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A genome-wide screen identifies SCAI as a modulator of the UV-induced replicative stress response [1]

['Jean-François Lemay', 'Centre De Recherche', 'De L Hôpital Maisonneuve-Rosemont', 'Montréal', 'Québec', 'Edlie St-Hilaire', 'Daryl A. Ronato', 'Genome Stability Laboratory', 'Chu De Québec Research Center', 'Oncology Division']

Date: 2022-10

Helix-destabilizing DNA lesions induced by environmental mutagens such as UV light cause genomic instability by strongly blocking the progression of DNA replication forks (RFs). At blocked RF, single-stranded DNA (ssDNA) accumulates and is rapidly bound by Replication Protein A (RPA) complexes. Such stretches of RPA-ssDNA constitute platforms for recruitment/activation of critical factors that promote DNA synthesis restart. However, during periods of severe replicative stress, RPA availability may become limiting due to inordinate sequestration of this multifunctional complex on ssDNA, thereby negatively impacting multiple vital RPA-dependent processes. Here, we performed a genome-wide screen to identify factors that restrict the accumulation of RPA-ssDNA during UV-induced replicative stress. While this approach revealed some expected “hits” acting in pathways such as nucleotide excision repair, translesion DNA synthesis, and the intra-S phase checkpoint, it also identified SCAI, whose role in the replicative stress response was previously unappreciated. Upon UV exposure, SCAI knock-down caused elevated accumulation of RPA-ssDNA during S phase, accompanied by reduced cell survival and compromised RF progression. These effects were independent of the previously reported role of SCAI in 53BP1-dependent DNA double-strand break repair. We also found that SCAI is recruited to UV-damaged chromatin and that its depletion promotes nascent DNA degradation at stalled RF. Finally, we (i) provide evidence that EXO1 is the major nuclease underlying ssDNA formation and DNA replication defects in SCAI knockout cells and, consistent with this, (ii) demonstrate that SCAI inhibits EXO1 activity on a ssDNA gap in vitro. Taken together, our data establish SCAI as a novel regulator of the UV-induced replicative stress response in human cells.

Funding: This work was supported by the following Canadian Institutes of Health Research (CIHR) grants: 201709PJT-388346 to H.W., 201603PJT-364096 to E.A.D., MOP-133442 to F.A.M. and FDN-388879 to J.Y.M., as well as by the National Sciences and Engineering Research Council of Canada: RGPIN-2019-05082 to H.W. and RGPIN-2018-05414 to F.M.B. J.-F.L. is a recipient of a CIHR post-PhD fellowship. H.W. and F.M.B. are each recipients of a Fonds de la recherche du Québec-Santé Senior scholarship. J.Y.M. is a Canada Research Chair in DNA repair and Cancer Therapeutics. F.A.M. is a Canada Research Chair in Epigenetics of Aging and Cancer. Y.G. was supported by Fondation du CHU de Québec and FRQS PhD scholarships. D.A.R. and C.S. obtained FRQS PhD scholarships. C.S. received a PhD scholarship from the Cole Foundation. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Several mechanisms have been shown to generate ssDNA in response to replicative stress and DNA damage: (1) During S phase, blockage of DNA polymerases causes their uncoupling from the MCM replicative helicase, which continues to unwind DNA ahead of the stalled RF, resulting in abnormally large tracts of ssDNA [ 20 ]. (2) Formation of reversed RF (5) creates nascent DNA ends that can be substrates for degradation by nucleases, e.g., MRE11 and EXO1, thereby generating ssDNA [ 21 ]. Unchecked nascent DNA degradation, termed “replication fork protection defect,” is prevented by several replicative stress response factors, including Rad51 and BRCA1/2 [ 22 ]. (3) Defects in RF reversal or lesion bypass, e.g., TLS, can increase usage of PRIMPOL-dependent repriming downstream of the lesion, which generates ssDNA “gaps” behind RFs (6). (4) Following UV exposure, excision of lesion-containing oligonucleotides during NER transiently generates short stretches of ssDNA, which can be extended by the EXO1 nuclease to promote ATR activation [ 23 ].

Following genotoxin exposure, single-stranded DNA (ssDNA) generated at stalled RF is avidly bound by heterotrimeric Replication Protein A (RPA) complexes [ 7 ]. This not only protects the ssDNA from degradation, but such RPA-bound ssDNA (hereafter RPA-ssDNA) also signals rapid activation of ataxia telangiectasia–mutated (ATM) and Rad3-related (ATR) kinase, the master regulator of intra-S phase checkpoint signaling [ 8 , 9 ]. ATR phosphorylates a multitude of substrates that cooperate to mitigate DNA replication stress by (i) forestalling excessive accumulation of ssDNA at, and stabilizing, stalled RF [ 1 , 10 ] and (ii) preventing further RF blockage by repressing the activation of new origins of replication [ 7 , 11 ]. In addition, RPA is recruited to all active replication origins and advancing RF in the absence of genotoxic insult, where it coats/protects ssDNA resulting from normal minichromosome maintenance (MCM) helicase activity [ 12 ]. In view of the above, maintaining an adequate supply of RPA during S phase, irrespective of whether or not cells are exposed to DNA damaging agents, is essential for timely completion of DNA synthesis [ 13 ]. Lack of ATR activity leading to unrestrained origin firing causes abnormally elevated formation of RPA-ssDNA, which, in turn, engenders progressive exhaustion of the available nuclear pool of RPA and eventual formation of lethal DSB at RF in a phenomenon termed “replication catastrophe” [ 14 ]. Moreover, as RPA is also strictly required for NER [ 15 ], conditions that promote inordinate sequestration of RPA at stalled RF and/or at aberrantly activated replication origins post-UV were shown by our lab and others to cause S phase–specific defects in the removal of UV-induced DNA photoproducts [ 16 – 19 ].

A variety of ubiquitous environmental genotoxins and chemotherapeutic drugs generate helix-destabilizing DNA adducts, e.g., solar UV-induced cyclobutane pyrimidine dimers (CPDs) and 6–4 pyrimidine-pyrimidone photoproducts (6-4PPs). If not efficiently removed by nucleotide excision repair (NER), these adducts block the progression of advancing replicative DNA polymerases. This, in turn, creates a state of “DNA replication stress” that precludes timely completion of S phase with potential genotoxic and carcinogenic consequences [ 1 ]. In order to alleviate these outcomes, i.e., to promote DNA synthesis restart, cells can enlist any among multiple DNA damage tolerance pathways to bypass replication-blocking lesions, including (i) error-free homologous recombination-dependent template switching [ 2 ], or (ii) error-prone translesion synthesis (TLS) following recruitment of specialized DNA polymerases to stalled replication forks (RFs) [ 3 ]. In addition, Rad51-dependent RF reversal can promote reannealing of nascent DNA [ 4 , 5 ]. This brings replication-blocking lesions back into double-stranded DNA, thereby providing an opportunity to repair the lesion prior to eventual resumption of normal DNA replication. In addition, recent evidence demonstrates that repriming beyond damaged bases can be used to allow continuation of DNA RF progression [ 6 ].

