Summary

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Introduction

Replication fork reversal is a replication stress tolerance mechanism that promotes replication-coupled DNA repair or bypass of DNA damage (Berti et al., 2020a; Cortez, 2019). Multiple proteins regulate the formation and stabilization of reversed forks since unregulated fork reversal can slow replication elongation, increase the frequency of double strand breaks (DSBs), cause extensive degradation of nascent DNA, and result in genome instability.

The recombinase RAD51 is required to promote fork reversal (Zellweger et al., 2015), and protect the nascent DNA from degradation (Hashimoto et al., 2010; Schlacher et al., 2011). The RAD51 reversal function is thought to involve a metastable RAD51 filament (Berti et al., 2020a). In most cases, reversal does not require BRCA2, although it may be involved in circumstances in which RAD51 function is partly compromised (Liu et al., 2020). Instead, other RAD51 regulators including RAD51 paralogs assist fork reversal (Berti et al., 2020b), but exactly how RAD51 promotes reversal is unknown.

A reversed fork is the substrate for nascent strand degradation since inactivating fork reversal enzymes like SMARCAL1, ZRANB3, HLTF, or FBH1 blocks degradation (Higgs et al., 2015; Kolinjivadi et al., 2017; Lemacon et al., 2017; Liu et al., 2020; Mijic et al., 2017; Taglialatela et al., 2017). Fork protection is thought to require RAD51 nucleoprotein filaments that are stabilized by BRCA2. These stable filaments protect the nascent DNA from nucleases including MRE11 and DNA2 (Hashimoto et al., 2010; Schlacher et al., 2011; Thangavel et al., 2015). Many proteins in addition to BRCA2 and RAD51 prevent nascent strand degradation, and they organize into at least two pathways depending on whether the translocases SMARCAL1, ZRANB3 and HLTF or FBH1 cooperate with RAD51 to remodel the fork (Liu et al., 2020). Moreover, different levels of RAD51 function are required for fork reversal and fork protection (Bhat and Cortez, 2018; Bhat et al., 2018). Consistent with this hypothesis, reducing the concentration of RAD51 in cells or reducing its ability to bind DNA using a chemical inhibitor, B02, causes deprotection of reversed forks; whereas, potently silencing RAD51 expression prevents fork reversal and inhibits nascent strand degradation (Bhat et al., 2018; Taglialatela et al., 2017).

RADX is a single-strand DNA (ssDNA) binding protein important for replication fork stability (Dungrawala et al., 2017; Schubert et al., 2017). In the absence of added replication stress, RADX inactivation causes slow replication elongation and increased DSBs (Dungrawala et al., 2017; Schubert et al., 2017). RADX competes with RAD51 for ssDNA, directly interacts with RAD51, destabilizes the RAD51 nucleofilament, and inhibits RAD51-dependent processes in vitro (Adolph et al., 2021). Inactivating RAD51 rescues the slow replication elongation and elevated DSBs observed in RADX-deficient cells (Dungrawala et al., 2017). These data suggest that RADX antagonizes RAD51 to maintain genome stability during DNA replication.

In cells experiencing persistent replication stress caused by hydroxyurea (HU), RADX overexpression reduces the amount of RAD51 at forks and promotes fork degradation (Bhat et al., 2018). In addition, RADX inactivation restores fork protection in BRCA2, BRCA1, or FANCD2-deficient cells treated with HU (Bhat et al., 2018; Dungrawala et al., 2017). Based on the antagonistic relationship between RADX and RAD51 found in biochemical experiments and in unstressed cells, and the observations that RADX inactivation or overexpression increases or reduces RAD51 levels at forks respectively, we proposed that the rescue of fork protection by RADX inactivation could be due to restoration of sufficient RAD51 activity to protect reversed forks in these cells (Bhat and Cortez, 2018; Bhat et al., 2018; Dungrawala et al., 2017). In other words, we hypothesized that removing a negative regulator of RAD51 like RADX may compensate for inactivating positive regulators like BRCA2. However, this hypothesis does not explain why RADX accumulates at stalled forks (Dungrawala et al., 2017; Schubert et al., 2017).

