A local ATR-dependent checkpoint pathway is activated by a site-specific replication fork block in human cells

  1. Sana Ahmed-Seghir
  2. Manisha Jalan
  3. Helen E Grimsley
  4. Aman Sharma
  5. Shyam Twayana
  6. Settapong T Kosiyatrakul
  7. Christopher Thompson
  8. Carl L Schildkraut  Is a corresponding author
  9. Simon N Powell  Is a corresponding author
  1. Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, United States
  2. Department of Cell Biology, Albert Einstein College of Medicine, United States
  3. Molecular Biology Program, Memorial Sloan Kettering Cancer Center, United States

Abstract

When replication forks encounter DNA lesions that cause polymerase stalling, a checkpoint pathway is activated. The ATR-dependent intra-S checkpoint pathway mediates detection and processing of sites of replication fork stalling to maintain genomic integrity. Several factors involved in the global checkpoint pathway have been identified, but the response to a single replication fork barrier (RFB) is poorly understood. We utilized the Escherichia coli-based Tus-Ter system in human MCF7 cells and showed that the Tus protein binding to TerB sequences creates an efficient site-specific RFB. The single fork RFB was sufficient to activate a local, but not global, ATR-dependent checkpoint response that leads to phosphorylation and accumulation of DNA damage sensor protein γH2AX, confined locally to within a kilobase of the site of stalling. These data support a model of local management of fork stalling, which allows global replication at sites other than the RFB to continue to progress without delay.

eLife assessment

This manuscript reports important data on the cellular response to a single site-specific replication fork block in human MCF7 cells. Compelling evidence shows the efficacy of the bacterial Tus-Ter system to stall replication forks in human cells. Fork stalling led to lasting ATR-dependent phosphorylation of histone H2AX but not of ATR itself and its downstream targets RPA and CHK1.

https://doi.org/10.7554/eLife.87357.3.sa0

Introduction

Genomic DNA is constantly exposed to exogenous and endogenous damaging agents, forming DNA lesions that challenge the progression of replication forks (RFs) during the S-phase of the cell cycle. The genome contains natural impediments, which cause replication stalling such as common fragile sites, repeated sequences, non-B structures, sites of collision between transcription and replication machinery, or DNA-protein complexes (Mirkin and Mirkin, 2007). These replication fork barriers (RFB) can impede the progression of the replication machinery and, if unresolved, lead to genomic instability, a common hallmark of aneuploidy, neurological and neuromuscular disorders, and cancer.

Replication stress can impede the progression of the replication fork. During replication stress, the intra-S phase checkpoint pathway can be activated at a local and global level, which is a critical step to process the DNA lesions and maintaining genome integrity. The S-phase checkpoint involves the uncoupling of replicative polymerases from the replicative helicase and the generation of ssDNA, which rapidly gets coated with RPA. In turn, this activates ATR/ATRIP kinase signaling with its effector kinase CHK1, plus phosphorylation of H2AX at Ser139 in response to a variety of lesions that promote cell cycle arrest, fork stabilization, and restart (Iyer and Rhind, 2017; Iyer and Rhind, 2013; Willis and Rhind, 2009; Zeman and Cimprich, 2014).

Previous studies suggest that the local and global intra-S checkpoints are two distinct mechanisms that can be distinguished by the signaling intensity of detecting DNA damage (Saxena and Zou, 2022). It is thought that a threshold of DNA damage must be reached to activate the global checkpoint and impact cell cycle progression during replication (Bantele et al., 2019; Shimada et al., 2002). The local checkpoint reacts to a unique replication fork stalling site, which does not impact overall cell cycle progression. The proteins involved in the global replication stress response have been identified using DNA damaging agents that lead to multiple DNA lesion formation randomly throughout the genome. Nevertheless, the mechanism and timing of the checkpoint pathway at a local scale remain largely unknown and are critical to understanding the series of processes at a single RF stall.

In the Escherichia coli genome, the DNA pausing sequences called terminator (Ter) are recognized by a protein called Tus to cause a polar site-specific arrest of the RF at the end of the bacterial chromosome replication (Hiasa and Marians, 1994; Hidaka et al., 1988; Mulcair et al., 2006; Roecklein et al., 1991). The Tus-Ter complex leads to a temporarily locked complex on DNA that can be overcome by the fork arriving in the opposite direction, displacing Tus to terminate replication. The artificial E. coli-based Tus/Ter system has previously been employed in mouse embryonic stem cells to measure homologous recombination repair products and investigate the DNA repair pathway choice (Chandramouly et al., 2013; Willis et al., 2017; Willis et al., 2014).

In this study, we integrated a plasmid with five repeats of the TerB sequence in the non-permissive orientation, referred to as pWB15, at a unique site within chromosome 12 of the breast cancer cell line MCF7 (MCF7 5C-TerB clone). We utilized the integrated Tus-TerB system in human MCF7 cells to artificially generate individual RFBs and investigated the activation of the S-phase checkpoint signaling mechanism. We show that Tus/Ter creates an efficient site-specific RFB in human cells and observed local activation of ATR signaling, which was responsible for the phosphorylation of DNA damage marker γH2AX at the stall sites. When a replication fork pauses at the local Tus-TerB block, we do not detect any alteration in global replication profiles. Our system allows us to study the ATR-checkpoint activity as a local response to a single RFB.

Results

Tus is found enriched at TerB sites integrated in human cells

The 23 bp TerB sequence in interaction with the Tus protein have been successfully used in yeast and mice to create artificial RFBs (Larsen et al., 2014a; Willis et al., 2018; Willis et al., 2017; Willis et al., 2014; Willis and Scully, 2016). To study the effect of site-specific replication blocks at a genomic locus in human cells, we used a previously established MCF7 clone with a unique integrated copy of the pWB15 plasmid carrying two sets of five TerB repeat sequences in opposite, non-permissive orientations with respect to the incoming replication forks (Figure 1A and Figure 1—figure supplement 1). Whole-genome sequencing of the clone was carried out to confirm the single-copy integration of the plasmid at hg38 chr12:95325001. Firstly, we checked whether Tus protein was able to bind TerB sequences in this artificial system in human cells. Chromatin immunoprecipitation (ChIP) followed by quantitative polymerase chain reaction (qPCR) was performed using an antibody against the His-tag 24 hr after MCF7 5C-TerB cells were transfected with a Tus-His expression plasmid. The Tus signal is specifically enriched over 20 times compared to the vector control (VC) condition in proximity to the TerB sequences (PP9, 2, and 47) (Figure 1B).

Figure 1 with 1 supplement see all
Tus is bound to TerB sites integrated in human cells.

(A) Schematic of linearized plasmid (pWB15) integrated into MCF7 cells (MCF7 5C-TerB) with two cassettes containing five tandem TerB sequences in the non-permissive orientation (black arrows). Black half-arrowheads depict polymerase chain reaction (PCR) products expected from stated primer pairs used in (B). (B) Chromatin immunoprecipitation (ChIP) with His antibody on MCF7 5C-TerB cells transfected with either a vector control (VC) or HA-Tus-His expression plasmids. ChIP-qPCR was conducted using the indicated primer pairs. Data show the percentage of input (n = 3). (C) Schematic of proximity ligation assay (PLA) to visualize Tus bound to TerB sites using HA and His antibodies. (D) Representative images of the PLA foci across stated conditions. Scale Bars, 10  μm (E) Percentage of cells with 1–2 PLA foci per nucleus. (n = 3, ≥150 cells per experiment). Bar graphs represents mean ± standard error of the mean (SEM), Student t-test (see methods).