We next tested the impact of SCAI on ssDNA gap formation using S1 nuclease DNA fiber assays. We found that S1 nuclease-dependent reduction in the size of DNA labeled post-UV, which reflects cleavage at ssDNA gaps [ 57 ], is unchanged in SCAI-depleted versus control cells ( Fig 7D and 7E ), suggesting that SCAI does not influence the frequency of ssDNA gap generation. Finally, we found that siRNA-mediated depletion of PRIMPOL, which promotes repriming and postreplicative gap formation after genotoxic stress [ 58 ], rescued ssDNA-RPA accumulation post-UV in SCAI-depleted cells ( Fig 7F–7H ). While the above data do not exclude that SCAI might modulate TLS post-UV, they suggest that this protein acts to limit EXO1-dependent nucleolytic extension of ssDNA gaps that are formed as a consequence of repriming downstream of replication-blocking lesions ( Fig 7I ).

Lack of TLS has been shown to favor alternative pathways to tolerate DNA damage during S phase, including PRIMPOL-dependent repriming [ 6 , 51 ]. Interestingly, recent data indicate that SCAI interacts with the REV3L subunit of TLS polymerase zeta [ 55 , 56 ], raising the possibility that REV3L-dependent TLS defects might contribute to the phenotypes of SCAI by favoring the generation of ssDNA gaps post-UV upon which EXO1 can act. Concordant with the above, we found that siRNA-mediated depletion of EXO1 rescued ssDNA-RPA accumulation in cells lacking either POLH, REV1, or REV3L ( S7A–S7I Fig ). However, our data also indicate that siRNA-mediated depletion of both SCAI and REV3L produces additive effects on ssDNA-RPA accumulation post-UV compared to the situation for depletion of either factor alone ( S7J–S7L Fig ). While this result does not exclude the possibility that dysregulation of REV3L may contribute in part to the phenotypes of cells lacking SCAI, it is nonetheless compatible with the notion that SCAI does have REV3L-independent roles post-UV, e.g., modulation of EXO1-dependent ssDNA gap extension.

(A) Recombinant SCAI protein was purified from insect cells, separated by SDS-PAGE and visualized by Coomassie blue staining. ( B) SCAI preferentially binds ssDNA over dsDNA. 5′-[ 32 P]-labeled ssDNA, dsDNA, splayed arm, or gapped DNA were incubated with purified recombinant SCAI at increasing concentrations and the reaction products separated by acrylamide gel electrophoresis and visualized by autoradiography (see S6A Fig ). Quantification of the percentage of SCAI-mediated DNA binding on ssDNA, dsDNA, and splayed arm substrates from 3 independent experiments. ( C) Left: in vitro DNA resection assays using a 3′-[ 32 P]-labeled gapped DNA substrate in the absence of any proteins, or with WT or a catalytically inactive version of EXO1 (D173A) supplemented with purified recombinant SCAI. Right: quantification of the percentage of DNA resection from 3 independent experiments. ( D, E) Depletion of SCAI does not increase ssDNA gap generation post-UV. ( D) Schematic of the DNA fiber assay used to assess RF progression post-UV. Cells were incubated with CldU (red) for 30 min, irradiated with UV (20 J/m 2 ), and then incubated with IdU (green) for 90 min. Cells were then treated or not with S1 nuclease. ( E) Left panel: dot plot and median (red line) of IdU/CldU ratio from cells transfected with siRNAs as indicated. Middle panel: dot plot and median of IdU tract lengths (DNA fiber dot plot are combined from n = 3 with similar result). Right panel: histogram of median values of IdU track length derived from independent experiments. Means and SEM are plotted as lines and whiskers. ( F-H) Depletion of PrimPol rescues ssDNA-RPA accumulation in cells lacking SCAI. ( F) Validation of siRNA-mediated KD of PrimPol and SCAI by immunoblot. ( G) Representative immunofluorescence flow cytometry plots from cells transfected with the indicated siRNA treated with 1 J/m 2 UV or mock-treated and allowed to recover for 6 h. The dashed red box delineates DNA-bound RPA high cells. ( H) Quantification from (G). Histogram values represent the mean ± SEM from 3 independent experiments. (I) Proposed model. After UV exposure, PrimPol-dependent repriming generates ssDNA gaps behind RF. These gaps recruit RPA and SCAI, and SCAI acts to restrain the resection activity of EXO1. Possible modulation of Polζ/REV3L-dependent TLS by SCAI cannot be excluded. Statistics used: Kruskal–Wallis with Dunn’s multiple comparisons test (E; left and middle panels), unpaired t test corrected for multiple comparisons using the Holm–Šídák method (E; right panel, H). ns: nonsignificant, *: p ≤ 0.05, **: p ≤ 0.01, ****: p ≤ 0.0001. The data underlying the graphs shown in the figure can be found in S1 Data . KD, knockdown; RF, replication fork; RPA, Replication Protein A; SEM, standard error of the mean; ssDNA, single-stranded DNA; TLS, translesion synthesis; WT, wild type.

EXO1 has been shown to extend ssDNA gaps left behind RF during replicative bypass of damaged DNA bases via PRIMPOL-dependent repriming [ 52 ]. Previously published data also suggest that SCAI possesses the capacity to bind ssDNA [ 38 ], raising the possibility that SCAI might directly influence the activity of EXO1 at ssDNA gaps. We purified SCAI and assessed its ability to bind various ssDNA-containing substrates in vitro ( Fig 7A ). Our data indicate that SCAI readily binds linear ssDNA or “splayed arm” ssDNA-containing structures but displays much lower affinity for dsDNA (Figs 7B and S6A ). Interestingly, SCAI binds to a splayed arm substrate containing 44 but not 30 nt of ssDNA (Figs 7B and S6B and S6C ), suggesting that the length of ssDNA influences the ability of SCAI to bind DNA. We further found that SCAI can bind a substrate containing a 34 base-long ssDNA gap ( Fig 7B ) and that incubation with a concentration of SCAI permitting the detection of such binding leads to significantly reduced EXO1 nucleolytic activity on this substrate ( Fig 7C ). In contrast, SCAI was unable to restrict EXO1 activity on the splayed arm substrate despite the ability to bind it ( S6D and S6E Fig ), raising the possibility that inhibition of this nuclease by SCAI requires a ssDNA/dsDNA junction, a structure present at both reversed forks and ssDNA gaps.