In this study, we find that consistent with our previous hypothesis, RADX inhibits aberrant fork reversal and prevents fork collapse in unstressed cells. However, we now find that RADX promotes the formation of reversed fork structures in cells experiencing persistent replication stress. Our results indicate that RADX has two apparently opposite functions to regulate fork reversal depending on the amount of replication stress. To explain these observations, we propose a unifying model in which the RADX-dependent destabilization of the RAD51 nucleofilament can either inhibit or promote fork reversal depending on whether forks are actively elongating or stalled. These results are reminiscent of N. gonorrhoeae RecX which stimulates functions of the recombinase RecA (the RAD51 orthologue) in cells even though it inhibits its biochemical activities (Gruenig et al., 2010).

Results

RADX inhibits inappropriate fork reversal in the absence of exogenous stress

RADX inactivation causes slow replication elongation and increased fork collapse in the absence of added replication stress (Dungrawala et al., 2017; Schubert et al., 2017). Inactivating the fork reversal enzymes SMARCAL1, ZRANB3, or HLTF using siRNA or CRISPR-Cas9 gene editing rescues the slow fork elongation phenotype caused by RADX silencing in U2OS and hTERT-RPE-1 cells (Figures 1A and S1A–D). Additionally, inhibiting PARP with the small molecule inhibitor Olaparib, which was previously reported to prevent fork reversal by promoting fork restoration (Berti et al., 2013), also rescued the fork elongation defect in RADXΔ cells (Figure 1B). In contrast, silencing MUS81 or inhibiting MRE11 did not rescue fork speeds indicating that the fork elongation defects in the absence of RADX are unlikely to be due to fork cleavage or nuclease degradation (Figures S1E and S1F). Finally, inactivating SMARCAL1, ZRANB3, or HLTF also reduces the frequency of DSBs observed in U2OS cells lacking RADX suggesting these breaks are a consequence of aberrant fork remodeling (Figure 1C and (Dungrawala et al., 2017)).

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Figure 1.

RADX inhibits inappropriate fork reversal in the absence of replication stress.

(A) U2OS cells transfected with the indicated siRNAs (NT=non-targeting) were labeled with CldU followed by IdU and DNA combing was used to measure elongation rates. A one-way ANOVA with Tukey’s multiple comparison test was used to calculate p values in all DNA fiber experiments. (B) Replication fork elongation was monitored by DNA combing in RADXΔ cells treated with 10μM Olaparib as indicated. (C) DSBs were measured by neutral comet assay in wild-type or HLTFΔ U2OS cells transfected with the indicated siRNAs. A Kruskal-Wallis test was used to calculate p values in all comet assays. All fiber and comet assays are representative experiments of at least n=3 biological replicates. See also Figure S1. (D) Example of a reversed replication fork imaged by EM and the mean+/−SEM percentage of reversed forks from three experiments is shown (Inset, magnified four-way junction at the reversed fork; P, parental strands; D, daughter strands; R, reversed strands). The number of replication intermediates analyzed for each condition is indicated in parenthesis. A Welch’s test was used to calculate p values. See also Table S1 and Figure S2.

These data collectively suggest that RADX prevents inappropriate fork reversal at unchallenged forks that otherwise would impair replication elongation and increase the frequency of fork collapse. To further test this hypothesis, we analyzed replication intermediates using electron microscopy (EM). As predicted, silencing RADX either by siRNA transfection (siRADX) or by CRISPR-Cas9 mediated gene editing (RADXΔ) increased the frequency of reversed forks observed by EM in the absence of added replication stress (Figures 1D and S2 and Table S1). Thus, we conclude that RADX inhibits fork reversal in the absence of replication stress.

RADX inactivation blocks nascent strand degradation at persistently stalled replication forks without restoring RAD51 localization

We previously reported that inactivating RADX in U2OS cells blocks nascent strand degradation when BRCA2 or BRCA1 are also inactivated (Bhat et al., 2018; Dungrawala et al., 2017). We confirmed that this effect was not cell type specific, since RADX silencing by RNA interference also prevents nascent strand degradation in BRCA2-depleted hTERT-RPE-1 cells and BRCA2Δ DLD1 cells (Figures 2A and ​and2B).2B). We also found that despite causing elevated levels of DSBs when silenced by itself in HU-treated cells, RADX silencing reduces the fork breakage observed in BRCA2-deficient cells, which previously was reported to be dependent on fork reversal (Figures 2C and S3A) (Lemacon et al., 2017).

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Figure 2.

RADX silencing protects stalled forks from degradation in the absence of BRCA2 without restoring RAD51 localization.

