RIF1 is a large protein of almost 2500 amino acids in mammalian cells (Figure 1—source data 1) with no known enzymatic activity. While information about RIF1 structural organization is limited, analyses of RIF1 homologs across species identified two motifs that are highly conserved from yeast to mammals: the N-terminal Huntingtin, Elongation factor 3, A subunit of protein phosphatase 2A, and Tor1 (HEAT) repeats, and the SILK-RVxF motif, whose sequence location shifted from the N-terminus to the C-terminal end during the evolution of unicellular to multicellular organisms (Figure 1A; Sreesankar et al., 2012; Xu et al., 2010). In vertebrates, RIF1 also exhibits a conserved C-terminal domain with a tripartite structure (Figure 1A; Xu et al., 2010). The region spanning between these N- and C-terminal motifs is poorly conserved and is characterized by a high degree of intrinsic disorder (Figure 1A). Additionally, RIF1 contains multiple serine-glutamine/threonine-glutamine (SQ/TQ) motifs, which are consensus sites for phosphorylation by the DNA damage response kinases ATM and ATR (Blackford and Jackson, 2017; Figure 1A, Figure 1—source data 2).

To identify post-translational modifications (PTMs) that modulate RIF1 functions in the maintenance of genome stability, we took advantage of the Isotopic Differentiation of Interactions as Random or Targeted (I-DIRT) experiment that we recently performed to define RIF1 interactome in mature B lymphocytes activated to differentiate ex vivo (Delgado-Benito et al., 2018). In addition to experiencing programmed DSB formation and repair during Ig CSR, activated B cells undergo a proliferative burst that renders them susceptible to DNA replication stress and damage (Figure 1B). Furthermore, activated B cells express considerably higher levels of RIF1 than their mouse embryonic fibroblast (MEF) counterparts (Figure 1C). The I-DIRT approach employed primary cultures of splenocytes from mice harboring a FLAG-2xHA-tagged version of RIF1 (RIF1^FH, Cornacchia et al., 2012; Delgado-Benito et al., 2018), which is expressed at physiological levels (Figure 1C; Delgado-Benito et al., 2018). For the RIF1 I-DIRT experiment, activated splenocytes cultures were also irradiated, which would simultaneously increase the level and broaden the range of DNA damage-induced PTMs (Figure 1D; Delgado-Benito et al., 2018). Furthermore, αFLAG-mediated pull-down of RIF1 was performed under conditions that preserved bona fide protein interactions and native complex formation (Figure 1D; Delgado-Benito et al., 2018). The RIF1 I-DIRT experiment generated a list of high-confidence interactor candidates with functions ranging from DSB repair to transcriptional regulation of gene expression (Figure 1D; Delgado-Benito et al., 2018). Moreover, a differential filtering criteria analysis uncovered an extended network of factors contributing to DNA replication initiation, elongation, and fork protection (Figure 1D). Altogether, these observations indicate that activated B cells provide an ideal model system to probe for RIF1 multiple biological functions and prompted us to re-evaluate RIF1 I-DIRT datasets for potentially relevant PTMs.

Analysis of post-translationally modified RIF1 peptides from different I-DIRT preparations revealed phosphorylation to be the predominant PTM, with the majority of phosphoresidues being serines followed by either a proline or a glutamic acid (SP or SE) (data not shown). Among all SQ/TQ motifs present in mouse RIF1, S¹³⁸⁷Q, S¹⁴¹⁶Q, and S¹⁵²⁸Q were reproducibly found to be phosphorylated across independent RIF1 I-DIRT datasets (Figure 1E and F). These SQ motifs exhibit a relatively high degree of conservation across species (Figure 1—source data 1 and Figure 1—source data 2). More interestingly, S¹³⁸⁷Q, S¹⁴¹⁶Q, and S¹⁵²⁸Q (S¹⁴⁰³Q, S¹⁴³¹Q, and S¹⁵⁴²Q in hRIF1) are located in close proximity to each other and form a defined cluster of SQ sites in the IDR of both mouse and human RIF1 (IDR-CII SQs) (Figure 1G).