(A) Depletion of EXO1, and to a lesser extent MRE11, rescues RPA-ssDNA accumulation post-UV in cells lacking SCAI. Cells were treated with 1 J/m 2 UV or mock-treated and allowed to recover for 6 h. The dashed red box delineates DNA-bound RPA high cells. ( B) Quantification from (A). Values represent the mean ± SEM from 3 independent experiments. ( C) Immunoblot analysis showing EXO1 or MRE11 depletions from whole-cell extracts from U-2 OS (WT) or SCAI-KO (#1) cells transfected with siRNAs. ( D) Top: schematic of the DNA fiber assay used to assess RF progression post-UV. Cells were incubated with CldU (red) for 15 min, irradiated with UVC (20 J/m 2 ), and then incubated with IdU (green) for 60 min. Bottom: dot plot of IdU/CldU ratio and median (red line) from U-2 OS transfected with siNT or siRNA against EXO1 (data combined from n = 2 with similar results). ( E) Top: schematic of the DNA fiber assay to monitor RF protection defects (nascent DNA degradation) after HU. Cells were incubated successively with CldU (red) and IdU (green) for 20 min each and then exposed to 4 mM HU for 4 h. Bottom: dot plot of IdU/CldU ratio and median (red line) from U-2 OS (WT) and SCAI-KO (#1) cells transfected with siRNAs against BRCA2 (data combined from n = 2 with similar results). ( F) Similar experiment as in (D) but from U-2 OS transfected with siNT or siRNA against BRCA2 (data combined from n = 2 with similar results). ( G-I) Lack of BRCA1/2 does not cause RPA-ssDNA accumulation under our experimental conditions. (G) Validation of BRCA1 and BRCA2 KDs by immunoblot. ( H) Experiments were performed as in (A) but with cells transfected with siRNAs against BRCA1 or BRCA2 +/− siSCAI. ( I ) Quantification from (H). Values are the mean ± SEM from 3 independent experiments. Statistics used: unpaired t test corrected for multiple comparisons using the Holm–Šídák method (B), two-tailed unpaired Student t test (I), Kruskal–Wallis with Dunn’s multiple comparisons test (D-F). ns: nonsignificant, **: p ≤ 0.01, ***: p ≤ 0.001, ****: p ≤ 0.0001. The data underlying the graphs shown in the figure can be found in S1 Data . HU, hydroxyurea; KD, knockdown; KO, knockout; RF, replication fork; RPA, Replication Protein A; siNT, nontargeting siRNA; siSCAI, SCAI-targeting siRNA; ssDNA, single-stranded DNA; WT, wild type.

We next tested directly whether RF protection defects, i.e., degradation of nascent DNA at reversed forks, contribute to the accumulation of RPA-ssDNA post-UV under our experimental conditions. As mentioned above, BRCA1/2 are well known to contribute to the protection of nascent DNA at stalled RF [ 21 , 54 ]. However, our screen did not identify BRCA1/2 ( S1 Table ), and, moreover, cells in which either of these proteins was depleted, either alone or in combination with SCAI, did not display significant elevation of RPA-ssDNA ( Fig 6G–6I ). Intriguingly, while depletion of BRCA2 has no significant effect on DNA-bound RPA levels post-UV, we found that lack of BRCA1 strongly suppressed ssDNA-RPA accumulation in S phase cells under our experimental conditions ( Fig 6G–6I ). While the basis for the latter observation is unclear, overall, our results indicate that nascent DNA degradation is unlikely to contribute to UV-induced accumulation of RPA-ssDNA in cells lacking SCAI.

Several nucleases, including EXO1 and MRE11, generate ssDNA at stalled RF [ 22 , 50 ]. Moreover, recent reports indicate that replicative stress leads to the formation of unreplicated ssDNA gaps behind forks, which can be extended by EXO1 and MRE11 [ 6 , 51 , 52 ]. We therefore tested whether these nucleases might promote RPA-ssDNA formation in cells lacking SCAI. Strikingly, accumulation of DNA-bound RPA post-UV was abrogated upon siRNA-mediated depletion of EXO1 in SCAI-KO cells, whereas the effect of MRE11 was more modest (Figs 6A–6C and S5A ). Based on this, we focused further characterization on the relationship between SCAI and EXO1-dependent DNA degradation and found that EXO1 KD rescues UV-induced RF progression defects caused by lack of SCAI (Figs 6D and S5B ). We reasoned that depletion of SCAI might favor EXO1-dependent nucleolytic degradation of nascent DNA at stalled RF [ 50 ], leading to reduction in RF progression and accumulation of RPA-ssDNA. Consistent with this, we found that cells lacking SCAI displayed nascent DNA instability comparable to that caused by depletion of BRCA2 (Figs 6E and S5C ) [ 21 ]. Codepletion of both factors caused an additive effect with regard to RF protection (Figs 6E and S5C ), suggesting that SCAI and BRCA2 act via distinct mechanisms to protect stalled RF from nucleolytic degradation. While we observed a similar trend with respect to RF progression after UV, the additive effect caused by codepletion of BRCA2 and SCAI versus either of these proteins alone did not reach statistical significance (Figs 6F and S5D ). More experiments will therefore be necessary to firmly ascertain whether these proteins act independently or redundantly in the context of UV-induced replicative stress. Taken together, the above data suggest that nascent DNA degradation is likely to contribute to the observed RF progression defects during genotoxic stress in cells lacking SCAI. Since RF protection/progression defects have been associated with sensitivity to genotoxins [ 53 ], we tested whether depletion of EXO1 might restore UV resistance in cells lacking SCAI and found that this is not the case ( S5E Fig ). It is therefore possible that EXO1-independent roles of SCAI, e.g., in DNA DSB repair [ 38 , 39 ], gene regulation [ 40 ], or heretofore unknown mechanisms, may have an overriding influence on UV sensitivity in SCAI-null cells. For example, lack of EXO1 may generate DNA lesions post-UV, e.g., DSB, which require SCAI for their processing.

We next sought to investigate whether SCAI can be found in close proximity to DNA repair or replicative stress proteins in vivo. To this end, we used a variation of the BioID assay (TurboID) coupled to mass spectrometry, which permits rapid biotinylation, purification, and mass spectrometry–based identification of proteins in close spatial proximity to a protein of interest ( S2 Table ) [ 47 , 48 ]. Consistent with a previous report indicating a role for SCAI in modulating transcription [ 40 ], our analysis revealed that proteins involved in gene expression and chromatin organisation are biotinylated by TurboID-SCAI both in untreated and UV-exposed cells ( Fig 5I–5K and S2 Table ). As expected, several peptides of the known SCAI-interacting DNA repair protein 53BP1 [ 38 , 39 ] were also recovered; however, the abundance of 53BP1 peptides in BioID control data sets from the Contaminant Repository for Affinity Purification database (CRAPome v2.0) [ 49 ] led to its exclusion from confirmed “hits” in our analyses. Importantly, our analyses identified several other proteins involved in DNA repair/replicative stress responses (e.g., REV1, REV3L, MCM10, BRCA2, PLK1, SLX4, UBR5, CLASPIN) as being in close physical proximity to SCAI ( Fig 5K ). Such proximity was observed in both UV- and mock-treated cells; while the reason for this is currently unknown, it is possible that either (i) SCAI is constitutively found in proximity/complex with the abovementioned proteins or that (ii) spontaneous DNA lesions formed in U-2 OS cells are sufficient to allow the detection of damage-induced SCAI-containing complexes. In any case, the above data support the notion that SCAI is localized in the vicinity of replicative stress response and DNA repair proteins in U-2 OS cells.

