Slow Replication Fork Velocity of Homologous Recombination-Defective Cells Results from Endogenous Oxidative Stress

International audienceReplications forks are routinely hindered by different endogenous stresses. Because homologous recombination plays a pivotal role in the reactivation of arrested replication forks, defects in homologous recombination reveal the initial endogenous stress(es). Homologous recombination-defective cells consistently exhibit a spontaneously reduced replication speed, leading to mitotic extra centrosomes. Here, we identify oxidative stress as a major endogenous source of replication speed deceleration in homologous recombination-defective cells. The treatment of homologous recombination-defective cells with the antioxidant N-acetyl-cysteine or the maintenance of the cells at low O2 levels (3%) rescues both the replication fork speed, as monitored by single-molecule analysis (molecular combing), and the associated mitotic extra centrosome frequency. Reciprocally, the exposure of wild-type cells to H2O2 reduces the replication fork speed and generates mitotic extra centrosomes. Supplying deoxynucleotide precursors to H2O2-exposed cells rescued the replication speed. Remarkably, treatment with N-acetyl-cysteine strongly expanded the nucleotide pool, accounting for the replication speed rescue. Remarkably, homologous recombination-defective cells exhibit a high level of endogenous reactive oxygen species. Consistently, homologous recombination-defective cells accumulate spontaneous γH2AX or XRCC1 foci that are abolished by treatment with N-acetyl-cysteine or maintenance at 3% O2. Finally, oxidative stress stimulated homologous recombination, which is suppressed by supplying deoxynucleotide precursors. Therefore, the cellular redox status strongly impacts genome duplication and transmission. Oxidative stress should generate replication stress through different mechanisms, including DNA damage and nucleotide pool imbalance. These data highlight the intricacy of endogenous replication and oxidative stresses, which are both evoked during tumorigenesis and senescence initiation, and emphasize the importance of homologous recombination as a barrier against spontaneous genetic instability triggered by the endogenous oxidative/replication stress axis

Signaling from Mus81-Eme2-Dependent DNA Damage Elicited by Chk1 Deficiency Modulates Replication Fork Speed and Origin Usage

International audienceMammalian cells deficient in ATR or Chk1 display moderate replication fork slowing and increased initiation density, but the underlying mechanisms have remained unclear. We show that exogenous deoxyribonucleosides suppress both replication phenotypes in Chk1-deficient, but not ATR-deficient, cells. Thus, in the absence of exogenous stress, depletion of either protein impacts the replication dynamics through different mechanisms. In addition, Chk1 deficiency, but not ATR deficiency, triggers nuclease-dependent DNA damage. Avoiding damage formation through invalidation of Mus81-Eme2 and Mre11, or preventing damage signaling by turning off the ATM pathway, suppresses the replication phenotypes of Chk1-deficient cells. Damage and resulting DDR activation are therefore the cause, not the consequence, of replication dynamics modulation in these cells. Together, we identify moderate reduction of precursors available for replication as an additional outcome of DDR activation. We propose that resulting fork slowing, and subsequent firing of backup origins, helps replication to proceed along damaged templates

Stepwise Activation of the ATR Signaling Pathway upon Increasing Replication Stress Impacts Fragile Site Integrity

<div><p>Breaks at common fragile sites (CFS) are a recognized source of genome instability in pre-neoplastic lesions, but how such checkpoint-proficient cells escape surveillance and continue cycling is unknown. Here we show, in lymphocytes and fibroblasts, that moderate replication stresses like those inducing breaks at CFSs trigger chromatin loading of sensors and mediators of the ATR pathway but fail to activate Chk1 or p53. Consistently, we found that cells depleted of ATR, but not of Chk1, accumulate single-stranded DNA upon Mre11-dependent resection of collapsed forks. Partial activation of the pathway under moderate stress thus takes steps against fork disassembly but tolerates S-phase progression and mitotic onset. We show that fork protection by ATR is crucial to CFS integrity, specifically in the cell type where a given site displays paucity in backup replication origins. Tolerance to mitotic entry with under-replicated CFSs therefore results in chromosome breaks, providing a pool of cells committed to further instability.</p></div

ATR-depletion enhances formation of ssDNA foci and fork asymmetry.

