity in individual cells of squashed roots with linear model fit. Number of RAD51 transcripts normalized by corresponding cell area, log10 of this value used for the corresponding axis. γ-H2AX fluorescence intensity measured as lg(Integrated Density) with prior normalization to DAPI Integrated density. Correlation coefficient (R) and p-value shown on a graph. DOF indicates deviance of fit calculated for the model. Dataset contains n=40, n=40, n=25, n=22 measurements for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively.
<figure><img src="image_10.png" /><figcaption>b</figcaption></figure>

<figure><img src="image_11.png" /><figcaption>c</figcaption></figure>

<figure><img src="image_12.png" /><figcaption>d</figcaption></figure>

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<figure><img src="image_9.png" /><figcaption>b</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>c</figcaption></figure>

<figure><img src="image_11.png" /><figcaption>d</figcaption></figure>

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Representative images of whole-mount smFISH for RAD51 mRNA in Col-0 roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. (i) Images of cell wall staining using Renaissance 2200 dye. (ii) Images of RAD51 mRNA detection. Scale bars, 20 µm. (b) Heatmaps representing quantification of RAD51 mRNA molecules detected in individual cells. (c) Frequency distribution of RAD51 mRNA molecules per cell, for plants treated with different zeocin concentrations. Bin groups created using a step of 10 transcripts. (d) Number of RAD51 mRNA molecules per cell in each of the selected cell types (Epidermis, Cortex, Endodermis, Stele) after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. Two-way ANOVA revealed statistically significant difference in RAD51 molecule number by both zeocin concentration (F(3)=119.93, *p*=2e-16)) and cell lineage (F(3)=25.39, *p*=1.71e-15)). Letters indicate results of TukeyHSD test of two-way ANOVA results with 95% confidence level. Measurements for (c) and (d) performed using data from images (a) and (b).
Representative images of whole-mount smFISH for RAD51 mRNA in Col-0 roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. (i) Images of cell wall staining using Renaissance 2200 dye. (ii) Images of RAD51 mRNA detection. Scale bars, 20 µm. (b) Heatmaps representing quantification of RAD51 mRNA molecules detected in individual cells. (c) Frequency distribution of RAD51 mRNA molecules per cell, for plants treated with different zeocin concentrations. Bin groups created using a step of 10 transcripts. (d) Number of RAD51 mRNA molecules per cell in each of the selected cell types (Epidermis, Cortex, Endodermis, Stele) after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. Two-way ANOVA revealed statistically significant difference in RAD51 molecule number by both zeocin concentration (F(3)=119.93, *p*=2e-16)) and cell lineage (F(3)=25.39, *p*=1.71e-15)). Letters indicate results of TukeyHSD test of two-way ANOVA results with 95% confidence level. Measurements for (c) and (d) performed using data from images (a) and (b).
<figure><img src="image_14.png" /><figcaption>b</figcaption></figure>

<figure><img src="image_15.png" /><figcaption>c</figcaption></figure>

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<figure><img src="image_13.png" /><figcaption>b</figcaption></figure>