Biochemical purification of newly replicated DNA using iPOND did not identify SCAI as a component of stalled RF [ 43 ]. Nevertheless, it remains possible that recruitment of SCAI in the vicinity of RF occurs infrequently or transiently, thereby precluding detection of SCAI using this method. We exploited a cell biology approach relying on the introduction of a 256XLacO array in U-2 OS cells expressing an mCherry-tagged LacR construct [ 44 ]. Recruitment of LacR proteins to the 256XLacOarray has been shown to be associated with induction of replicative stress markers in this genomic region [ 44 – 46 ]. We used cell lines harboring the 256XLacO array and expressing mCherry-LacR and either GFP-SCAI or control GFP. We confirmed that our GFP-SCAI fusion was functional by testing its ability to form nuclear foci in response to IR-induced DSB [ 38 , 39 ] ( S4A Fig ). As expected, we observed elevated signals for chromatin-bound RPA32 (reflecting ssDNA formation) and phosphorylated histone H2AX (generated by ATR and ATM kinase activity, and which can therefore reflect either replicative stress or DSB) at the LacO array in EdU-labeled (S phase) cells ( S4B and S4C Fig ). However, such replication stress/DNA damage markers were also observed in EdU-negative cells ( S4B and S4C Fig ), suggesting that replicative stress-induced DNA lesions formed at the LacO array during S may persist in subsequent phases of the cell cycle. We also observed that the intensity of GFP-SCAI colocalized with the mCherry-LacR-labeled region is significantly higher than for GFP alone in both EdU-positive and EdU-negative cells ( Fig 5D–5F ). Overall, the above data are consistent with the notions that (i) binding of mCherry-LacR to LacO produces localized replicative stress and (ii) that SCAI colocalizes with the LacO array in S phase cells. Since both the colocalization of SCAI with the LacO array and replication stress-induced DNA damage can be observed outside of S phase, we cannot exclude that some recruitment of GFP-SCAI to the LacO array might reflect the activity of this protein after DNA damage has been generated, e.g., during DSB repair or at postreplicative ssDNA gaps. We next tested whether SCAI is recruited to chromatin in cells experiencing UV-induced replicative stress. To this end, we used fluorescence microscopy coupled to stringent washes that remove proteins that are not bound to DNA in cells expressing either GFP alone or GFP-SCAI. We found that UV elevates the binding of SCAI to chromatin to a similar extent as IR in EdU-labeled S phase cells ( Fig 5G ). Importantly, our data also indicate that such significant increases in chromatin-bound SCAI post-UV do not result from elevated expression of this protein ( Fig 5H ). Taken together, the above data are consistent with the notion that SCAI can be recruited to genomic regions experiencing DNA replication stress.

(A) Schematic of the DNA fiber assay used to assess RF progression post-UV. Cells were incubated with CldU (red) for 15 min, irradiated with UV (20 J/m 2 ), and then incubated with IdU (green) for 60 min. ( B) Left panel: dot plot and median (red line) of IdU/CldU ratio from control (siNT or WT) or SCAI-depleted (siSCAI or SCAI-KD) cells. Middle panel: dot plot and median of IdU tract lengths. Right panel: dot plot and median of CIdU tract lengths (data combined from n = 3 with similar result). ( C) 53BP1 does not influence RF progression post-UV in cells lacking SCAI. Similar experiment as in (B) (left panel; data combined from n = 2 with similar result). ( D) SCAI localizes to stalled RFs. Schematic of the assay used to evaluate recruitment of SCAI to stalled RF caused by binding of mCherry-LacR to a LacO array. ( E) Representative microscopy images for the assay described in (D). Scale bar = 10 μM. (F) Quantification of GFP or GFP-SCAI normalized signal intensity in the mCherry-LacR foci in non-S phase cells (EdU−) or S phase cells (EdU+). Each point represents a single cell. Lines represent the median. Data combined from 3 similar biological replicates. ( G) GFP-SCAI associates with DNA post-UV. Signal intensity from S phase cells was determined by flow cytometry +/− irradiation with 2 J/m 2 UV or 5 Gy IR. Cells were allowed to recover for 6 h (UV) or 5 h (IR). Red line represents the mean. Representative results from 3 independent experiments. ( H) Immunoblot analysis of the expression level of GFP-SCAI under the experimental conditions described in (G) ± induction by doxycycline. ( I) Interrogation of proximity interactome was performed through biotin labeling using TurboID-SCAI under untreated and UV-treated (2 J/m 2 ) conditions. Fold-change is relative to the CRAPome background controls (see Materials and methods ). Proteins with a SAINT score ≥0.7 and a BFDR ≤0.05 are considered significant. (J) GO term enrichment analysis of proteins found in the untreated and UV-treated conditions. (K) Proteins associated with the GO term “Cellular response to DNA damage stimulus” are shown as a dot plot in which node color represents the fold increase, node size represents the relative fold change between the experimental conditions, and node edges represent the SAINTexpress BFDR. Raw data are in S2 Table . Statistics used: Mann–Whitney test (B), Kruskal–Wallis with Dunn’s multiple comparisons test (C), two-tailed unpaired Student t test (F), one-way ANOVA corrected for multiple comparisons using Tukey’s test (G). ns: nonsignificant, *: p ≤ 0.05, **: p ≤ 0.01, ***: p ≤ 0.001, ****: p ≤ 0.0001. The data underlying the graphs shown in the figure can be found in S1 Data . a.u., arbitrary units; BFDR, Bayesian false discovery rate; GO, Gene Ontology; IR, ionizing radiation; RF, replication fork; SAINTexpress, Significance Analysis of INTeractome; siNT, nontargeting siRNA; siSCAI, SCAI-targeting siRNA; WT, wild type.

We next assessed the impact of SCAI on RF progression after UV irradiation using DNA fiber analysis. We found that both siRNA-mediated depletion and CRISPR-Cas9 KD of SCAI significantly compromised RF progression post-UV in U-2 OS cells ( Fig 5A and 5B ) as well as in 2 additional cancer cell lines: TOV-21G (ovarian cancer) and WM3248 (melanoma) ( S2C–S2E Fig ). SCAI depletion does not negatively influence RF progression in the absence of genotoxic treatment; in fact, unchallenged cells treated with siSCAI displayed modest increases in RF progression ( Fig 5B ). Our data also indicate that the negative impact of SCAI depletion on RF progression after UV treatment is independent of 53BP1 ( Fig 5C ), as was the case for SCAI-dependent modulation of RPA-ssDNA levels ( Fig 4I–4K ). Overall, these data demonstrate that SCAI influences RF progression after genotoxic stress.