We concluded that in activated B lymphocytes, RIF1 is phosphorylated at a conserved cluster of SQ motifs within its IDR.

Phosphorylation of residues within IDRs has been reported to affect protein functions in a variety of cellular contexts (Bah and Forman-Kay, 2016; Wright and Dyson, 2015). Given the conservation, proximity, and IDR location of S¹³⁸⁷Q, S¹⁴¹⁶Q, and S¹⁵²⁸Q motifs, as well as their identification as phosphoresidues in I-DIRT pull-downs, we decided to assess the contribution of the IDR-CII SQ phosphorylation to the regulation of RIF1 activities in DNA repair. RIF1 inhibits resection of DSBs downstream 53BP1 during both aberrant repair of DNA replication-associated DSBs in the absence of BRCA1 and physiological end joining of CSR breaks in G1 in B cells (Chapman et al., 2013; Di Virgilio et al., 2013; Escribano-Díaz et al., 2013; Feng et al., 2013; Zimmermann et al., 2013). Therefore, to determine if phosphorylation of the IDR-CII modulates RIF1’s role in DSB end protection, we monitored both types of repair in cells expressing phosphomutant RIF1.

To assess for aberrant (radial chromosome formation) and physiological (CSR) repair events in the same cellular context, we opted to perform our analysis in HR-deficient, yet CSR-proficient, CH12 cells bearing hypomorphic Brca1 mutations (Figure 2A). CH12 is a well-characterized mouse B cell lymphoma line that recapitulates the molecular mechanism and regulation of CSR (Nakamura et al., 1996). Furthermore, CH12 cells display a stable near-diploid genome that can be easily and efficiently manipulated by somatic gene targeting (Delgado-Benito et al., 2020; Delgado-Benito et al., 2018; Sundaravinayagam et al., 2019).

These features render CH12 the preferred model system over B cells isolated from the available BRCA1-mutated mouse models, which (1) are refractory to classic transfection methods, (2) do not allow for transduction-based reconstitution studies of large proteins like RIF1, and (3) whose primary nature precludes genetic manipulation for knock-in generation.

To generate BRCA1-mutated CH12 cells able to support CSR, we introduced in-frame deletions specifically within exon 11 of the Brca1 gene (Björkman et al., 2015; Bunting et al., 2010; Callen et al., 2013). Targeted deletion of Brca1 exon 11 in mice results in the expression of a splice variant (BRCA1- Δ11) that preserves intact N-terminal RING finger domain and C-terminal BRCT repeats but lacks key motifs that are essential for BRCA1 functions (Evers and Jonkers, 2006; Xu et al., 2001; Xu et al., 1999). BRCA1-Δ11-expressing B cells exhibit genome instability because of impaired HR but undergo CSR as proficiently as WT cells (Bunting et al., 2010; Callen et al., 2013). We employed two different nickase gRNA pairs directed towards the 5′- region of the exon (Figure 2—figure supplement 1A). All analyzed clones bore in-frame deletions, which are indicative of internally deleted, hypomorphic BRCA1 mutants (Brca1^mut, Figure 2—figure supplement 1B, and data not shown). To functionally confirm the partial loss of BRCA1 function, we assessed the levels of chromosomal aberrations in response to treatment with the PARP inhibitor olaparib (PARPi). PARPi increases the load of DNA replication-associated breaks, and in BRCA1-deficient backgrounds it triggers the accumulation of chromatid breaks and radial chromosomes (Farmer et al., 2005). These aberrations are caused by the inability to engage physiological repair by HR, in part because of suppressed DSB end resection (Bouwman et al., 2010; Bunting et al., 2010). The resulting genome instability is responsible for the increased cell lethality associated with PARPi treatment in this genetic background (Farmer et al., 2005; Rottenberg et al., 2008). Analysis of metaphase spreads revealed that in contrast to control cells PARPi-treated Brca1^mut CH12 cell lines accumulated chromatid breaks and radials with high frequency (Figure 2B, Figure 2—figure supplement 1C). Accordingly, all Brca1^mut cell lines displayed reduced viability in the presence of PARPi compared to their wild-type counterparts (Figure 2C). We concluded that Brca1^mut CH12 cells exhibit genome instability-driven cell death following PARPi treatment.