To corroborate this observation, we performed the proximity ligation assay (PLA) (see ‘Methods’). Two tagged-Tus (HA-Tus and Tus-His) or a VC expression plasmid were simultaneously transfected and antibodies against the two tags, HA and His, were used to generate a PLA signal (Figure 1C and D). We observed 19% of the cells transfected with the two tagged Tus were harboring one or two PLA signal versus 9% of the cells transfected with the VC (Figure 1E), a significant enrichment of the PLA signal when Tus was bound to TerB. These results indicate a specific interaction of the Tus protein with the TerB sequences in human cells.

The Tus-TerB barrier is an efficient block to incoming replication forks

It has previously been demonstrated that the Tus-TerB interaction can induce an RFB in mouse embryonic cells using a chromosomally integrated system (Willis et al., 2014). We reasoned that our integrated system in MCF7 cells would be more prone to replication fork arrests near the TerB sequences. We used single-molecule analysis of replicated DNA (SMARD) to investigate evidence of replication fork stalling in the genomic DNA (Norio and Schildkraut, 2001). The self-labeling SNAP-tag was used to label Tus. After induction of Myc-NLS-TUS-SNAP by using Dox for 3 d, asynchronous MCF7 5C-TerB cells were sequentially pulse-labeled using two nucleoside analogs (iododeoxyuridine [IdU] and chlorodeoxyuridine [CldU]) for 4 hr each (Figure 2A). The isolated DNA was digested by restriction endonuclease Sfil, and the 200 kb fragment obtained was stretched on slides for staining. Two DNA FISH probes were used to detect the 200 kb segment of interest containing the TerB arrays: one probe of 40 kb within the chromosome 12 (hg38 chr12:95,199,827–95,239,226) and one probe of 7 kb made from pWB15 (Figure 2B). The direction of replication forks could be determined using the labeling patterns and its position within the locus of interest using our FISH probes. After inducing the expression of Myc-NLS-TUS-SNAP from an integrated plasmid in MCF7 5C-TerB cells (Figure 2—figure supplement 1A and B), we analyzed its impact on the replication fork progression within the region containing TerB repeats (Figure 2C and D). The yellow arrows indicate the transition of labeling from IdU to CldU incorporation showing the replication fork direction. When Tus was expressed, these transition sites were found accumulated near the TerB regions (Figure 2D, arrows within the white oval) compared to the random signal spread on the 200 kb region when Tus is not expressed (Figure 2C). We quantified the percentage of molecules with replication forks at each 5 kb interval in the 200 kb SfiI segment containing TerB sequences (Figure 2E and F) and confirmed an accumulation of forks at the TerB sequences only when Tus was expressed (Figure 2F, black arrow), supporting that the binding of Tus on TerB arrays can impair the replisome progression at genomic loci. Interestingly, expression of the Tus protein alone does not alter the direction of replication fork progression along the 200 kb SfiI segment where forks predominantly progress from the 3′ to 5′ direction in cells both with and without Tus expression as analyzed by the percentage of molecules with IdU incorporation at each 5 kb interval (Figure 2—figure supplement 1C and D). Furthermore, no significant difference was observed in the global replication fork speed in cells with or without Tus expression (Supplementary file 1a). Together, these data indicate that when Tus bound the TerB sequences integrated within chromosome 12, it creates a physical barrier and an obstacle to the replication fork progression.

Figure 2 with 2 supplements see all
Replication fork pauses at the Ter sequence in the presence of the Tus protein.

(A) Schematic of pulse labeling of MCF7 5C-TerB cells with Tus protein expression for 3 d. (B) Locus map of a 200 kb SfiI segment from MCF7 5C-TerB Chromosome 12 with the integrated plasmid (pWB15) DNA (orange). A 40 kb FISH probe made from fosmid WI2-1478M20 and a 7.5 kb FISH probe made from plasmid pWB15 are shown in blue. (C, D) Top: locus map of the DNA segment containing Ter sequence with the location of the FISH probes. Bottom: photomicrographs of labeled DNA molecules from MCF7 5C-TerB, Tus not expressed (C) and MCF7 5C-TerB, Tus expressed (D). Yellow arrows indicate the position of replication forks at the transition of labeling from IdU to CldU incorporation showing replication fork direction. Molecules are arranged in the following order: forks progressing from 3′ to 5′, forks progressing from 5′ to 3′, termination events, and initiation events. Replication forks (yellow arrows) in the white oval are all at the same location (at the TerB sequence) on molecules from different cells indicating that replication forks are pausing at this TerB sequence. (E, F) Percentage of molecules with replication forks at each 5 kb interval in the 200 Kb SfiI segment containing TerB sequence, quantified from molecules in MCF7 5C-TerB (E) Tus not expressed and (F) Tus expressed. Percentage of molecules with replication forks progressing 3′ to 5′ (< blue) and 5′ to 3′ (> orange) are shown. In the cells expressing Tus, a high percentage of molecules contain replication forks pausing in both directions in the 5 kb interval that contains Ter sequences (black arrow [F], white oval [D]). (G) Schematic similar to Figure 1A with progression of the endogenous origins of replication within the chromosome 12 shown (green arrows). (H) Chromatin immunoprecipitation (ChIP) using MCM3 antibody on MCF7 5C-TerB cells transfected with vector control (VC) or HA-Tus-His plasmids. ChIP-qPCR was conducted using the indicated primer pairs. Data shows the fold enrichment relative to the IgG controls (n = 3). Bar graphs represents mean ± standard error of the mean (SEM), Student t-test (see methods).

The eukaryotic replisome is a multiprotein apparatus consisting of DNA helicases, DNA polymerases and accessory proteins to duplicate the genome. We hypothesized that an RFB would induce an accumulation of these proteins on chromatin around TerB sequences. To test our hypothesis, we performed ChIP using an antibody against mini-chromosome maintenance protein 3 (MCM3), a CMG core-complex protein, in the absence or presence of Tus. When Tus was expressed, MCM3 was threefold more enriched upstream of the TerB arrays at the PP0-2 and PP10 sites compared to the surrounding primer pairs (Figure 2G and H). Additionally, we find that there is an enrichment of the DNA repair scaffold protein FANCM at the TerB sites mirroring the MCM3 enrichment (Figure 2—figure supplement 2), indicative of the stalled replication forks as previously described (Collis et al., 2008; Nandi and Whitby, 2012; Panday et al., 2021; Xue et al., 2015). Together, these results show that Tus expression induces an RFB at the TerB arrays impairing the replisome progression during S-phase.

γH2AX is enriched at TerB sites after Tus expression before fork collapse

One of the earliest responses to replication stress is the phosphorylation of the histone variant H2AX on serine 139 (γH2AX) by members of the phosphoinositide 3-kinase (PI3K)-like family (PIKK) (Ward and Chen, 2001). As we demonstrated that TerB arrays can block incoming replication forks, we asked whether we could see a γH2AX signal enrichment due to fork arrest. Using a co-immunoprecipitation assay with the chromatin fraction of cross-linked cells expressing either GFP tag or GFP-Tus, we found that GFP-Tus and γH2AX were immunoprecipitated together using GFP or γH2AX antibodies (Figure 3A).

Figure 3 with 2 supplements see all
γH2AX is enriched at TerB sites after Tus expression.

(A) MCF7 5C-TerB cells transfected with GFP or GFP-Tus, γH2AX and IgG antibodies were used to immuno-precipitate proteins and analyzed by immunoblotting with indicated antibodies. (B) Schematic of proximity ligation assay (PLA) to visualize HA-Tus bound to TerB in the proximity of γH2AX sites using HA and γH2AX antibodies. (C) Representative images of the PLA foci across stated conditions. Scale Bars, 10  μm. (D) Percentage of cells with 1–2 PLA foci per nucleus (n = 3, ≥150 cells per experiment). (E) Schematic of MCF7 5C-TerB depicting the positions of the primer pairs with respect to the integrated TerB locus. (F) γH2AX levels along the integrated TerB plasmid were analyzed by ChIP-qPCR in MCF7 5C-TerB cells transfected with vector control (VC) or Tus expression plasmids using the indicated primer pairs. Data shows the fold enrichment relative to IgG controls (n = 3). Bar graphs represents mean ± standard error of the mean (SEM), Student t-test (see methods).