<p>(A) Percentage of S-phase cells with ssDNA foci in the indicated conditions of transfection and aphidicolin treatment. Mean ± s.e.m are presented. (B) Upper panel: Western blot analyses of total cell extracts 48 h post-transfection with a NONsi RNA or siRNAs specific to ATR or/and Mre11. Loading control: β-actin. Lower panel: percentage S-phase cells (mean ± s.e.m) displaying ssDNA foci in populations of cells depleted of ATR and treated or not with aphidicolin 0.3 µM. Mre11 status is indicated (siMre11 or treatment with Mirin: 100 µM). (C) Percentage of FISH signal co-localizing with ssDNA foci in cells depleted of ATR and treated with 0.3 µM aphidicolin. The immunostaining of ssDNA and PCNA was combined with FISH with BAC probes. Upper panel: JEFF lymphoblastoid cells, the BACs probe early replicating regions (875H7 and 456N14) and late replicating regions (321A23, 660J14, and 641C17 that corresponds to <i>FRA3B</i>, the major CFS in lymphocytes). Lower panel: MRC-5 fibroblasts, BAC 456N14 probe for an early replicating region, other BACs correspond to late replicating regions (875H7, 321A23, 641C17, and 660J14 which corresponds to the major CFS in fibroblast mapping at 3q13.3). (D) Distributions of fork asymmetry in cells transfected and treated as indicated. Forks travelling in the <i>FRA3B</i> locus are represented by red circles (<i>n</i> = 17) and in the bulk genome by black circles (untreated: NONsi, <i>n</i> = 128; siCHk1, <i>n</i> = 103; siATR, <i>n</i> = 147; with aphidicolin: NONsi, <i>n</i> = 121; siCHk1, <i>n</i> = 132; siATR, <i>n</i> = 123). Horizontal orange lines represent the medians of fork distributions. Medians and <i>P</i> values are indicated above the distributions (not significant, <i>ns</i>).</p

Moderate fork slowing triggers neither γ-H2AX foci nor ATM activation.

<p>(A) Western blot analysis of ATM-Ser1981 phosphorylation and γ-H2AX in exponentially growing cells, untreated (-) or treated as indicated. (B) Immunostaining of γ-H2AX together with chromatin-bound PCNA in untreated cells (-) and in cells treated with aphidicolin 0.6 µM for the indicated periods of time, or for 1 h with HU 1 mM.</p

Moderate fork slowing is not associated with formation of ssDNA foci.

<p>(A) ssDNA detection scheme. Cells hemi-substituted throughout their genome after growth in the presence of CldU for about 1.5 cell generations were treated or not with aphidicolin or HU for different periods of time prior to fixation. CldU was immuno-detected without DNA denaturation, which only permits visualization of substituted and single-stranded regions. (B) Co-detection of ssDNA (red) and chromatin bound PCNA (green) by immunostaining of untreated cells (-) and cells treated as indicated. Nuclei were counterstained with DAPI. (C) Typical pattern of PCNA and CldU foci in cells treated with HU 1 mM during 1 h. (D) Typical pattern of PCNA and CldU foci in cells depleted of ATR and treated with 0.3 µM of aphidicolin for 4 h.</p

Moderate fork slowing triggers chromatin loading of sensors and mediators of the ATR pathway.