(A and B) Fork protection assays were completed in (A) hTERT-RPE-1 or (B) DLD1 cells with and without BRCA2 after transfection of the indicated siRNAs. (C) Neutral comet assay in siRNA transfected U2OS cells treated with 3mM HU for 5 hours. (D) U2OS cells transfected with siRNA were labeled with 10μM EdU for 20 minutes, treated with 3mM HU for 5h, and stained for RAD51 and EdU. Representative images of RAD51 staining and the number of chromatin-bound RAD51 foci per nucleus is shown (mean +/− 95% confidence interval). Scale bar-10μm. A Mann-Whitney test was used to calculate p-values. See also Figure S3.

Since RADX inactivation increases the amount of RAD51 localized to replication forks (Dungrawala et al., 2017), we previously hypothesized that inactivating RADX at persistently stalled forks prevents nuclease-mediated nascent strand degradation by improving the stability of RAD51 filaments on the reversed forks. This hypothesis predicts that RADX inactivation should restore RAD51 localization to stalled forks in BRCA2-deficient cells. To test this prediction, we performed quantitative immunofluorescence imaging of chromatin-bound RAD51 in S-phase cells. As shown previously, silencing BRCA2 reduces the intensity and number of RAD51 foci while silencing RADX modestly increases the intensity and number of RAD51 foci in HU-treated cells (Figures 2D and S3B) (Dungrawala et al., 2017; Taglialatela et al., 2017). However, contrary to our expectation, silencing RADX in the absence of BRCA2 did not appreciably restore the number or intensity of RAD51 foci. Silencing RADX also did not increase RAD51 localization to stalled forks in BRCA1-deficient cells (Figures 2D and S3B). Thus, the ability of RADX inactivation to prevent nascent strand degradation and fork cleavage when fork protection factors are inactivated may not result from restoration of RAD51 localization to reversed forks.

To further test this idea, we reasoned that if RADX inactivation restores fork protection to reversed forks by stabilizing RAD51 filaments, then loss of RADX should not rescue nascent strand degradation in cases where fork protection is independent of RAD51 filament stability. For example, ABRO1 was reported to protect reversed forks independently of RAD51 (Xu et al., 2017). Thus, if RADX inactivation restores RAD51 filament stability on reversed forks, it should not rescue the nascent strand degradation in ABRO1-deficient cells. In contrast to this prediction, RADX silencing does restore fork protection in ABRO1-depleted cells (Figure S3C). Furthermore, we confirmed that a reversed fork is the substrate for degradation in cells lacking ABRO1 since inactivating the fork reversal proteins ZRANB3 or HLTF also prevents nascent strand degradation when ABRO1 is silenced (Figures S3D and S3E). These observations are inconsistent with the idea that RADX inactivation restores fork protection by increasing the stability of RAD51 filaments on reversed forks.

Partial depletion of RADX causes nascent strand degradation

We did not observe nascent strand degradation in either our RADXΔ cells generated by CRISPR-Cas9 or after transfection with RADX siRNA. However, Schubert et al., reported nascent strand degradation when they silenced RADX with siRNA (Schubert et al., 2017). To further explore how RADX inactivation affects fork stability, we tried to reconcile these observations. RNA interference may not fully inactivate RADX, so we considered the possibility that partial RADX loss might yield a different effect than complete loss of function as we had previously found for RAD51 (Bhat et al., 2018). To test this hypothesis directly, we utilized different concentrations of RADX siRNA to obtain varying levels of silencing. 5nM of RADX siRNA caused efficient knockdown with little RADX protein visible by immunoblotting, while 0.5nM of the same siRNA yielded only partial knockdown (Figure 3A). Strikingly, the partial RADX knockdown cells treated with HU underwent extensive nascent strand degradation even though the cells with more efficient RADX knockdown did not (Figure 3A). The same result was obtained with a second siRNA to RADX (Figure 3A). Partial knockdown of RADX did not yield fork degradation in RADXΔ cells indicating that these are not off-target effects of the siRNAs (Figure S4A). Moreover, nascent strand degradation upon partial RADX inactivation was also observed in hTERT-RPE-1 cells indicating that this effect is not cell-type specific (Figure S4B).

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Figure 3.

Partial depletion of RADX causes nascent strand degradation.

(A-D) Fork protection assays were performed in wild-type, SMARCAL1Δ, HLTFΔ or ZRANB3Δ U2OS cells transfected with the indicated amount of each siRNA. Cells were treated with (B) 100μM Mirin or (D) 25μM B02 where indicated. A one-way ANOVA with Tukey’s multiple comparison test was used to calculate all p values. All experiments were completed at least twice. See also Figure S4.