To gain insight into the γH2AX signal at the RFB induced by the Tus-TerB interaction, cells were transfected with either VC or HA-Tus expression plasmids, and PLA was performed using antibodies against HA and γH2AX (Figure 3B). We found that 16% of the HA-Tus transfected cells were harboring at least one PLA signal versus 3% of the VC transfected cells (Figure 3C and D).

To better characterize the γH2AX enrichment around TerB sites, we performed ChIP-qPCR assays using a γH2AX antibody 24 hr after Tus or VC expression. We observed an enrichment of γH2AX, strictly co-localizing with the Tus-Ter interaction sites (PP9 and PP47), suggesting a local role of γH2AX in response to stalled RFs (Figure 3E and F). This is in stark contrast to the observed spreading of the γH2AX signal after a site-directed DSB (Berkovich et al., 2007; Chailleux et al., 2014; Clouaire et al., 2018; Savic et al., 2009). To corroborate this observation in our integrated cassette, we generated a site-specific DSB by expressing Cas9 targeted to the integrated cassette (Figure 3—figure supplement 1A–C). We assessed γH2AX using the ChIP-qPCR assay 24 hr after transfection and noticed enrichment at the distal primer pairs (PP0-2 and PP10), contrasting with the very tight local signal observed at the Tus-TerB RFB (Figure 3—figure supplement 1D). However, we did not find any significant γH2AX signal between VC or Tus expression (Figure 3—figure supplement 2A and B), supporting the lack of global response. This suggests that the RFB generated by the interaction between Tus and the TerB arrays activates a stress response that stimulates the phosphorylation of H2AX, concentrated around the fork block.

H2AX is phosphorylated in an ATR-dependent manner without a global activation of the intra-S checkpoint

ATR is known to phosphorylate H2AX in response to replication stress and plays a pivotal role in the surveillance of DNA replication (Ward and Chen, 2001). To understand whether the global ATR-dependent checkpoint was activated in response to the Tus-TerB RFB, we analyzed the levels of phospho-proteins involved in the intra-S checkpoint activation pathway (RPA, CHK1, and ATR) by immunoblotting fractionated cell extracts 24 hr after VC or Tus expression. Cells treated with 2 mM hydroxyurea (HU) for 4 hr demonstrated an increase in phospho-protein levels, acting as a control for checkpoint activation (Figure 4A, lanes 3 and 4). Tus expression alone did not activate the global ATR-dependent checkpoint as the level of phospho-protein remained the same as the VC control (Figure 4A, lanes 1 and 2). We did not find any observable difference in cell cycle progression between VC or Tus expression alone (Figure 4—figure supplement 1A and B), supporting the lack of global response.

Figure 4 with 1 supplement see all
γH2AX phosphorylation at Tus-TerB is ATR-dependent.

(A) MCF7 5C-TerB cells transfected with vector control (VC) or HA-Tus were treated for 4 hr with 2 mM hydroxyurea (HU) before lysis and fractionation (whole-cell extract [WCE], soluble fraction, and chromatin fraction). Total and phosphorylated protein levels were examined by immunoblotting as indicated. Tus expression was confirmed in WCE. (B) Depletion of pATR Th1989 in MCF7 5C-TerB cells treated with 2 mM HU ± 40 nM of ATR inhibitor was examined with immunoblotting. (C) Schematic of MCF7 5C-TerB depicting the positions of the primer pairs with respect to the integrated TerB locus. (D, E) γH2AX levels were analyzed by ChIP-qPCR in MCF7 5C-TerB cells transfected with (D) VC expression plasmid (E) Tus expression plasmid, ±4 hr treatment with ATRi with indicated primer pairs. Data shows the fold enrichment over the IgG controls (n = 3). Bar graphs represents mean ± standard error of the mean (SEM), Student t-test (see methods).

We hypothesized that if the local γH2AX enrichment at TerB sites was ATR-dependent, a reduction in ATR activity would lead to a decreased γH2AX signal. To validate the ATR inhibitor activity (ATRi), VE-822 (Fokas et al., 2012), we performed immunoblotting using total protein extracts of cells treated with 2 mM HU and compared the level of phospho-ATR TH1989 with or without an ATR inhibitor. A 4 hr treatment with ATRi reduced the level of phospho-ATR TH1989, reflecting a decrease in ATR activity (Figure 4B). ChIP-qPCR assays were performed using a γH2AX antibody 24 hr after VC or Tus expression, and cells were harvested after a 4 hr ATRi treatment. Consistent with our hypothesis, the γH2AX enrichment in the vicinity of TerB sites after Tus expression was remarkably decreased upon ATRi treatment (Figure 4C–E, PP9 and PP47), implying that the Tus-TerB RFB activates a localized stress response that stimulated the local phosphorylation of H2AX. Together, these data suggest a local activation of the intra-S checkpoint via the ATR kinase and rapid phosphorylation of H2AX, which could mediate the recruitment of repair factors near the damage site.

Discussion

Replication stress occurs during the S-phase of the cell cycle, when the replication machinery encounters DNA lesions that cause stalling of replicative polymerases and can be a significant cause of genomic instability. If the stalled forks cannot be processed, they can result in DNA breakage, mutations, and chromosomal rearrangements, leading to the development of many different human cancers (Gaillard et al., 2015; Tubbs and Nussenzweig, 2017). In this study, we used the reconstituted E. coli-derived protein-DNA barrier, Tus-TerB, to study the replication stress response at a single RFB. We have shown that the Tus protein efficiently binds at the TerB sequences using ChIP-qPCR and PLA in MCF7 cells (Figure 1). We confirmed that this integrated system was able to cause site-specific RFB at TerB sequences when Tus was introduced with the use of the SMARD technique and observed the accumulation of replication fork protein MCM3 upstream of TerB sequences (Figure 2). Our results indicate that the integrated Tus-TerB system acts as an efficient site-specific RFB, providing valuable insights into the local processing of a stalled single fork in human cells and its difference from global replication stress (Figure 5).

A local ATR-dependent checkpoint is activated by the Tus-TerB replication fork barrier (RFB).

A model depicting the cellular response to a site-specific DSB using CRISPR-Cas9 and a global replication stress with hydroxyurea, both of which lead to a DNA damage response (DDR, yellow triangles), increased gH2AX (green) levels globally in the cell and, if left unresolved, can alter cell cycle progression. In contrast, the site-specific replication fork block, Tus/TerB, elicits a local ATR-dependent DDR, which is responsible for the phosphorylation and accumulation of γH2AX at the stalled site. This local signaling does not affect the progression of the cell cycle and is not altered during the local checkpoint response.