As mentioned previously, SCAI physically interacts with 53BP1 to modulate DSB repair [ 38 , 39 ]. We therefore evaluated whether this functional interaction is relevant in the context of UV-induced RPA-ssDNA accumulation in S phase cells. Compared to the situation for UV, DSB-inducing ionizing radiation (IR) did not cause noticeable accumulation of RPA on DNA in either control or SCAI-depleted cells ( Fig 4G and 4H ), indicating that DSB processing, i.e., end resection, does not cause significant accumulation of RPA-ssDNA in S phase cells under our experimental conditions. We further found that depletion of 53BP1, alone or in combination with that of SCAI, did not influence levels of RPA-ssDNA post-UV in our assay ( Fig 4I–4K ). Taken together, these data indicate that the abnormal response to replicative stress in cells lacking SCAI is unlikely to be related to defective 53BP1-dependent DSB repair.

(A) Immunofluorescence flow cytometry was used to measure repair synthesis-associated EdU incorporation in G1/G2 cells (y-axis) and total DNA content (x-axis; DAPI signal). Cells transfected with siNT, SCAI-, or XPC-targeting siRNAs were irradiated with 20 J/m 2 UV and allowed to recover for 3 h in medium containing 5 μM EdU. The red and blue dashed lines are positioned, respectively, in the middle of the EdU signal of the G1 and G2 cell populations of the siNT-treated cells to facilitate comparison. ( B) Quantification from (A). Values are the mean ± SEM from at least 2 independent experiments and are relative to siNT-treated cells. ( C) Representative images of 5-EU incorporation from cells transfected with the indicated siRNA. Cells were either mock- or UV-treated (6 J/m 2 ) and samples collected 3 and 24 h after irradiation. Scale bar = 20 μM. White arrows indicate cells with reduced incorporation of 5-EU. ( D) Quantification from (C). The red lines represent the median. The assay was repeated twice independently with similar result. ( E) Validation of siRNA-mediated KD of XPA using immunoblot. ( F) Quantification of UV-induced 6-4PP removal as a function of cell cycle using a flow cytometry–based assay from cells transfected with siNT or siSCAI. Values are the mean ± SEM from 3 independent experiments. ( G) Immunofluorescence flow cytometry was used to measure DNA-bound RPA32 (y-axis) and DNA content (x-axis; DAPI signal). Cells were irradiated with either 1 J/m 2 UV or 5 Gy IR and allowed to recover for 6 h prior to sample collection. The dashed red box delineates DNA-bound RPA high cells. Representative flow cytometry plots are shown. ( H) Quantification from (G). Values represent the mean ± SEM from 3 experiments. ( I) Immunoblot analysis from cells transfected with the indicated siRNAs. ( J) siNT-, siSCAI-, si53BP1-, and siSCAI/si53BP1-transfected cells were irradiated with 1 J/m 2 UV and allowed to recover for 6 h before immunofluorescence flow cytometry analysis. The dashed red box delineates DNA-bound RPA high cells. Representative flow cytometry plots are shown. ( J ) Quantification from (I). Values represent the mean ± SEM from 3 independent experiments. Statistics used: two-tailed unpaired Student t test (B, H), two-tailed Mann–Whitney test (D), unpaired t test corrected for multiple comparisons using the Holm–Šídák method (F, I). ns: nonsignificant, *: p ≤ 0.05, **: p ≤ 0.01, ****: p ≤ 0.0001. The data underlying the graphs shown in the figure can be found in S1 Data . DSB, double-strand break; IR, ionizing radiation; KD, knockdown; NER, nucleotide excision repair; siNT, nontargeting siRNA; siSCAI, SCAI-targeting siRNA; 6-4PP, 6–4 pyrimidine-pyrimidone photoproduct.

Several NER genes were recovered in our screen (Figs 1 and 2 ). Indeed, defective removal of UV-induced DNA lesions is expected to exacerbate RF stalling and accumulation of RPA-ssDNA in S phase cells. To address the possibility that SCAI might regulate NER efficiency, we evaluated the DNA repair synthesis step of this pathway by quantifying unscheduled incorporation of the nucleoside analog EdU in G1/G2 cells post-UV [ 41 , 42 ]. As expected, siRNA-mediated depletion of the essential NER factor XPC strongly attenuated repair synthesis compared to cells transfected with nontargeting siRNA (siNT) ( Fig 4A and 4B ). In contrast, EdU incorporation post-UV was not reduced in SCAI-depleted versus control cells, suggesting that the global genomic NER subpathway is not compromised by lack of SCAI ( Fig 4A and 4B ). Similarly, we found that recovery of RNA synthesis post-UV as measured by incorporation of the nucleoside analog EU [ 41 , 42 ], an indicator of the efficiency of the transcription-coupled NER subpathway, was similar in control versus SCAI-depleted cells but clearly defective in cells in which the essential NER factor XPA was knocked down ( Fig 4C–4E ). Finally, we used a flow cytometry–based NER assay, which was originally developed in our lab to directly evaluate repair of UV DNA photoproducts as a function of cell cycle [ 16 ]. We did not observe any significant difference between siNT- and SCAI-targeting siRNA (siSCAI)-treated cells with respect to removal of 6–4 photoproducts in U-2 OS cells (Figs 4F and S3 ). We further note that, consistent with our previously published data [ 16 ], U-2 OS cells are profoundly defective in the removal of UV-induced lesions during S phase ( Fig 4F ). Overall, the above results indicate that lack of SCAI does not cause replicative stress by compromising NER-mediated removal of UV-induced DNA lesions.

Consistent with the elevated formation of RPA-ssDNA observed in Fig 3A–3D , native immunofluorescence of incorporated BrdU, which directly reflects ssDNA accumulation, was elevated in SCAI-depleted S phase cells post-UV as compared to control cells ( Fig 3G ). Exposure to the replicative stress-inducing drugs cisplatin (CDDP) and 4-nitroquinoline 1-oxide (4-NQO) was also found to increase RPA-ssDNA during S phase in SCAI-depleted compared to control cells ( Fig 3H and 3I ). Finally, we found that SCAI-KD and SCAI-KO cells are sensitized to UV and CDDP (Figs 3J and S1E ) and that reexpression of SCAI in KO cells completely rescues their sensitivity to UV ( Fig 3K ). Taken together, these data show that upon exposure to genotoxins that cause replicative stress, SCAI acts to alleviate (i) abnormal accumulation of RPA-ssDNA in S phase cells and (ii) loss of cell viability and/or reduced proliferation.