<p>(A) Measurement of replication fork speed. Typical examples of combed DNA molecules displaying replications forks in control cells (Ctl) and cells treated with aphidicolin. Cells were pulse-labelled with IdU then CldU before DNA was purified and combed (see Methods). The analogues were revealed on stretched molecules as previously described <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1003643#pgen.1003643-Anglana1" target="_blank">[44]</a> and fork speed was determined by measuring the length of IdU (red) and CldU (green) tracks. Blue: counterstaining of the DNA used to select unbroken IdU and CldU tracks (B) Mean fork speed (upper panel), kb per min ± standard error of the mean (s.e.m.), and chromatin recruitment of checkpoint proteins (lower panel). Mean fork speed and chromatin recruitment of ATR and RPA2 were studied after 4 h of treatment with the indicated aphidicolin concentrations or 1 h of treatment with HU 1 mM. Asterisks indicate that fork speed cannot be measured. (C) Western blot analysis of chromatin extracts from exponentially growing cells, untreated (-) or treated as indicated.</p

ATR depletion compromises genome stability.

<p>(A) Quantification of metaphases (M) with breaks in cells transfected as above and treated for 16 h with the indicated aphidicolin concentrations. Mean ± s.e.m are presented. (B) Correlations between replication fork speed and chromosome instability. The data presented in A and <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1003643#pgen-1003643-g005" target="_blank">Figure 5C</a> were used to plot the percentage of metaphase plates with chromosome breaks against fork speed in each condition of transfection and treatment. (C) Breaks occurring at <i>FRA3B</i> quantified as in C. (D) Correlations between replication speed and <i>FRA3B</i> instability evaluated as in B.</p

Moderate fork slowing does not trigger phosphorylation of ATR targets.

<p>(A) Western blot detection of Chk1-Ser317 and Ser345 phosphorylations in total extracts of cells treated 4 h with aphidicolin or 1 h with HU 1 mM (upper panel). Quantification of Chk1-Ser317P and -Ser345P from 3 independent experiments (lower panel). <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1003643#s2" target="_blank">Results</a> (mean ± s.e.m) are expressed as percentage of the level of phosphorylation found in cells treated with 1 mM HU. Lanes 1–6 and lanes 7–9 correspond to different blots. (B) Western blot detection of RPA2-Ser33 phosphorylation, p53 and p53-Ser15 phosphorylation in total extracts of cells treated for 4 h with aphidicolin or 1 h with HU 1 mM. Loading control: β-actin.</p

Smarcal1-Mediated Fork Reversal Triggers Mre11-Dependent Degradation of Nascent DNA in the Absence of Brca2 and Stable Rad51 Nucleofilaments.

Brca2 deficiency causes Mre11-dependent degradation of nascent DNA at stalled forks, leading to cell lethality. To understand the molecular mechanisms underlying this process, we isolated Xenopus laevis Brca2. We demonstrated that Brca2 protein prevents single-stranded DNA gap accumulation at replication fork junctions and behind them by promoting Rad51 binding to replicating DNA. Without Brca2, forks with persistent gaps are converted by Smarcal1 into reversed forks, triggering extensive Mre11-dependent nascent DNA degradation. Stable Rad51 nucleofilaments, but not RPA or Rad51T131P mutant proteins, directly prevent Mre11-dependent DNA degradation. Mre11 inhibition instead promotes reversed fork accumulation in the absence of Brca2. Rad51 directly interacts with the Pol α N-terminal domain, promoting Pol α and δ binding to stalled replication forks. This interaction likely promotes replication fork restart and gap avoidance. These results indicate that Brca2 and Rad51 prevent formation of abnormal DNA replication intermediates, whose processing by Smarcal1 and Mre11 predisposes to genome instability.This work was funded by the Associazione Italiana per Ricerca sul Cancro (AIRC), European Research Council (ERC) Consolidator Grant 614541, the Armenise-Harvard Foundation career development award, Epigen Progetto Bandiera (4.7), AICR-Worldwide Cancer Research (13-0026), and Fondazione Telethon Grant GGP13-071 (to V.C.); by Wellcome Trust Investigator Award 104641/Z/14/Z (to L.P.); by the Czech Science Foundation (GACR 17-17720S and 13-26629S) and Project LQ1605 from the National Program of Sustainability II (MEYS CR) (to L.K.); and by NIH grant R01CA197774 and Susan G. Komen CCR grant CCR16377030 (to A.C.). V.S. is funded by a Fondazione Veronesi (FUV) personal postdoctoral fellowship