We show that the site-specific replication stress leads to the accumulation of γH2AX at the site of fork block when Tus was expressed by co-immunoprecipitation of Tus and γH2AX (Figure 3A). Additionally, we confirmed that γH2AX is localized at the site of Tus-TerB using PLA and ChIP-qPCR assays (Figure 3D and F). We suggest that the γH2AX signal is being constrained to a region of less than a kilobase from the TerB sites by tortional stress generated when the replication fork encounters the barrier and is not influenced by well-defined topological domains that are one measure of the functional units of the genome (Dixon et al., 2012; Rao et al., 2014). Furthermore, there was no observable difference in global replication profiles or the activation of global checkpoint markers such as pRPA, pCHK1, and pATR, with or without Tus protein expression, contrasting with the differences observed in the presence and absence of HU-induced replication stress (Figure 4A, Supplementary file 1a, Figure 3—figure supplement 2, and Figure 4—figure supplement 1). We conclude that Tus-TerB-induced replication fork stall does not activate global ATR-dependent S-phase checkpoint signaling, suggesting that a local response is occurring at the stalled replication fork. Interestingly, upon ATR inhibition, we observed that the enrichment of phosphorylated H2AX at TerB sites was significantly reduced when Tus was expressed, which indicates local ATR signaling is responsible for the phosphorylation of H2AX at the stalled site (Figure 4C and E). A similar localized response had previously been observed in yeast, with a site-specific DSB in G1 cells, where there was no activation of the global checkpoint signaling, but γH2AX signaling was observed (Janke et al., 2010). This localized ATR-dependent γH2AX differs from the well-observed ATM-dependent spreading of γH2AX signal after a site-directed double-strand break (Berkovich et al., 2007; Chailleux et al., 2014; Clouaire et al., 2018; Savic et al., 2009) and suggests a distinct local intra-S phase checkpoint is being activated in response to the single RFB.

There are additional mechanisms to be understood in this newly described local replication stress response, which includes whether the activation of ATR is dependent on ATRIP bound to ssDNA (Ball et al., 2005; Zou and Elledge, 2003). In the Tus-TerB system, there is no known dissociation of the replicative helicase from the polymerase, which is responsible for the ssDNA signal. Therefore, just like ATM can be activated independently of the MRE11 complex (Bakkenist and Kastan, 2003), we expect that ATR is activated independently of ATRIP, likely due to local changes in chromatin that will be investigated in future studies.

During extensive DNA damage, the genome-wide checkpoint response activates ATR/CHK1 globally, which leads to the slowing of all replication forks in the cell, inhibition of cell cycle progression, and suppression of late origin firing; to provide sufficient time for DNA repair. It has been proposed that a local checkpoint response occurs when one or a few replication forks encounter a DNA lesion and activates ATR/CHK1 signaling at local sites of fork stalling. The local ATR response has been hypothesized to be transient, in which the fork moves slowly only at stalled sites to promote fork stabilization, restart the stalled fork, and suppress recombination without triggering the global checkpoint response (Iyer and Rhind, 2017; Iyer and Rhind, 2013; Kaufmann et al., 1980; Merrick et al., 2004; Saxena and Zou, 2022; Willis and Rhind, 2009; Zeman and Cimprich, 2014).

We predict that the local ATR-dependent checkpoint signaling can result in a rapid and controlled response to allow RFB resolution through the recruitment of repair factors to the damaged site. The factors might include those involved in fork stabilization, fork reversal, fork cleavage, and homology-dependent replication restart (Saxena and Zou, 2022). The Tus-TerB system has previously been employed in mouse embryonic stem cells to measure and investigate the DNA repair pathway choice (Chandramouly et al., 2013; Panday et al., 2021; Willis et al., 2017; Willis et al., 2014). This included both error-free and error-prone homologous recombination induced by a mammalian chromosomal RFB, as well as identifying the role of the structure-specific endonuclease complex SLX4-XPF and FANCM in the repair of microhomology-mediated tandem duplications that occur at replication arrest (Elango et al., 2022; Nandi and Whitby, 2012; Willis et al., 2017). The Tus-TerB block is robust and is not dissociated by the action of DNA helicases such as PIF1, corroborated in previous yeast studies that have demonstrated that the ‘sweepase’ RRM3 is unable to resolve this RFB (Larsen et al., 2014b). Therefore, it is likely that the RFB is resolved either by incoming replication forks leading to termination at the TerB site or by fork cleavage (Larsen et al., 2014b; Willis et al., 2017; Willis et al., 2014). Fork reversal at the Tus-TerB site has been observed in the yeast system (Larsen et al., 2014b; Marie and Symington, 2022), but not for the mammalian systems, and it is unlikely that shifting the position of the RFB would make it easier to resolve. Therefore, fork cleavage is the most likely mechanism for resolving the RFB.

The upstream pathway of activation of the DNA damage response, as a result of a single RFB, is not completely understood. Previous work has shown that FANCM acts as a scaffolding protein for recruitment of different repair protein complexes involved in the stalled replication fork rescue (Panday et al., 2021; Willis et al., 2017). We observed that the FANCM protein is enriched at the site of the RFB induced by Tus-TerB (Figure 2—figure supplement 2), confirming a similar phenomenon observed in mouse cells. However, it is yet to be determined whether FANCM recruitment to the RFB precedes local H2AX phosphorylation by ATR or whether FANCM is recruited as a consequence of the γH2AX signal to help elicit the downstream DNA damage response (DDR). We also anticipate that the protein complex, 9-1-1 (RAD9A, HUS1, RAD1), and the subsequently recruited TOPBP1, may also play a role at this local ATR-dependent intra-S checkpoint (Helt et al., 2005; Parrilla-Castellar et al., 2004).

While the Tus-TerB system is an important tool in deciphering signaling at individual replication forks, it will be important to test whether the local-S phase model remains supported when exploring endogenous lesions that occur when replication forks encounter secondary structures such as R-loops, G-quadruplexes, and common fragile sites characterized by trinucleotide repeat expansion [(CAG)n/(CTG)n] (Brickner et al., 2022; Bryan, 2019; Kim et al., 2017). Here the mechanisms surrounding the identification and resolution of the RFB may differ. Overall, our results indicate that Tus-TerB system acts as an efficient RFB and activates local ATR checkpoint signaling at the stall site, leading to phosphorylation and accumulation of the DNA damage sensor protein γH2AX, which is dependent on the ATR kinase. The local γH2AX accumulation at the stalled region would lead to the recruitment of DNA repair factors for resolution of the fork (Figure 5). Together, our findings reveal the signaling mechanism of the Tus/TerB-induced replication block and showed that the site-specific fork block is dependent on the local ATR S-phase checkpoint signaling.

Methods

Cell culture and transfection conditions

MCF7 5C-TerB cells were grown at 37°C and 5% CO2 in complete DMEM high glucose supplemented with 10% FBS, 2 mM GlutaMAX, 20 mM HEPES, 100 I.U./ml penicillin, and 100 μg/ml streptomycin supplemented with 400 µg/ml zeocin. The cell line was characterized using the Cell Line Authentication by STR analysis at the Integrated Genomics Operation, MSKCC. Cells were routinely tested for mycoplasma contamination using the Universal Mycoplasma Detection Kit (ATCC 30-1012K).

For PLA, cells were seeded at 1 × 105 in a 12-well cell culture plate. For T7 assay, cells were seeded at 2.5 × 105 in a 6-well cell culture plate. For ChIP, IP, and cellular fractionation, cells were seeded at 2.5 × 105 in a 10 cm cell culture dish.

MCF7 5C-TerB cells were seeded and transfected the day after with 10 μg of pCMV3xnls Tus or pCMV3xnls using the Mirus Bio TransIT-LT1 reagents for 24 hr more. For the generation of DSB, cells were transfected 6 hr prior to harvesting with 120 pmol of the sgRNA TerB1 and 120 pmol of the Cas9 protein (obtained from QB3 Macrolab, Berkeley) using the Lipofectamine CRISPRMAX Cas9 transfection reagents.

Generation of stable cell lines

PvuI linearized pWB15 was transfected into MCF7 (ATCC HTB-22) cells to generate the stably integrated clone, MCF7 5C-TerB (Figure 1—figure supplement 1). Then, 800 μg/ml of zeocin was used to select positively integrated cells and monoclonal colonies were isolated and subsequently maintained in 400 μg/ml zeocin. Single-integrant validation was performed by whole-genome sequencing of the clone.