The fork degradation caused by partial depletion of RADX siRNA is prevented by treatment with mirin which inhibits MRE11 (Dupre et al., 2008), C5 which inhibits DNA2 (Liu et al., 2016), or by inactivating SMARCAL1, ZRANB3, or HLTF (Figures 3B, ​,3C,3C, and S4C). Thus, the resection substrate in these cells is likely a reversed fork that is degraded by MRE11 or DNA2.

These results phenocopy the effects previously reported for RAD51 (Bhat et al., 2018). RAD51 partial knockdown by siRNA, inhibition with a small molecule inhibitor, or mutation also yields nascent strand degradation that is dependent on fork reversal (Bhat et al., 2018; Taglialatela et al., 2017; Wang et al., 2015). Strikingly, either efficient or partial knockdown of RADX in cells treated with the RAD51 inhibitor, B02, largely restores fork protection (Figure 3D). A parsimonious explanation for these results is that in the presence of HU, RADX functions like RAD51 to help generate a reversed fork substrate for degradation even though in the absence of added replication stress, it inhibits fork reversal.

RADX is required for fork-reversal dependent telomere catastrophe in cells experiencing added replication stress

We next looked for additional contexts to test the hypothesis that RADX inhibits fork reversal in the absence of added replication stress but is needed to promote fork reversal in the presence of replication stress. One such context is at telomeres where telomerase binding to reversed forks in RTEL1-deficient cells causes telomere catastrophe (Figure 4A) (Margalef et al., 2018). If RADX is needed to promote fork reversal, then we might expect RADX inactivation to rescue the telomere catastrophe in RTEL1-deficient cells. Conversely, if RADX prevents fork reversal, then its inactivation would have no effect or increase telomeric dysfunction. As previously reported, inactivating RTEL1 in HeLa cells with long telomeres increased the frequency of telomere fragility and heterogeneity, and co-depleting the fork reversal enzyme ZRANB3 rescues these phenotypes linking them to fork reversal (Figures 4B-​-D)D) (Margalef et al., 2018). Inactivating RADX by itself did not cause a significant increase in telomere heterogeneity or fragility (Figures 4C and ​and4D).4D). When RADX and RTEL1 were co-depleted, we observed a small but statistically insignificant decrease in fragility and no change in heterogeneity compared to RTEL1 inactivation by itself (Figures 4C and ​and4D).4D). These data suggest that in the absence of added replication stress agents, loss of RADX has only minor effects on telomere stability and is not required for formation of reversed forks at telomere sequences in RTEL1-deficient cells. We next repeated the experiments in the presence of aphidicolin to increase replication stress levels and generate more persistently stalled forks. As expected, addition of aphidicolin increases telomere catastrophe in RTEL1-deficient cells (Figures S4D and S4E). In this condition, RADX inactivation by itself had no effect on telomere fragility or heterogeneity; but inactivating RADX in the aphidicolin-treated, RTEL1-deficient cells rescued the telomere fragility and heterogeneity similar to inactivating ZRANB3 (Figures 4E and ​and4F).4F). These results are consistent with the idea that in the presence of added replication stress, RADX is needed to generate the reversed forks that are a source of the telomere instability in RTEL1-deficient cells.

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Figure 4.

RADX inactivation rescues fork reversal-dependent telomere instability only in cells treated with aphidicolin.

(A) Schematic model indicating that fork reversal causes telomere catastrophe in RTEL1-deficient cells (Margalef et al., 2018). (B) Representative images of normal, fragile, and unequal telomeres. Scale bar-1μm. (C) Quantification of fragile telomeres and (D) telomere heterogeneity in cells transfected with siRNAs. (E) Quantification of fragile telomeres and (F) telomere heterogeneity in cells transfected with siRNAs and treated with 0.2μM aphidicolin for 40h. Each data point represents a metaphase spread. All experiments were completed at least three times. Parentheses indicate the number of total metaphases analyzed. P values were calculated using a Kruskal-Wallis test. See also Figure S4.