(A, C) Immunoblot of whole-cell extracts from control U-2 OS, SCAI-depleted cells (A) and SCAI KO cells (C). ( B, D) Immunofluorescence flow cytometry measurements of DNA-associated RPA32 in control, SCAI-depleted cells (B, left plot: siSCAI, right plot: SCAI-KD), or SCAI-KO (D) cells 6 h after 1 J/m 2 UV irradiation. RPA high cells are delineated by a dashed box. ( E, F) Quantification from (B) and (D), respectively. Values are the mean ± SEM from at least 3 independent experiments. (G) Depletion of SCAI increases ssDNA generation post-UV. Control and SCAI-depleted cells were exposed to BrdU for 48 h, and then irradiated with UV as indicated. Native BrdU signal was assessed by immunofluorescence flow cytometry at 6 h post-UV. Median is presented (red line), and error bars indicate the interquartile range. Data combined from 2 independent experiments with similar results. ( H) Immunofluorescence flow cytometry measurements of DNA-associated RPA32 in control and SCAI-depleted cells (as in (B)). U-2 OS transfected with siNT or siSCAI were treated with 0.5 μM 4-NQO for 1 h and allowed to recover for 5 h or continuously exposed for 6 h to 5 μM cisplatin (CPPD). ( I ) Quantification from (H). Values are the mean ± SEM from 3 independent experiments. ( J) SCAI-KD/KO cells are sensitive to UV as measured by clonogenic survival. Values are the mean ± SEM from 3 independent experiments. ( K) Rescue of UV sensitivity of SCAI-KO cells by transient overexpression of SCAI as determined by clonogenic survival. Values are the mean ± SEM from 2 independent experiments. Right: immunoblot of whole-cell extracts from control U-2 OS (WT), SCAI KO cells, and SCAI-KO that transiently overexpress SCAI. Statistics used: two-tailed unpaired Student t test (E, I), unpaired t test corrected for multiple comparisons using the Holm–Šídák method (J, F), and Mann–Whitney (G). *: p ≤ 0.05, **: p ≤ 0.01, ***: p ≤ 0.001, ****: p ≤ 0.0001. The data underlying the graphs shown in the figure can be found in S1 Data . KD, knockdown; KO, knockout; siNT, nontargeting siRNA; siSCAI, SCAI-targeting siRNA; WT, wild type.

The SCAI gene was recovered at multiple time points in our RPA-ssDNA screen ( Fig 1F ). SCAI has been reported to interact with 53BP1 to modulate DSB repair [ 38 , 39 ] and also to influence gene transcription [ 40 ]. However, any effect of SCAI on the response to genotoxin-induced replicative stress was unknown. We found that U-2 OS cells in which SCAI was either knocked down or knocked out via CRISPR-Cas9 (SCAI-KD or SCAI-KO, respectively), or down-regulated using multiple independent siRNA, exhibited elevated RPA-ssDNA post-UV as compared to control cells (Figs 3A–3D and S1A–S1C ). Like other genes identified in our screen, accumulation of RPA on DNA was observed primarily during S phase in cells lacking SCAI (Figs 3B and 3D and S1B ), suggesting that this factor might modulate the response to replicative stress. siRNA-mediated SCAI depletion also caused a similar phenotype in TOV-21G ovarian cancer cells ( S2A and S2B Fig ).

Interestingly, all 3 subunits of the RPA complex were also identified as “hits” in the screen ( S1 Table ). This is consistent with published data indicating that partial siRNA-mediated depletion of RPA, i.e., to levels that do not compromise unchallenged DNA replication, exacerbates replicative stress [ 13 ]. As such, although it might seem counterintuitive, reduction of RPA availability was shown to ultimately elevate DNA-bound RPA in cells treated with hydroxyurea and ATR inhibitor [ 13 ]. We speculate that progressive CRISPR-Cas9-mediated depletion of RPA subunits might lead to a similar situation upon UV irradiation. In other words, while the total level of RPA is decreased due to CRISPR-Cas9 gene inactivation, the proportion of RPA that is DNA-bound is likely to be elevated due to replicative stress post-UV. Taken together, the results indicate that our screening strategy is competent in identifying mediators of the UV-induced DNA replication stress response.

(A) Main functional groups derived from genes recovered in the screen. Genes selected for further validation are shaded in grey. ( B) Representative immunofluorescence flow cytometry assays and immunoblots after siRNA-mediated depletion of selected genes. Cells transfected with nontargeting (siNT) or gene-specific siRNAs were mock- or UV-treated (1 J/m 2 ). % RPA high cells (dashed box) were assessed 6 h after irradiation. ( C) Quantification of (B). Values represent the mean ± SEM of at least 3 independent experiments. Statistics used: unpaired t test corrected for multiple comparisons using the Holm–Šídák method. *: p ≤ 0.05, **: p ≤ 0.01, ***: p ≤ 0.001, ****: p ≤ 0.0001. The data underlying the graph shown in the figure can be found in S1 Data .

We next evaluated siRNA-mediated depletion of individual “hits” from our screen on ssDNA-RPA formation post-UV. Genes from various functional groups were selected ( Fig 2A ). As expected, knockdown (KD) of RAD18, POLH, and XPC caused elevated ssDNA-RPA post-UV ( Fig 2B and 2C ). Our screen also identified factors whose potential roles in the UV-induced replicative stress response are incompletely characterized ( Fig 2B and 2C ): (i) the TRiC chaperonin complex (CCT2 and CCT8 subunits), which possesses several DNA repair/replication proteins as substrates [ 34 ]; (ii) the RUVBL1 chromatin remodeler, recently suggested to play roles in modulating the replicative stress response [ 35 ]; and (iii) RIF1, a DNA double-strand break (DSB) repair factor that also regulates DNA replication origin activity [ 36 , 37 ]. Down-regulation of the above factors caused elevation in RPA-ssDNA specifically in S phase cells, consistent with the notion that most of the genes recovered in our screen act by mitigating replicative stress. We note that siRNA against RIF1 also caused elevated RPA-ssDNA in the absence of UV, which might reflect the role of this gene in negatively regulating the activation of DNA replication origins in unperturbed cells [ 36 ].

We found that sgRNA associated with the RPA high fraction changed from day 6 to 15 ( Fig 1F and S1 Table ), likely reflecting loss of sgRNA targeting essential and growth-promoting genes from the cell populations. Nevertheless, several genes were recovered at more than one time point ( Fig 1F ). Seven genes recovered at every time point encode factors with previously documented roles in the response to replicative stress and/or UV-induced DNA damage, as follows: RFWD3, a ubiquitin ligase that regulates both TLS and RPA recruitment to stalled RFs [ 29 , 30 ]; DNA polymerase eta, a TLS polymerase that mediates bypass of UV-induced CPD (3); RAD18, a PCNA ubiquitin ligase involved in DNA damage tolerance [ 31 ]; RAD9, a component of the intra-S phase checkpoint 911 complex [ 32 ]; and the NER pathway proteins XPA and XPC [ 33 ]. Gene Ontology (GO) term analysis of genes identified in our screen returned terms related to known pathways influencing the cellular response to UV-induced replicative stress, including error-prone TLS, nucleotide excision repair, DNA replication, and postreplication repair ( Fig 1G ).