Stable MCF7 5C-TerB cells-inducible-expressing Myc-NLS-TUS-SNAP were generated by lentiviral transduction. Cells were infected with lentiviral particles containing pInducer10L or pInd Tus-SNAP, and single-cell clonal colonies were selected in complete DMEM containing 1 µg/ml puromycin and 400 µg/ml zeocin. Myc-NLS-TUS-SNAP protein was expressed in stable lines by induction with 1 µg/ml doxycycline for 3 d.

Plasmid construction

The doxycycline-inducible lentiviral plasmid used to express N-terminally myc-tagged, C-terminally SNAP-tagged nuclear localized, human codon-optimized wild-type Tus (Myc-NLS-TUS-SNAP) was generated as follows. The N-terminally myc epitope-tagged, nuclear-localized, codon-optimized wild-type Tus cDNA sequence was PCR-amplified from pcDNA3-β-MYC-NLS-Tus using primers forward 5′-AGTCGGTACCGAATTCGCCACCATGGAACAAAAGCTG-3′ and reverse 5′-AGTCGGCGGCCGCGCCGCTACCGTCAGCCACGTACAGGTGCA and the SNAP-tag cDNA sequence PCR amplified using primers forward 5′-AGTCGCGGCCGCCGGCCACATGGACAAAGACTGCGAAATGAAGC-3′ and reverse 5′-ACTGCTCGAGTCAACCCAGCCCAGGCTTGC-3′. The Tus and SNAP PCR-amplified fragments were digested with KpnI and NotI, and NotI and XhoI, respectively, and inserted into the inducible expression lentiviral vector pInducer10L (Drosopoulos et al., 2020) directly downstream of the doxycycline-inducible promoter, to generate pInd Tus-SNAP. This construct was sequenced to confirm that unintended mutations were not introduced during PCR and cloning.

Immunoblotting

Cells were harvested by trypsinization, suspended in complete DMEM, washed with PBS, then pelleted and flash frozen in liquid N2 and stored at –80°C. For SDS-PAGE, pellets were thawed on ice and lysed by resuspending in Laemmli buffer (60 mM Tris-HCl pH 6.8, 400 mM 2-mercaptoethanol, 2% SDS, 10% glycerol, 0.01% bromophenol blue) to a final concentration of 106 cells/ml. Lysates were denatured (5 min at 100°C) and passed through a 25-gauge needle (5×) then spun for 2 min at full speed in a microfuge. Aliquots of lysate corresponding to 105 cells were resolved on 4–15% gradient SDS-PAGE gels, proteins transferred to nitrocellulose membrane and blocked in PBS with 5% Blotting-grade Blocker and 0.1% Tween 20. Membranes were then incubated with primary antibodies diluted in PBS with 5% Blotting-grade Blocker. Primary antibodies used were anti-Myc tag and anti-actin. Following incubation with primary antibodies, membranes were washed with PBS + 0.1% Tween 20. Membranes were then incubated with fluorescently labeled goat anti-mouse IRDye 680LT and goat anti-rabbit IRDye800CW secondary antibodies and then washed in PBS + 0.1% Tween 20. Immunoblots were imaged on an Odyssey Lc Infrared scanner.

SMARD assay

SMARD was performed essentially as previously described (Drosopoulos et al., 2012; Norio and Schildkraut, 2001; Twayana et al., 2021). Exponentially growing cells in complete DMEM with 1 µg/ml doxycycline were sequentially pulse-labeled with IdU (4 hr), followed by CldU (4 hr) each to a final concentration of 30 μΜ. Following pulsing, cells were suspended in PBS at a concentration of 3 × 107 cells per ml. An equal volume of 1% molten TopVision low-melting point agarose in PBS was added. The resulting cell suspension in 0.5% TopVision agarose was poured into wells of a chilled plastic mold to make plugs of size 0.5 cm × 0.2 cm × 0.9 cm, each containing 106 cells. Cells in the plug were lysed at 50°C in a buffer containing 1% n-lauroylsarcosine, 0.5 M EDTA, and 0.2 mg/ml proteinase K. Plugs were rinsed in TE and washed with 200 μM phenylmethanesulfonyl fluoride (PMSF). Genomic DNA in the plugs was digested with SfiI overnight at 37°C. Digested genomic DNA was cast into 0.7% SeaPlaque GTG agarose gel, and DNA was separated by pulsed field gel electrophoresis (PFGE) using a CHEF-DRII system (Bio-Rad). The 200 kb DNA segment containing TerB sequences within the gel was located by performing Southern hybridization on a portion of the gel using a specific probe made from the pWB15 plasmid. This identified the pulse field gel slice containing TerB sequences, which was excised and melted (20 min at about 72°C). The DNA in the gel solution was stretched on microscope slides coated with 3-aminopropyltriethoxysilane, denatured with sodium hydroxide in ethanol, fixed with glutaraldehyde, and hybridized overnight with biotinylated DNA FISH probes at 37°C in a humidified chamber. Biotinylated DNA FISH probes were made from the fosmid WI2-1478M20 and from the plasmid pWB15. Following hybridization, slides were blocked with 3% BSA for at least 20 min and incubated with the avidin Alexa Fluor 350. This was followed by two rounds of incubation with the biotinylated anti-avidin D for 20 min, followed by the avidin Alexa Fluor 350 for 20 min. Slides were then incubated with an antibody specific for IdU; a mouse anti-BrdU antibody, an antibody specific for CldU; a rat monoclonal anti-BrdU antibody, and the biotinylated anti-avidin D for 1 hr. This was followed by incubation with secondary antibodies: Alexa Fluor 568 goat anti-mouse IgG (H+L), Alexa Fluor 488 goat anti-rat IgG (H+L), and the avidin Alexa Fluor 350 for 1 hr. Slides were rinsed in PBS with 0.03% IGEPAL CA-630 after each incubation. After the last rinse in PBS/CA-630, coverslips were mounted on the slides with ProLong Gold Antifade reagent. A Zeiss fluorescent microscope and IP Lab software (Scanalytics) were used to capture images of IdU/CldU incorporated DNA molecules. Images were processed using Adobe Photoshop and aligned (using Adobe Illustrator) based on the positions of FISH signals that identify the 200 kb segments that contain the ter sequences.

Proximity ligation assay (PLA)

MCF7 5C-TerB cells were seeded on poly-L-lysine-coated coverslips and transfected the day after as described before. Then, 24 hr after transfection, cells were washed with PBS, and fixed and permeabilized with 4% paraformaldehyde containing 0.2% Triton X-100 for 20 min on ice and blocked with PBS-BSA 3% overnight at 4°C. Coverslips were incubated with primary antibodies for 1 hr at room temperature (RT). Proximity ligation was performed using the Duolink In Situ Red Starter Kit Mouse/Rabbit (Sigma-Aldrich) according to the manufacturer’s protocol. The oligonucleotides and antibody-nucleic acid conjugates used were those provided in the Sigma-Aldrich PLA kit. Images were quantified by counting the number of foci per nucleus using Nikon software.

Immunoprecipitation

For immunoprecipitation, MCF7 5C-TerB cells were pre-extracted using CSK100 buffer (100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 10 mM PIPES pH 6.8, 1 mM EGTA, 0.2% Triton X-100, anti-protease and anti-phosphatase) for 5 min on ice, fixed in 1% formaldehyde for 10 min on ice, and a 1% glycine solution was used to stop the reaction. After scraping the cells in ice-cold PBS and centrifugation, pellets were lysed in SDS lysis buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.1% SDS, anti-proteases, and anti-phosphatases) for 10 min on ice and sheared for 3 min. Samples were cleared by centrifugation for 5 min at 4°C. Immunoprecipitations were performed on 10-fold diluted lysates in dilution buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 5 mM EDTA, 0.2% Triton X-100, anti-proteases and anti-phosphatases) with antibodies against GFP, γH2AX, or IgG overnight at 4°C on a wheel. Beads were extensively washed in the dilution buffer and denatured in 2× Laemmli buffer.