RADX promotes fork reversal in the presence of persistent replication stress

The simplest explanation of the fork protection and telomere stability data is that RADX is needed to generate reversed forks in cells experiencing persistent replication stress. To directly test this hypothesis, we utilized EM to observe the frequency of fork reversal in HU-treated cells. As reported previously, treatment with HU yields a high frequency of reversed forks (Figures 5A and ​and5B,5B, Table S1). Depletion of RADX by siRNA (siRADX) or inactivation by CRISPR-Cas9 (RADXΔ) consistently decreased the percentage of reversed forks although not to the levels of unstressed cells (Figure 5B, Table S1).

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Figure 5.

RADX inactivation suppresses fork reversal in cells experiencing persistent replication stress.

(A) Example of a reversed fork observed by EM (Inset, magnified four-way junction at the reversed fork; P, parental strands; D, daughter strands; R, reversed strands). (B) Quantitation of fork reversal measured by EM in the indicated U2OS cells treated with 3mM HU for 5h (n=3; Mean +/− SEM). The number of replication intermediates analyzed for each condition is indicated in parenthesis. P values were calculated using a Welch test. See also Table S1 and Figure S2.

RADX binds RAD51 and ssDNA to promote fork reversal in the presence of persistent replication stress

In unstressed cells, the ability of RADX to prevent fork instability is dependent on its direct interactions with RAD51 and ssDNA (Adolph et al., 2021; Dungrawala et al., 2017). To test whether the same mechanisms operate in stressed cells, we analyzed the activities of RADX separation of function mutants with diminished ssDNA binding activity (RADX OB2m, (Dungrawala et al., 2017) or an inability to interact with RAD51 (RADX QVPK, (Adolph et al., 2021)). First, we examined if transient overexpression of these proteins caused nascent strand degradation in HU-treated cells as previously observed for wild type RADX (Dungrawala et al., 2017). In contrast to wild-type RADX, the RADX QVPK or RADX OB2m mutants do not cause nascent strand degradation when overexpressed in U2OS cells (Figures 6A and S5A). The RADX overexpression-induced nascent strand degradation is rescued by overexpressing RAD51 suggesting that it is due to a reduction in the stability of RAD51 filaments on reversed replication forks (Figure S5A).

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Figure 6.

A direct interaction of RADX and RAD51 is required to maintain fork stability in cells experiencing persistent replication stress.

(A) Fork protection assays with 3mM HU were performed in U2OS cells immediately after infection with lentiviruses to overexpress wild-type (WT) RADX or RADX QVPK. Arrows in the immunoblot indicate the endogenous and GFP-tagged RADX proteins. (B) Wild-type or RADXΔ U2OS cells complemented with WT RADX or RADX QVPK were transfected with non-targeting or BRCA2 siRNA and examined for fork protection. Note that the wild-type RADX but not the RADX QVPK mutant expression in the RADXΔ complemented cells decreases after a few passages as previously reported (Adolph et al., 2021). A one-way ANOVA with Tukey’s multiple comparison test was used to calculate all p values. All experiments were completed at least three times. Extra lanes were removed.

We next examined fork protection in RADXΔ cells complemented with wild-type, QVPK, or OB2m RADX. As we previously reported, expression of the wild-type RADX protein decreases after a few passages to a level that no longer causes fork deprotection and permits nascent strand degradation when BRCA2 is silenced (Figure 6B) (Adolph et al., 2021). In contrast, cells only expressing the RADX QVPK or RADX OB2m proteins have stable forks when BRCA2 is silenced similar to RADXΔ cells consistent with the idea that RADX binding to RAD51 and ssDNA is needed to enable fork reversal in cells experiencing persistent stress (Figure 6B and S5B).

To explain the paradoxical observations for RADX function in unstressed and stressed cells, we reasoned that RADX could either “switch” from an antagonist of RAD51 in the absence of stress to a positive regulator of RAD51 with persistent stress or that it may continue to antagonize RAD51 biochemically in the presence of stress but that this function may help generate the “metastable” RAD51 filament proposed to be needed to reverse forks (Berti et al., 2020a). To test if RADX switched from an antagonist of RAD51 to a positive regulator of RAD51, we purified GFP-Flag-RADX from unstressed and HU-treated 293T cells (Fig. S6A). Both RADX proteins inhibited RAD51-mediated strand exchange and D-loop formation as efficiently as MBP-RADX purified from insect cells while the RADX-QVPK protein did not (Figures S6B and S6C, (Adolph et al., 2021)). Thus, RADX purified from untreated and HU-treated cells have similar biochemical activities. Furthermore, the ability of RAD51 overexpression to rescue the phenotypic effect of RADX overexpression in HU-treated cells (Figure S5A) also suggests it remains a RAD51 antagonist in these conditions.