We devised a CRISPR-Cas9 screening strategy employing the genome-wide GeCKOv2 lentiviral library [ 25 , 26 ] in conjunction with the above-described flow cytometry assay ( Fig 1E ). U-2 OS cells were infected with the GeCKOv2 library and propagated for periods of 6, 9, 12, or 15 d to allow phenotypic expression. Cells were then either exposed to 1 J/m 2 UV or mock-treated. At 6 h post-UV, cells were fixed and labeled with anti-RPA32 antibodies followed by FACS to sort RPA high cells (i.e., within the red dotted rectangle in Fig 1A and 1C ). Following extraction of DNA from untreated and RPA high cells, barcode sequences were amplified by PCR, and corresponding guide RNAs (sgRNA) identified by high-throughput sequencing. Results were then analyzed using the MAGeCK pipeline to identify sgRNA that are overrepresented in the RPA high population versus untreated controls [ 27 , 28 ].

(A) Immunofluorescence flow cytometry was used to measure ssDNA-bound RPA32 (y-axis) and total DNA content (x-axis; DAPI signal). Cells were treated with 1, 3, or 5 J/m 2 UV or mock-treated, and samples were collected 1, 3, or 6 h post-UV. The dashed red box delineates DNA-bound RPA high cells. ( B) Quantification from (A). Values are the mean ± SEM from 2 independent experiments. ( C) Proof-of-concept using the ATR inhibitor VE-821. Cells were mock-treated or irradiated with 1 J/m 2 UV +/− 2 μM of VE-821. Samples were harvested 6 h posttreatment. ( D) Quantification from (C). Values are mean ± SEM from 3 experiments. ( E) Schematic overview of the FACS-based CRISPR-Cas9 screen. Cells were irradiated with 1 J/m 2 UV at 6, 9, 12, and 15 d postinfection with the GeCKOv2 lentiviral library (see Materials and methods ). At each time point, mock-treated cells were collected to assess sgRNA representation. ( F) Venn diagram of the distribution of the genes recovered at each time point. ( G) GO term enrichment analysis of genes identified at all time points. Statistics used: unpaired t test corrected for multiple comparisons using the Holm–Šídák method. **: p ≤ 0.01, ***: p ≤ 0.001. The data underlying the graphs shown in the figure can be found in S1 Data . ATR, ATM and Rad3-related; GO, Gene Ontology; RPA, Replication Protein A; ssDNA, single-stranded DNA.

We sought to identify gene networks that restrict RPA accumulation on DNA during genotoxin-induced replicative stress. To this end, we optimized an existing method coupling flow cytometry, stringent washes, and immunofluorescence to measure ssDNA-associated (as opposed to free) RPA32 (one of the 3 subunits of the RPA complex) in U-2 OS human osteosarcoma cells in response to 254 nm UV (hereafter UV; Fig 1A ) [ 24 ]. Exposure to 1 J/m 2 UV caused detectable RPA recruitment to DNA at 1 and 3 h post-UV, which was largely resolved by 6 h ( Fig 1A and 1B ). In contrast, higher UV doses (3 or 5 J/m 2 ) led to persistent accumulation of RPA (close to signal saturation) at all time points post-UV that we tested ( Fig 1A and 1B ). The dynamic range of this assay, within a 6-h window, is therefore much larger at low (1 J/m 2 ) vs higher doses of UV in U-2 OS cells ( Fig 1B ). As proof of principle for our experimental conditions, we treated cells with VE-821, a pharmacological ATR inhibitor that derepresses replication origins post-UV, thereby generating abundant ssDNA [ 13 ]. As expected, ATR inhibition caused a strong increase in DNA-associated RPA in response to 1 J/m 2 UV ( Fig 1C and 1D ).

Discussion

We developed a genome-wide screening strategy to identify genes limiting the formation of RPA-ssDNA in response to replication-blocking UV-induced DNA lesions. RPA-ssDNA serves as a platform for recruitment/activation of the intra-S phase checkpoint kinase ATR and other effectors of the replicative stress response [59,60]. One important role of the ATR-mediated intra-S phase checkpoint is to limit the generation of RPA-ssDNA during genotoxin-induced replication stress by prohibiting origin activation. This, in turn, preserves adequate pools of RPA, thereby forestalling genome-wide induction of DSB at persistently stalled RF [13,14]. We note that the precise molecular mechanisms underlying the formation of replication-associated DSB at stalled RF under conditions of limited RPA availability remain incompletely understood. ssDNA is known to be more susceptible to spontaneous cytosine deamination than dsDNA, leading to formation of abasic sites, which may promote further RF stalling if left unrepaired [61]. Moreover, ssDNA generated in the absence of ATR, which causes exhaustion of RPA pools, was found to be susceptible to cytosine deamination by APOBEC enzymes [62]. Finally, reducing the abundance of RPA stimulates the formation of secondary structures in ssDNA, which can lead to its nucleolytic degradation [63]. The literature therefore clearly indicates that ssDNA is intrinsically less stable than dsDNA and that its generation must be tightly controlled during replicative stress.

As expected, our ssDNA-RPA screen recovered several genes, which, by virtue of their participation in the activation of the intra-S phase checkpoint, are important determinants of RPA-ssDNA generation. Indeed, this signalling cascade is known to limit the accumulation of RPA-ssDNA during replicative stress in several ways. As mentioned earlier, the intra-S phase checkpoint signalling inhibits the initiation of new origins of replication, thereby restricting the number of stalled RF and consequent ssDNA formation [11,64]. Data from yeast also clearly demonstrate that intra-S phase checkpoint mutants accumulate much longer stretches of ssDNA than wild-type cells at individual stalled RF, although the precise mechanisms are not entirely clear [10]. Importantly, these stretches of ssDNA result at least in part from EXO1-dependent degradation, which is inhibited by the intra-S phase checkpoint kinase Rad53 in yeast [65].

A second category of “hits” from our RPA-ssDNA screen is involved in DNA damage tolerance via TLS. TLS polymerase eta is required for bypass of UV-induced CPD, and we previously demonstrated that lack of this enzyme causes strong accumulation of RPA on DNA post-UV [18]. Moreover, we and others showed that ssDNA accumulation is sufficiently elevated in TLS-deficient cells to cause S phase–specific defects in UV photoproduct removal by sequestering RPA away from damaged sites, thereby preventing its essential function in NER [18,19,66]. Interestingly, recently published data indicate that defective TLS enhances the formation of postreplicative ssDNA gaps by favoring PRIMPOL-dependent repriming beyond damaged bases [6,67]. Moreover, formation of such ssDNA gaps has been shown to enhance sensitivity to replicative stress [68,69]. Finally, data presented here clearly indicate that EXO1-mediated degradation, presumably at ssDNA gaps, promotes ssDNA-RPA accumulation in cells lacking TLS polymerase activity (S7 Fig). It therefore seems likely that formation of ssDNA gaps, as well as their extension via EXO1-dependent nucleolytic degradation, at least partially explains the strong representation of TLS polymerases, and regulators thereof, in our screen.