Proteins were separated on 4–12% acrylamide SDS-PAGE, transferred on nitrocellulose membrane, and detected with the indicated antibodies described in Supplementary file 1b and ECL reagents.

Cellular fractionation

For the cellular fractionation, MCF7 5C-TerB cells were scraped in PBS, divided into two different tubes (1/3 of the volume for the whole-cell extract [WCE] and the remaining 2/3 for the fractionation) and centrifuged to keep the pellets.

For the WCE, cells were lysed in 1 volume of lysis buffer (50 mM Tris pH 7.5, 20 mM NaCl, 1 mM MgCl2, 0.1% SDS, anti-protease and anti-phosphatase) for 10 min at RT on a wheel and denaturated in 2× Laemmli buffer.

For the fractionation, MCF7 5C-TerB cells were pre-extracted in 2 volumes of CSK100 (100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 10 mM PIPES pH 6.8, 1 mM EGTA, 0.2% Triton X-100, anti-protease and anti-phosphatase) for 15 min on ice, and centrifuged. The supernatant (SN) representing the soluble fraction was kept in a new tube (soluble fraction). The pellet was washed with CSK50 (50 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 10 mM PIPES pH 6.8, 1 mM EGTA, 0.2% Triton X-100, anti-protease and anti-phosphatase) and resuspended in 2 volumes of CSK50 containing benzonase for 10 min on a rotating wheel, and the SN was kept after centrifugation (chromatin fraction). All the fractions were denaturated in 2× Laemmli buffer.

Proteins were separated on 4–12% acrylamide SDS-PAGE, transferred on nitrocellulose membrane, and detected with the indicated antibodies described in Supplementary file 1b and ECL reagents.

Chromatin immunoprecipitation assay

ChIPs were performed using the ChIP-IT express according to the manufacturer’s instructions. Cells were crosslinked, washed, harvested by scraping, and lysed according to the kit’s instructions. Chromatin was sonicated to 200–1500 bp fragments and DNA concentration determined. Equal amount up to 15 ug of DNA was then used for each immunoprecipitation. The antibodies used are listed in Supplementary file 1b. DNA fragments were eluted, purified, and analyzed by SYBR Green real-time PCR. The sequences of primers used for qPCR are given in Supplementary file 1b. Experiments were repeated at least three times, and each real-time qPCR reaction was performed in duplicate.

Detection of DSBs using the T7 endonuclease assay

MCF7 5C-TerB cells were seeded and transfected the day after with 120 pmol of Cas9 protein and 120 pmol of the sgRNA TerB1 using the Lipofectamine CRISPRMAX Cas9 transfection reagents (previously described). Then, 24 hr post-transfection, cells were lysed using a lysis buffer (100 mM NaCl, 10 mM Tris-HCl pH 8, 25 mM EDTA pH 8, 0,5% SDS) and 50 ug of Proteinase K overnight at 50°C with shaking. After addition of NaCl and mix for 1 min, the supernatant was put in a new tube and an ethanol precipitation was performed. A PCR to amplify the 900 bp region surrounding the sgRNA TerB1 site was conducted using PP9F and R. Heteroduplex were formed by heating and cooling down the samples, and a T7 endonuclease assay digestion was performed prior to electrophoresis for detection.

Cell cycle progression

MCF7 5C-TerB cells were transfected as described before. Control cells were treated with 2 mM HU for 4 hr before harvesting. Cells were washed with PBS and fixed in 1 ml cold 70% ethanol for 30 min. Cells were pelleted and washed with PBS. Then, 100 µg/ml RNase A was added and 50 µg/ml PI solution was added directly to the pellet, mixed well, and incubated for 30 min at RT in the dark. A total of 50,000 cells per condition were analyzed by flow cytometry.

Immunofluorescence (γH2AX)

Cells were fixed with 4% PFA/PBS for 20  min, permeabilized with 0.5% Triton-X 100 for 10  min, washed three times in 1× PBS, and blocked in 10% goat serum overnight at 4°C. The primary antibody (1:500) was incubated for 2 hr at RT. Cells were then washed three times in 1× PBS. The secondary antibody (1:1500) was incubated for 1 hr at RT in dark. Cells were then washed again three times in 1× PBS. Stained cells were mounted with mounting medium containing DAPI. Slides were imaged at 60× (immersion oil) using Nikon A1 spinning disk confocal microscope.

Image analysis

For PLA and gH2AX experiments, slides were imaged at 60× (immersion oil) with Nikon spinning disk confocal microscope. PLA foci per nucleus and gH2AX foci per nucleus were calculated using Nikon Elements AR Analysis Explorer (version 5.21.03), where DAPI was used as a mask for the nucleus. The number of PLA foci per nucleus was quantified to get the percentage of cells with one or two foci indicating a positive signal at the site-specific replication block. The number of gH2AX foci was counted for each DAPI to obtain the average number of gH2AX foci in each condition.

NGS sequencing of the MCF7 5C-TerB clone

Genomic DNA was extracted from the cells and submitted to Novogene for sequencing. The samples were processed and whole-genome sequencing was performed using their hWGS service pipeline.

Data analysis and Statistics

Statistical analysis for the various experiments was performed using GraphPad Prism version 9.4.0. Results are presented as mean ± standard error of the mean (SEM). A p value of <0.05 by Student’s t-test was considered statistically significant. ns, nonsignificant, *p<0.05, **p<0.005, ***p<0.001.

Data availability

The list of antibodies, plasmids and oligonucleotides used in this study are provided in Supplementary file 1b. WGS sequencing files generated in this study have been deposited at the NCBI BioProject database under accession number PRJNA868342 and are publicly available as of the date of publication. Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

The following data sets were generated

References

Peer review

Joint Public Review:

The authors report the first use of the bacterial Tus-Ter replication block system in human cells. A single plasmid containing two divergently oriented five-fold TerB repeats was integrated on chromosome 12 of MCF7 cells. ChIP and PLA experiments convincingly demonstrate the occupancy of Tus at the Ter sites in cells. Using an elegant Single Molecule Analysis of Replicated DNA (SMARD) assay, compelling data demonstrate the replication block at Ter sites dependent on the presence of the protein. As an orthogonal method to demonstrate fork stalling, ChIP data show the accumulation of the replicative helicase component MCM3 and the repair protein FANCM around the Ter sites. Previous published work from the Scully and Hickson laboratories showed that Ter sites do not perturb replication fork progression and consistently the data show that the observed effects are dependent on expression of the Tus protein. The SMARD data reveal that about one third of the forks are arrested at Tus/Ter but it is unclear for how long forks remain stalled. Fork stalling led to a highly localized gammaH2AX response, as monitored by ChIP using primer pairs spread along the integrated plasmid carrying the Ter sites. This response was shown to be dependent on ATR using the ATR inhibitor VE-822. This contrasts with a single Cas9-induced DSB between the two Ter sites, which causes a more spread gammaH2AX response measured at two sites flanking the DSB. The difference between the DSB and the Tus-induced stall is very significant. Interestingly, despite evidence for ATR activation through the gammaH2AX response, no evidence for phosphorylation of ATR-T1989, CHK1-S345, or RPA2-S33 could be found under fork stalling conditions. The global replication inhibitor hydroxyurea (HU) elicited phosphorylation of ATR-T1989, CHK1-S345, or RPA2-S33. In this context, it would have been of interest to examine if a single DSB in the Ter region leads to phosphorylation of ATR-T1989, CHK1-S345, or RPA2-S33 and cell cycle arrest. The replication inhibitor HU led to an increase in gamma H2AX foci consistent with a global replication stress response. Overall, this is a well written manuscript, and the data provide convincing evidence that the Tus-Ter system poses a site-specific replication fork block in MCF7 cells leading to a localized ATR-dependent DNA damage checkpoint response that is distinct from the more global response to HU or DSBs.

https://doi.org/10.7554/eLife.87357.3.sa1

Author response

The following is the authors’ response to the original reviews.