To directly test whether RADX could promote fork reversal by decreasing the stability of RAD51 filaments, we examined the ability of SMARCAL1 to catalyze fork reversal on model replication fork substrates in the presence of RAD51, RPA and RADX (Figure 7A). Individually, MBP-RADX or RAD51 inhibited fork reversal when added before SMARCAL1 (Figures 7B and ​and7C)7C) but as previously reported, RPA stimulates SMARCAL1 (Figure 7B; (Betous et al., 2012). As expected, addition of RPA, RAD51 and MBP-RADX together in the absence of SMARCAL1 did not catalyze fork reversal (Figure 7D). Moreover, adding RPA by itself after RAD51 did not improve SMARCAL1-mediated reversal (Figure 7E). In contrast, adding increasing amounts of MBP-RADX in the presence of RPA to a fork substrate with pre-bound RAD51 stimulated fork reversal by SMARCAL1 (Figures 7E). Importantly, adding the MBP-RADX QVPK mutant that cannot bind RAD51 or destabilize RAD51 filaments did not stimulate reversal (Figure 7F). These data suggest that RADX can promote fork reversal in at least some conditions by destabilizing RAD51.

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Figure 7.

RADX promotes fork reversal by destabilizing RAD51 filaments in vitro

(A) Schematic of the fork reversal assay. (B). Addition of RPA stimulates SMARCAL1- dependent fork reversal, while addition of RAD51 inhibits fork reversal. (C) Increasing concentrations of RADX by itself inhibits fork reversal by SMARCAL1. (D). Addition of RPA, RADX, and RAD51 do not catalyze fork reversal in the absence of SMARCAL1. (E) Addition of increasing concentrations of RADX in the presence of RPA overcomes the fork reversal inhibition caused by RAD51. Top, quantifications (n=4; mean+/−SEM). Bottom, representative experiment. (F) Unlike wild type RADX, RADX QVPK does not stimulate fork reversal in the presence of RAD51, RPA, and SMARCAL1. Top, quantifications (n=3; mean+/−SEM). A representative experiment is shown. A repeated measures one-way ANOVA with Sidaks multiple comparisons test was used to calculate p values. The line in blots indicate lanes were removed.

Discussion

Our findings indicate that RADX regulates replication fork reversal in a context-dependent manner. In the absence of added replication stress, RADX inhibits fork reversal to prevent fork slowing and fork breakage. However, in the presence of added replication stress, RADX helps to generate reversed forks. Therefore, RADX inactivation blocks nascent strand degradation and telomere dysfunction in fork protection-deficient or RTEL1-deficient cells experiencing persistent replication stress since reversed forks are the substrates for the nuclease processing in these circumstances (Kolinjivadi et al., 2017; Lemacon et al., 2017; Margalef et al., 2018; Mijic et al., 2017; Taglialatela et al., 2017). Both RADX-dependent inhibition of fork reversal in the absence of stress and generation of reversed forks in the presence of stress depend on its ability to interact directly with RAD51 and ssDNA. Based on these findings and our observation that RADX can stimulate fork reversal in vitro in conditions where RAD51 binding a fork substrate is inhibitory, we conclude that RADX likely confines fork reversal to persistently stalled forks by destabilizing RAD51 nucleofilaments. We suggest that the difference in outcome at elongating versus stalled forks is due to the difference in amount of ssDNA.

Our new data also explains the discrepancies in the literature on the stability of nascent DNA when RADX is inactivated. We reported that silencing RADX with RNA interference or knocking out RADX with CRISPR-Cas9 caused fork slowing but did not yield nascent strand degradation in the presence of persistent replication stress (Bhat et al., 2018; Dungrawala et al., 2017). Yet, Schubert et al., reported that RADX siRNA caused nascent strand degradation in HU-treated cells (Schubert et al., 2017). We now find that inefficient RADX silencing causes nascent strand degradation indicating the differences in published results were due to how efficiently RADX was inactivated. In this respect, RADX phenocopies RAD51 since partial silencing of either causes nascent strand degradation but efficient inactivation yields stable forks in HU-treated cells.