As expected, we also recovered genes encoding NER factors as determinants of RPA-ssDNA generation upon UV irradiation. In all phases of the cell cycle, NER-mediated removal of damaged DNA generates ssDNA gaps during the repair synthesis step, which can be extended via the action of nucleases [23]. Nevertheless, the absence of NER activity presumably results in a larger number of persistent replication-blocking UV-induced lesions, leading to ssDNA formation specifically in S phase cells, which is what we observed (Fig 2B). We note, however, that the extent of RPA-ssDNA generation caused by NER defects was less pronounced than those caused by deficiencies in the intra-S phase checkpoint or TLS pathways. This suggests that (i) a large fraction of persistent UV-induced DNA lesions can be readily bypassed by DNA damage tolerance pathways during S phase, and consequently, (ii) NER defects per se only cause modest elevation in replicative stress in human cells under our experimental conditions.

Our screen also identified several factors whose roles in modulating the cellular response to UV-induced replicative stress has not been as well documented compared with the above examples. TRiC is a chaperone complex that assists in protein folding [34,70,71] and has been reported to influence various cellular pathways including gene expression [72], cellular signalling [73], and protection against proteotoxic stress [74]. Interestingly, recent data indicate that activation of the integrated stress response, a signalling cascade that responds to protein misfolding, leads to inhibition of histone gene synthesis and consequent formation of R-loops that are known to inhibit DNA RF progression [75]. Curiously, however, published data also show that inhibition of RF progression caused by lack of histone synthesis is not associated with dramatic elevation of RPA-ssDNA [76]. Since TRiC assists in the folding of many proteins, the molecular mechanisms explaining its influence on DNA replication stress and RPA-ssDNA formation are likely complex, and their elucidation would require further experiments.

Rif1 plays several roles that might allow this factor to limit accumulation of RPA on DNA: (i) regulation of DSB repair by interacting with the critical nonhomologous end-joining factor 53BP1 [77]; (ii) inhibition of DNA replication origin activation by promoting dephosphorylation of the MCM complex [36,78]; and (iii) prevention of nascent DNA degradation at stalled RF [79]. Our results indicate that lack of 53BP1-dependent DSB repair does not cause an important accumulation of RPA-ssDNA in S phase cells. Furthermore, our data indicate that degradation of nascent DNA at stalled RF, i.e., defective RF protection, does not strongly contribute to RPA accumulation on DNA under our experimental conditions. We therefore speculate that, as is the case for cells lacking ATR [13], abnormal activation of DNA replication origins may contribute to elevated RPA-ssDNA generation in cells lacking Rif1. We note that our data are at odds with published reports indicating that cells lacking BRCA2, which are known to display strong fork protection defects, generate elevated ssDNA in response to hydroxyurea (HU) [80]. While the basis of this discrepancy is unknown, it is possible that degradation of nascent DNA in response to UV generates only a modest amount of ssDNA, which cannot be readily detected under our experimental conditions. We also note that our data are consistent with the demonstration that lack of BRCA1, which is well known to cause severe RF protection defects, does not elicit S phase–specific NER defects due to sequestration of RPA at stalled RF [17].

While our work was in preparation, 2 independent groups reported that SCAI interacts with the pol zeta subunit REV3L and plays important roles in the repair of DNA interstrand crosslinks [55,56]. One of these studies also found that SCAI is recruited to chromatin and promotes survival post-UV [56]. Our work is generally consistent with the aforementioned reports and, moreover, extends them by identifying SCAI as a regulator of DNA RF progression and ssDNA gap processing in response to UV-induced helix-destabilizing lesions. We note that Adeyemi and colleagues reported that lack of EXO1 modestly rescues the sensitivity of SCAI-null cells to the chemotherapeutic drug cisplatin [55]. While our results (S5E Fig) are seemingly inconsistent with this, it is possible that the impact of EXO1 on the viability of SCAI-null cells might be more important in response to drugs causing interstrand crosslinks such as cisplatin, relative to the situation for UV where these adducts are not induced. We also note that EXO1 extension of ssDNA gaps is known to promote Rad51-dependent homologous recombination and sister-chromatid exchange during replicative stress [52]. It is therefore possible that defects in such homology-dependent mechanisms in cells lacking EXO1 might ultimately cause DSB that require SCAI for their processing. Finally, our data do not exclude the possibility that compromised REV3L-dependent TLS might contribute to UV sensitivity in cells lacking SCAI, irrespective of EXO1-dependent nucleolytic extension. Indeed, REV3L-depleted cells are known to be very sensitive to UV [81].

As mentioned above, the REV3L subunit of TLS polymerase zeta was recently reported to physically associate with SCAI [55,56]. This was found to be independent of REV7, the other pol zeta subunit, suggesting that pol zeta per se is not involved in restricting ssDNA accumulation during replicative stress. While our screen did identify several TLS factors, including both REV3L and REV7, as negative regulators of ssDNA accumulation, epistasis experiments suggest that SCAI and REV3L play nonredundant roles in response to UV-induced replicative stress. Indeed, our in vitro data indicate that SCAI can act alone to limit EXO1 activity at ssDNA gaps, consistent with the notion that REV3L and SCAI exert distinct roles during ssDNA gap processing. Nevertheless, our results do not exclude the possibility that defective REV3L-dependent TLS might contribute to ssDNA-RPA accumulation in cells lacking SCAI. Further investigations will be required to evaluate the precise function of the SCAI-REV3L interaction in response to UV irradiation.

Consistent with the aforementioned recent study [55], we found that lack of SCAI leads to degradation of nascent DNA, i.e., RF protection defects. Moreover, depletion of EXO1 rescued UV-induced reduction in RF progression in cells lacking SCAI, suggesting that SCAI promotes DNA replication by limiting nucleolytic degradation of nascent DNA at stalled RF. We also note that depletion of BRCA2 in SCAI-null cells caused additive defects in RF protection upon HU treatment, suggesting that these proteins act in a nonredundant manner to protect RF from nucleolytic activity. Since nascent DNA degradation at stalled RF in cells lacking BRCA2 does not cause significant accumulation of RPA-ssDNA post-UV under our experimental conditions, RF protection defects are unlikely to account for the observed ssDNA accumulation in SCAI-KO cells. Interestingly, our in vitro data indicate that SCAI binds ssDNA with much greater affinity than dsDNA. This is in agreement with published data showing interaction of SCAI with ssDNA and with its colocalization with RPA in the context of DSB repair [38]. Moreover, we found that SCAI inhibits EXO1 activity on a ssDNA gap in vitro. Extension of ssDNA gaps by EXO1 and other nucleases has been shown to occur in response to lesions in template DNA [52] and to significantly contribute to the formation of ssDNA upon replicative stress [67–69]. Taken together, the above leads us to propose that interaction between SCAI and ssDNA at postreplicative gaps might prevent nucleolytic extension of these gaps by EXO1. While we did not formally investigate the impact of SCAI on EXO1-mediated degradation of nascent DNA at reversed forks, we note that this nuclease is known to act on both stalled RF and ssDNA gaps [50,52]. It is therefore tempting to speculate that SCAI-dependent reduction of ssDNA formation at gaps or reversed RF might entail a similar mechanistic basis. Nevertheless, further experiments will be necessary to fully characterize the mechanisms through which SCAI impacts the generation of ssDNA in human cells.

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