Public Review:

The authors report the first use of the bacterial Tus-Ter replication block system in human cells. A single plasmid containing two divergently oriented five-fold TerB repeats was integrated on chromosome 12 of MCF7 cells. ChIP and PLA experiments convincingly demonstrate the occupancy of Tus at the Ter sites in cells. Using an elegant Single Molecule Analysis of Replicated DNA (SMARD) assay, convincing data demonstrate the replication block at Ter sites dependent on the presence of the protein. As an orthogonal method to demonstrate fork stalling, ChIP data show the accumulation of the replicative helicase component MCM3 and the repair protein FANCM around the Ter sites. It is unclear whether the Ter sites integrated by a single copy plasmid have any effect on the replication of this region but the data show that the observed effects are dependent on expression of the Tus protein. The SMARD data do not reveal what proportion of forks are arrested at Tus/Ter, or how long the fork delay is imposed. Fork stalling led to a highly localized gammaH2AX response, as monitored by ChIP using primer pairs spread along the integrated plasmid carrying the Ter sites. This response was shown to be dependent on ATR using the ATR inhibitor VE-822. This contrasts with a single Cas9-induced DSB between the two Ter sites, which causes a more spread gammaH2AX response. While this was monitored only at a single distal site, the difference between the DSB and the Tus-induced stall is very significant. Interestingly, despite evidence for ATR activation through the gammaH2AX response, no evidence for phosphorylation of ATR-T1989, CHK1-S345, or RPA2-S33 could be found under fork stalling conditions. The global replication inhibitor hydroxyurea (HU) elicited phosphorylation of ATR-T1989, CHK1-S345, or RPA2-S33. In this context, it would have been of interest to examine if a single DSB in the Ter region leads to phosphorylation of ATR-T1989, CHK1-S345, or RPA2-S33 and cell cycle arrest. It is not shown whether the replication inhibitor HU leads to the same widely spread gamma H2AX response. Overall, this is a well written manuscript, and the data provide convincing evidence that the Tus-Ter system poses a site-specific replication fork block in MCF7 cells leading to a localized ATR-dependent DNA damage checkpoint response that is distinct from the more global response to HU or DSBs.

Author response to public review:

“It is unclear whether the Ter sites integrated by a single copy plasmid have any effect on the replication of this region but the data show that the observed effects are dependent on expression of the Tus protein.”

-The lack of perturbation of the TerB sequence on fork progression has extensively been studied previously in both Willis et al, 2014 and Larsen et. al, 2014. Furthermore, as the detection of the SMARD signal at the TerB sites is dependent on the 7.5kb probe that spans the TerB sites (orange probe, Fig 2B & 2D), it would be impossible to study the effect on replication in this region, with and without the integration of the single copy plasmid.

“The SMARD data do not reveal what proportion of forks are arrested at Tus/Ter, or how long the fork delay is imposed.”

-The percentage of fork stalling at the TerB sites, with and without Tus expression, has been quantified in Figure 2E & 2F. Essentially, 36% forks stall at the TerB block, i.e. 18% of the forks stall in both the 5’ to 3’ (orange) and 3’ to 5’ (blue) direction when the Tus-TerB block is active.

“It is not shown whether the replication inhibitor HU leads to the same widely spread gamma H2AX response.”

-While we have not shown gH2AX accumulation via ChIP after HU treatment, Supplementary Figure 5A & 5B clearly show increased gH2AX foci when the cells are treated with HU, suggesting a global replication stress response that is in stark contrast to the response to Tus-TerB.

Recommendations for the authors:

Lines 78, 95: In the experimental set-up there are two divergent 5-TerB sites in the orientation that is non-permissive for the fork progression notwithstanding the direction. This raises an obvious question: How an intervening (~1kb-long) DNA segment in being replicated? Does it stay under-replicated and then break?

-The reviewers pose an important question about how the intervening sequence flanked by the two TerB sites is replicated, and if this leads to formation of anaphase bridges resulting in breaks. We think this is very plausible and this very question is part of ongoing studies in the lab with the aim to understand how the cell resolves a site-specific block. Unfortunately, this falls outside the scope of the current study.

Also, it is unclear what is meant with non-permissive orientation. This depends on the predominant replication direction. As the construct has Ter repeats in opposite orientation, any direction is non-permissive. These descriptions could be rephrased to avoid confusion

-The text has been edited to clarify this.

Fig 1A: It would be helpful to annotate the map to show the position of each primer relative to the Ter array. Why is there no signal for pp52?

-Figure 1A has the map of the locus with the annotated primer pairs and their relative positions to the TerB array.

-pp52 is positioned beyond the TerB array so binding of the Tus-His protein there is unlikely, confirming the specificity of the Tus binding to only the TerB array and not to the adjacent chromatin.

Figure 1B: Change Tus to Tus-His to make it easier to understand that the anti-His ChIP is targeting Tus. Provide information what normalization method was used in the ChIP experiments.

-Figure 1B has been edited to reflect this change

Line 113: Willis et al. 2014 also worked with chromosomal Ter sites, which should be acknowledged here.

The text has been modified to indicate this. We apologize for the oversight.

Line 126: Define pWB15 and its significance in text.

-The text has been edited to clarify this and mentions pWB15.

Figure 2E, F: Define legend (blue, orange boxes and arrow heads).

-The figure legend corresponding to Figure 2 has a detailed description of the boxes and the arrows.

Figure 3E, 4C: Add map of primers like in Figures 1 and 2.

-The map added to Figures 3 & 4 and text updated.

Figure 4: Showing that the gammaH2AX response is spread like with the single DSB would bolster the conclusion about the difference between a local and global response. Fig 4A, Lane-3: A loading control for the chromatin fraction is missing.

-Measuring gH2AX chromatin spread after global replication stress can be challenging. We have tried to address the question of global and local gH2AX response post replication stress by quantifying gH2AX foci in cells treated with and without hydroxyurea, comparing it with cells that have a functional Tus-TerB block (Supplementary Figure 5A& 5B). A single fork block seems to only elicit a local response while a global replication stress leads to gH2AX accumulation globally in the cell.

-Lamin A/C has been added to Fig 4A as a loading control for the chromatin fraction.

Figure S4: Analyzing ATR, CHK1 and RPA phosphorylation as well as cell cycle profile under single DSB condition may reveal that different localized responses exist. I mention this because it was reported in yeast that a single DSB in G1 cells leads to a similarly localized Mec1 (ATR) -dependent response that does not elicit phosphorylation of Rad53 (CHK1) and other downstream targets, but leads to H2A phosphorylation as well as phosphorylation of RPA and the Rad51 paralog Rad55 (see PMCID: PMC2853130). It might be of interest to the reader to discuss this publication and the commonalities and differences between both localized checkpoint response

-The reviewers raise an interesting question about the phosphorylation of ATR/CHK1/RPA and its effect on cell cycle after a single DSB. The aim of using the Cas9 break site in this study was merely to corroborate previously published observations pertaining to the spread of gH2AX after a DSB and to contrast that with the local response seen with Tus-TerB. Thus, while an intriguing question, we do not think this particular experiment will help in the understanding of the localized checkpoint response after a single replication fork block. However, we have included the observations previous published in the yeast system (PMC2853130) in our discussion as it helps compare and contrast fork blocks and DSBs further. It is of worth though that the yeast studies were looking at the cellular response to a DSB in G1.