<figure><img src="image_14.png" /><figcaption>c</figcaption></figure>

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**Fig. 3** Simultaneous detection and quantification of RAD51-GFP line roots. (a) Representative confocal images and quantification of RAD51 mRNA (ii) and RAD51-GFP protein signals (iii) after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. (i) Imaging of cell wall staining using Renaissance 2200 dye. (ii, iii) Imaging of (ii) RAD51 mRNA by smFISH and (iii) RAD51-GFP signals. (iv, v) Heatmaps representing the levels of the corresponding mean signal intensity per cell (MI). (vi) Signal intensity of RAD51 mRNA molecules. (v) Signal intensity of RAD51-GFP protein. (vi) Heatmaps representing the ratio between the RAD51 mRNA and RAD51-GFP protein signal intensities in each cell. Scale bars, 20 µm. (b) RAD51 mRNAs per cell after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. ANOVA revealed statistically significant difference in RAD51 mRNA signal mean intensity by zeocin concentration (F(3)=558.7, p<2e-16)). Letters indicate results of TukeyHSD test with 95% confidence level. (c) RAD51-GFP mean signal intensity per cell after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. ANOVA revealed statistically significant difference in RAD51-GFP signal mean intensity by zeocin concentration (F(3)=547.2, p<2e-16)). Letters indicate results of TukeyHSD test with 95% confidence level. Graphs on (b) and (c) created using dataset from several images containing (n=640, n=1391, n=854, n=457 cells for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively) individual cell measurements.
**Fig. 3** Simultaneous detection and quantification of RAD51-GFP line roots. (a) Representative confocal images and quantification of RAD51 mRNA (ii) and RAD51-GFP protein signals (iii) after exposure to 0 µM, 10 µM, 50 µM, 170 µM concentrations of zeocin. (i) Imaging of cell wall staining using Renaissance 2200 dye. (ii, iii) Imaging of (ii) RAD51 mRNA by smFISH and (iii) RAD51-GFP signals. (iv, v) Heatmaps representing the levels of the corresponding mean signal intensity per cell (MI). (vi) Signal intensity of RAD51 mRNA molecules. (v) Signal intensity of RAD51-GFP protein. (vi) Heatmaps representing the ratio between the RAD51 mRNA and RAD51-GFP protein signal intensities in each cell. Scale bars, 20 µm. (b) RAD51 mRNAs per cell after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. ANOVA revealed statistically significant difference in RAD51 mRNA signal mean intensity by zeocin concentration (F(3)=558.7, p<2e-16)). Letters indicate results of TukeyHSD test with 95% confidence level. (c) RAD51-GFP mean signal intensity per cell after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. ANOVA revealed statistically significant difference in RAD51-GFP signal mean intensity by zeocin concentration (F(3)=547.2, p<2e-16)). Letters indicate results of TukeyHSD test with 95% confidence level. Graphs on (b) and (c) created using dataset from several images containing (n=640, n=1391, n=854, n=457 cells for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively) individual cell measurements.
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**Fig. 4 Dynamics of RAD51 mRNA transcription throughout the cell cycle.** (a) Scheme of experimental setup used for quantification of S-phase cells. Seedlings were treated with different concentrations of zeocin concentrations for 10 hours, followed by additional treatment with zeocin and EdU for two hours. (b) Confocal images of roots acquired using sequential WM-smFISH/EdU staining protocol after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. Cell wall staining using Renaissance 2200 dye. (i) Detection of S-phase cells by EdU staining. (ii) RAD51 mRNA detection by smFISH. Scale bars, 20 µm. (c) RAD51 mRNA and EdU staining images of 10 µM zeocin sample with higher magnification showing RAD51 transcripts on EdU-negative cells. (i) Merged image showing RAD51 mRNA and EdU signals. White dashed box delineates magnified area. Scale bar, 20 µm. (ii) Magnified area showing RAD51 mRNA signals. Scale bar, 5 µm. (iii) Magnified area showing EdU signals. Scale bar, 5 µm. (d) Percentage of EdU positive cells in roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. ANOVA revealed statistically significant difference in EdU positive cell numbers by zeocin concentration (F(3)=24.38, *p*=1.67e-08)) in our measurements (n=1123, n=1356, n=1519, n=1208 measurements for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively). Error bars indicate standard deviation. Letters indicate results of TukeyHSD test with 95% confidence level. (e) Schematic representation of the cell cycle and its phases. Cell cycle phases labelled by corresponding fluorescent reporter indicated for each of the plant lines used for the cell cycle analysis. (f) Representation of cells in different phases of the cell cycle in Cytrap line roots, value shown in %. Representation of G1 cells is an approximation calculated by exclusion according to Cytrap line description. Dataset containing 1002, 816, 800, 998 individual measurements for 0 µM, 50 µM and 170 µM concentrations correspondingly was used. (g) Percentage of G1-phase cells in roots after for the different concentrations of zeocin in roots of PlaCCI plant line. ANOVA revealed statistically significant difference in EdU G1-phase cell numbers by zeocin concentration (F(3)=15.15, *p*=4.56e-07)) in our measurements (n=2526, n=2438, n=1997, n=1698 measurements for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively). Error bars indicate standard deviation. Letters indicate results of TukeyHSD test with 95% confidence level. (h) Representative confocal image of root cells from CDT1a-CFP plant line after exposure to 50 µM zeocin, showing with RAD51 mRNA signal detection via WM-smFISH. (i) Detection of CDT1a-CFP reporter. (ii) Detection RAD51 mRNA signals. Asterisks indicate transcription sites. (iii) Overlay of (i) and (ii) images. Scale bars, 20 µm.
**Fig. 4 Dynamics of RAD51 mRNA transcription throughout the cell cycle.** (a) Scheme of experimental setup used for quantification of S-phase cells. Seedlings were treated with different concentrations of zeocin concentrations for 10 hours, followed by additional treatment with zeocin and EdU for two hours. (b) Confocal images of roots acquired using sequential WM-smFISH/EdU staining protocol after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. Cell wall staining using Renaissance 2200 dye. (i) Detection of S-phase cells by EdU staining. (ii) RAD51 mRNA detection by smFISH. Scale bars, 20 µm. (c) RAD51 mRNA and EdU staining images of 10 µM zeocin sample with higher magnification showing RAD51 transcripts on EdU-negative cells. (i) Merged image showing RAD51 mRNA and EdU signals. White dashed box delineates magnified area. Scale bar, 20 µm. (ii) Magnified area showing RAD51 mRNA signals. Scale bar, 5 µm. (iii) Magnified area showing EdU signals. Scale bar, 5 µm. (d) Percentage of EdU positive cells in roots after exposure to 0 µM, 10 µM, 50 µM, 170 µM zeocin. ANOVA revealed statistically significant difference in EdU positive cell numbers by zeocin concentration (F(3)=24.38, *p*=1.67e-08)) in our measurements (n=1123, n=1356, n=1519, n=1208 measurements for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively). Error bars indicate standard deviation. Letters indicate results of TukeyHSD test with 95% confidence level. (e) Schematic representation of the cell cycle and its phases. Cell cycle phases labelled by corresponding fluorescent reporter indicated for each of the plant lines used for the cell cycle analysis. (f) Representation of cells in different phases of the cell cycle in Cytrap line roots, value shown in %. Representation of G1 cells is an approximation calculated by exclusion according to Cytrap line description. Dataset containing 1002, 816, 800, 998 individual measurements for 0 µM, 50 µM and 170 µM concentrations correspondingly was used. (g) Percentage of G1-phase cells in roots after for the different concentrations of zeocin in roots of PlaCCI plant line. ANOVA revealed statistically significant difference in EdU G1-phase cell numbers by zeocin concentration (F(3)=15.15, *p*=4.56e-07)) in our measurements (n=2526, n=2438, n=1997, n=1698 measurements for 0 µM, 10 µM, 50 µM, 170 µM zeocin respectively). Error bars indicate standard deviation. Letters indicate results of TukeyHSD test with 95% confidence level. (h) Representative confocal image of root cells from CDT1a-CFP plant line after exposure to 50 µM zeocin, showing with RAD51 mRNA signal detection via WM-smFISH. (i) Detection of CDT1a-CFP reporter. (ii) Detection RAD51 mRNA signals. Asterisks indicate transcription sites. (iii) Overlay of (i) and (ii) images. Scale bars, 20 µm.