Six observations are consistent with the conclusion that RADX prevents fork reversal in the absence of replication stress: 1) RADX is present at actively elongating replication forks placing it where it would need to operate to prevent RAD51-dependent reversal (Dungrawala et al., 2017); 2) RADX inactivation causes an increased abundance of RAD51 at replication forks (Dungrawala et al., 2017); 3) RADX inactivation slows replication elongation and increases fork collapse; 4) both fork slowing and fork collapse in the absence of RADX can be rescued by inactivating the fork reversal enzymes HLTF, ZRANB3, or SMARCAL1 or by silencing RAD51; 5) replication elongation rates in RADX-deficient cells are also rescued by addition of PARP inhibitors which inhibit fork reversal; and 6) direct visualization of replication intermediates by EM shows an accumulation of reversed forks in RADX-deficient cells that are not treated with any replication stress agent.

Six additional observations support the conclusion that RADX helps to generate reversed forks in the presence of replication stress: 1) RADX accumulates at stalled replication forks where it would need to operate to promote RAD51-dependent reversal (Dungrawala et al., 2017; Schubert et al., 2017); 2) RADX silencing or deletion prevents nascent strand degradation when fork protection factors including BRCA2, BRCA1, BOD1L, ABRO1, Fanconi anemia proteins, or DCAF14 are inactivated similar to the rescue observed by inactivating fork reversal enzymes (Bhat et al., 2018; Townsend et al., 2021); 3) the rescue of fork protection in these circumstances is not due to restoration of RAD51 localization to stalled replication forks; 4) RADX inactivation prevents fork reversal-dependent telomere catastrophe in RTEL1-deficient cells treated with aphidicolin to elevate replication stress levels; 5) RADX inactivation prevents reversal-dependent fork breakage that is observed in BRCA2-deficient cells; 6) the percentage of reversed forks observed by EM in HU-treated cells is decreased when RADX is inactivated.

We envision two models to explain the apparent difference in RADX functions in the absence or presence of replication stress (Figure S6). First, RADX could switch functions from a RAD51 antagonist to a RAD51 activator in response to replication stress. A post-translational modification or change in protein binding partners could mediate this change. Second, RADX could retain the same biochemical activity (reducing RAD51 nucleofilament stability) in both stressed and unstressed cells; however, the reduction in RAD51 nucleofilament stability induced by RADX generates the metastable filaments needed for fork reversal in stressed cells, while it prevents inappropriate RAD51 access to elongating forks in unstressed conditions. In this second model, other changes in the replication fork proteome or DNA structures when cells are treated with HU would be needed to yield the apparent difference in outcome. We hypothesize that the difference may be the amount of ssDNA at active versus stalled forks. When there is limited ssDNA for short times, such as what would be present on the lagging template strand at active replication forks or what would be generated by stochastic polymerase speed changes (Graham et al., 2017), RADX-destabilization could prevent RAD51 from accessing the forks to promote reversal. However, when there is persistent fork stalling yielding more ssDNA, then destabilization of RAD51 filaments that form on the template DNA strands could be needed to allow the fork reversal enzymes to reanneal the parental DNA and catalyze fork reversal. In other words, accumulation of RADX at stalled forks could help the reannealing of the parental DNA strands by making the RAD51 nucleofilament more dynamic so it does not act as a road-block to reversal and possibly to allow RPA to remain present where it can stimulate SMARCAL1 (Figure S7).

While both models could explain the apparent switch in RADX function, our data is more consistent with the second model for several reasons. First, RADX function in both stressed and unstressed cells requires a direct interaction with RAD51 and ssDNA. Second, loss of RADX increases RAD51 chromatin localization both in the absence and presence of stress. Third, RADX purified from unstressed or HU-treated cells inhibits RAD51-mediated strand exchange and D-loop activity. Fourth, overexpressing RAD51 overcomes the fork destabilization caused by overexpressing RADX suggesting they retain an antagonistic relationship in the presence of HU. Fifth, RAD51 added to fork reversal assays can inhibit SMARCAL1 presumably by acting as a steric block to enzyme translocation. Adding RADX to these reactions (but not RADX that cannot interact with RAD51) overcomes the RAD51-dependent inhibition when RPA is also present. The same purified RADX protein inhibits RAD51-dependent strand exchange and destabilizes RAD51 nucleofilaments (Adolph et al., 2021). Finally, a “less is more” model for RAD51 activity has precedence in the bacterial literature where the N. gonorrhoeae RecX protein biochemically inhibits N. gonorrhoeae RecA, but actually promotes its function in vivo (Gruenig et al., 2010). Although RADX has no sequence similarity to RecX, the RecA and RAD51 proteins are orthologs that likely require similar regulatory mechanisms. Taken together, we conclude that the RADX may promote fork reversal by making the RAD51 filament metastable, which was previously proposed to be required to generate reversed forks (Berti et al., 2020a).