Lines 256-260: In the discussion of ATRIP, unpublished data are discussed that show no increase in ssDNA. What is the effect of ATRIP depletion? Maybe delete this mention of unpublished data, if no new data can be provided. The authors are aware that this makes the mechanism of ATR activation at the 5-TerB site elusive.

-This statement has been deleted and the text has been modified.

Another possibility discussed by the authors is fork reversal. Since Tus/Ter complex block the CMG progression, fork reversal would result in a chicken foot structure with the long single-stranded 3'-overhang of an Okazaki fragment site. Such a structure should be protected by BRCA2 or RAD52 proteins from degradation. Any role for these proteins in the checkpoint activation at the TerB site?

-The reviewers suggest an interesting scenario where the Tus-TerB block induced reversed fork structure could be protected by the loading of known DNA repair proteins and this in turn could lead to a signaling mechanism and checkpoint activation. While we have not tested this hypothesis, nor studied the temporal dynamics of the formation if the reversed fork with respect to gH2AX accumulation, we think the localized gH2AX signal observed in the vicinity of the block is what initiates the downstream DDR response, promoting fork stabilization, followed either by fork reversal and restart or fork collapse. If the reversed fork was responsible for the gH2AX signaling, one would envision the spread to be more widespread, perhaps decorating the entire stretch of DNA between the block and the reversed fork. However, further studies are warranted to tease out this mechanism and the spatio-temporal dynamics.

Lines 292-294: The authors state that "unpublished work from our laboratory has demonstrated that replication forks are cleaved at or near the TerB site..." Unless the data are shown, it might be best to eliminate discussion of unpublished work, also because the occurrence of DNA ends at Ter sites was already described in Willis et al. 2017.

-The statement has been deleted and Willis et al. 2017 has been referenced.

Suppl Table 1: It would help to also show representative images of stretched fibers in addition to the summary data shown.

-Since the data is negative, the fiber images do not show any discernible differences and we do not think it adds useful information.

Suppl Fig 4. ChIP for gamma H2AX data. It would be helpful to show the distribution of the gamma H2AX signal along the chromosome for both the DSB response and the Tus/Ter response.

-The gH2AX ChIP signal at PP0-2 and PP10 has been included in Supplementary Fig4D. Though not significant for PP0-2, the data strongly suggests that there is increased spread of gH2AX along the chromosome after a DSB, strongly contrasting with the response after Tus-TerB block. The text has been modified to include both primer pairs.

https://doi.org/10.7554/eLife.87357.3.sa2

Article and author information

Author details

  1. Sana Ahmed-Seghir

    Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft
    Contributed equally with
    Manisha Jalan
    Competing interests
    No competing interests declared
  2. Manisha Jalan

    Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    Contribution
    Conceptualization, Formal analysis, Validation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Sana Ahmed-Seghir
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4467-4934
  3. Helen E Grimsley

    Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    Contribution
    Formal analysis, Methodology, Writing - original draft
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3485-0305
  4. Aman Sharma

    Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    Contribution
    Formal analysis, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1017-8307
  5. Shyam Twayana

    Department of Cell Biology, Albert Einstein College of Medicine, New York, United States
    Contribution
    Formal analysis, Methodology
    Competing interests
    No competing interests declared
  6. Settapong T Kosiyatrakul

    Department of Cell Biology, Albert Einstein College of Medicine, New York, United States
    Contribution
    Formal analysis, Methodology
    Competing interests
    No competing interests declared
  7. Christopher Thompson

    Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    Contribution
    Methodology
    Competing interests
    No competing interests declared
  8. Carl L Schildkraut

    Department of Cell Biology, Albert Einstein College of Medicine, New York, United States
    Contribution
    Formal analysis, Supervision, Funding acquisition, Methodology, Project administration, Writing - review and editing
    For correspondence
    carl.schildkraut@einsteinmed.edu
    Competing interests
    No competing interests declared
  9. Simon N Powell

    1. Department of Radiation Oncology and the Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    2. Molecular Biology Program, Memorial Sloan Kettering Cancer Center, New York, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Investigation, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    powells@mskcc.org
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8183-4765

Funding

National Institutes of Health (R01-CA187069)

  • Simon N Powell

National Institutes of Health (P50-CA247749)

  • Simon N Powell

National Institutes of Health (R01-GM045751)

  • Carl L Schildkraut

National Institutes of Health (R01-CA085344)

  • Carl L Schildkraut

National Cancer Institute (P30-CA008748)

  • Simon N Powell

National Cancer Institute (P30-CA013330)

  • Carl L Schildkraut

National Institutes of Health (T32AG023475)

  • Shyam Twayana

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are indebted to Ino de Bruijn, Jorge S Reis-Filho, and Yingjie Zhu for help with the genomics data. We thank the members of the Powell and Schildkraut lab for their comments on the manuscript. This work was supported in part by NIH grants R01-CA187069 and P50-CA247749 (to SNP); R01-GM045751 and R01-CA085344 (to CLS); the National Cancer Institute Cancer Center Support Grants, P30-CA008748 at MSK, and P30-CA013330 for use of a core facility at Albert Einstein COM. ST was supported by NIH Training Grant T-32 NIH T32AG023475. Figures were created using BioRender.com

Senior Editor

  1. Kevin Struhl, Harvard Medical School, United States

Reviewing Editor

  1. Wolf-Dietrich Heyer, University of California, Davis, United States

Version history

  1. Preprint posted: March 26, 2023 (view preprint)
  2. Sent for peer review: March 26, 2023
  3. Preprint posted: May 23, 2023 (view preprint)
  4. Preprint posted: August 21, 2023 (view preprint)
  5. Version of Record published: August 30, 2023 (version 1)

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You can cite all versions using the DOI https://doi.org/10.7554/eLife.87357. This DOI represents all versions, and will always resolve to the latest one.

Copyright

© 2023, Ahmed-Seghir, Jalan et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Sana Ahmed-Seghir
  2. Manisha Jalan
  3. Helen E Grimsley
  4. Aman Sharma
  5. Shyam Twayana
  6. Settapong T Kosiyatrakul
  7. Christopher Thompson
  8. Carl L Schildkraut
  9. Simon N Powell
(2023)
A local ATR-dependent checkpoint pathway is activated by a site-specific replication fork block in human cells
eLife 12:RP87357.
https://doi.org/10.7554/eLife.87357.3

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https://doi.org/10.7554/eLife.87357

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    1. Chromosomes and Gene Expression
    Rachel A Johnston, Katherine A Aracena ... Jenny Tung
    Research Advance

    Previously, we showed that a massively parallel reporter assay, mSTARR-seq, could be used to simultaneously test for both enhancer-like activity and DNA methylation-dependent enhancer activity for millions of loci in a single experiment (Lea et al., 2018). Here, we apply mSTARR-seq to query nearly the entire human genome, including almost all CpG sites profiled either on the commonly used Illumina Infinium MethylationEPIC array or via reduced representation bisulfite sequencing. We show that fragments containing these sites are enriched for regulatory capacity, and that methylation-dependent regulatory activity is in turn sensitive to the cellular environment. In particular, regulatory responses to interferon alpha (IFNA) stimulation are strongly attenuated by methyl marks, indicating widespread DNA methylation-environment interactions. In agreement, methylation-dependent responses to IFNA identified via mSTARR-seq predict methylation-dependent transcriptional responses to challenge with influenza virus in human macrophages. Our observations support the idea that pre-existing DNA methylation patterns can influence the response to subsequent environmental exposures—one of the tenets of biological embedding. However, we also find that, on average, sites previously associated with early life adversity are not more likely to functionally influence gene regulation than expected by chance.

    1. Chromosomes and Gene Expression
    Masaaki Sokabe, Christopher S Fraser
    Insight

    A new in vitro system called Rec-Seq sheds light on how mRNA molecules compete for the machinery that translates their genetic sequence into proteins.