Inactivating RADX in HU-treated cells reduces the percentage of reversed forks visualized by EM, but it does not fully prevent reversal suggesting either it is only important at a subset of stalled forks or it reduces the time that all stalled forks reside in the reversed state. Fork reversal is a dynamic process with reversed forks being “reset” by fork restoration reactions catalyzed by RECQ1 or by the limited action of nucleases like DNA2 (Berti et al., 2013; Hu et al., 2012; Thangavel et al., 2015). The fork reversal enzymes like SMARCAL1, can also catalyze fork restoration reactions (Betous et al., 2013). Moreover, other fork protection proteins like RAD52 can prevent excessive RAD51 function and engagement of translocases at replication forks to limit fork reversal upon stalling (Malacaria et al., 2019). The stability of the RAD51 filament and whether it is formed on gaps in the template DNA strands, on the nascent DNA of a reversed replication fork, or even its binding to double strand DNA at the fork will also determine the dynamics of the conversion between a three-way to four-way junction. The persistence of the replication block, amount of ssDNA, recruitment of other RAD51 regulators, and the activity of competing pathways all likely combine to yield the static snapshot of reversed fork frequency that is observed by EM. RADX may change the dynamics of RAD51 at HU-stalled forks enough to shift the equilibrium between the 3- and 4- way junctions towards reversal, but not be absolutely essential explaining why some reversed forks remain visible by EM.

Nonetheless, even though RADX inactivation only partially reduces fork reversal in cells exposed to persistent replication stress, it fully prevents nascent strand degradation in HU-treated cells in which fork protection factors are inactivated. Thus, whatever reversal happens in these circumstances is not enough to yield extensive nascent strand degradation. RADX, like RAD51, may be acting in two steps of the fork protection pathway at persistently stalled forks – promoting fork reversal and destabilizing RAD51 filaments on the reversed fork making it more susceptible to nucleases. Finally, nascent strand degradation may require multiple cycles of reversal and degradation. If RADX inactivation reduces this dynamic cycling, then that would also explain why fork degradation is completely dependent on RADX but fork reversal is only partially-dependent on RADX.

In summary, RADX antagonizes RAD51 in unstressed cells to prevent aberrant fork reversal. However, in stressed cells with persistently stalled forks, RADX interacts with RAD51 to promote reversal. In both cases, RADX may operate by destabilizing RAD51 filaments. By confining fork reversal to stalled forks, RADX prevents aberrant processing by nucleases that can generate genome instability.

STAR Methods

EXPERIMENTAL MODEL AND SUBJECT DETAILS

U2OS (female) and HEK293T (female) were cultured in DMEM with 7.5% Fetal Bovine Serum (FBS). hTERT-RPE-1 (female) cells were cultured in DMEM F12 with 7.5% FBS and 7.5% Sodium bicarbonate. HeLa1.3 (long telomere) was cultured in DMEM with 10% FBS. DLD-1 (male) was cultured in RPMI1640 with 10% FBS. U2OS RADXΔ, SMARCAL1Δ, ZRANB3Δ, HLTFΔ were generated as described previously (Dungrawala et al., 2017; Liu et al., 2020). U2OS, HEK293T, and hTERT-RPE-1 were purchased from ATCC, tested for mycoplasma, and authentication verified using short tandem repeat profiling. HeLa 1.3 (female) was a gift from Dr. Titia de Lange (Rockefeller University). DLD-1 wild type and DLD-1 BRCA2 knockouts were a gift from Dr. Douglas K. Bishop (University of Chicago). The RADXΔ cells expressing wild-type RADX or RADX QVPK were described previously (Adolph et al., 2021). Plasmid transfections were performed with polyethylenimine for 293T, FugeneHD for U2OS cells and siRNA transfections were performed according to manufacturer’s instructions with Dharmafect 1 (Dharmacon) for U2OS, 293T, and DLD-1 cells, or RNAimax (Thermofisher) for hTERT-RPE-1 and HeLa1.3 cells.

METHOD DETAILS

Supplementary Material

Acknowledgements:

Footnotes

References