Deletion of DSB end protection factors in BRCA1-deficient cells releases the inhibition on DNA end resection and partially rescues HR, genome stability, and, as a consequence, viability (Bouwman et al., 2010; Bunting et al., 2010; Cao et al., 2009; Chapman et al., 2013; Dev et al., 2018; Escribano-Díaz et al., 2013; Feng et al., 2013; Findlay et al., 2018; Ghezraoui et al., 2018; Gupta et al., 2018; Noordermeer et al., 2018; Xu et al., 2015; Zimmermann et al., 2013). Therefore, we monitored the consequences of ablating key components of the DSB end protection cascade in Brca1^mut CH12 cell lines. In-bulk targeting of RIF1 as well as of the up- and downstream pathway components 53BP1 and REV7, respectively, led to a significant rescue of viability in Brca1^mut cells (Figure 2D). Furthermore, Brca1^mutRif1^-/- clonal derivatives exhibited reduced levels of chromosomal aberrations and a marked increase in viability after PARPi treatment compared to Brca1^mut cells (Figure 2G, Figure 2—figure supplement 2A–D). We concluded that Brca1^mut CH12 cell lines recapitulate the RIF1-dependent genome instability and cell lethality typical of BRCA1-deficient backgrounds.

CH12 cells can be induced to undergo CSR to IgA with high efficiency after activation with αCD40, IL-4, and TGFβ (CIT cocktail, Nakamura et al., 1996). NHEJ repair of CSR breaks in CH12 mimics the molecular requirements of the physiological process in primary B cells. Accordingly, Brca1^mut CH12 cell lines were able to undergo stimulation-dependent CSR to levels comparable to WT CH12 cells (Figure 2E), whereas deletion of RIF1 in these cells dramatically impaired CSR (Figure 2—figure supplement 2E).

Altogether, these findings show that Brca1^mut CH12 cell lines allow for the investigation of both outcomes of RIF1-mediated DSB end protection: aberrant repair of DNA replication-associated DSBs and physiological end joining of CSR breaks.

To investigate whether the phosphorylation status of the conserved IDR-CII SQ cluster is required for RIF1’s ability to inhibit DSB end resection, we abrogated phosphorylation of S¹³⁸⁷Q, S¹⁴¹⁶Q, and S¹⁵²⁸Q motifs by serine to alanine substitutions in Brca1^mut CH12 cells via CRISPR-Cas9-mediated knock-in mutagenesis at the Rif1 locus (Figure 2F, Figure 2—figure supplement 2F and G). This knock-in approach allows the characterization of the PTM-dependent regulation of RIF1 biological functions under physiological levels of protein expression.

Despite the expected HR deficiency of Brca1^mut cells, we obtained several clonal derivatives that harbored the desired mutations (Brca1^mutRif1^S→A) and expressed wild-type levels of RIF1 (Figure 2F). To control for any clonality-related issue, we employed three independent clonal derivatives for all subsequent analyses.

We first asked whether preventing phosphorylation of the IDR-CII SQ cluster affected RIF1’s ability to inhibit resection during repair of DNA replication-associated DSBs. To do so, we assessed chromosomal aberrations and viability following PARPi treatment. All Brca1^mutRif1^S→A cell lines accumulated chromatid breaks and radial chromosomes to the same levels as the control Brca1^mut genotype (Figure 2G) and were as sensitive to the treatment (Figure 2H). In contrast, Brca1^mutRif1^-/- cells exhibited the expected reduction in chromosomal aberrations and rescue of viability (Figure 2G and H). These results show that abrogation of phosphorylation events at the conserved cluster does not affect RIF1’s role in promoting genome instability in BRCA1-deficient cells.

We next assessed the contribution of IDR-CII SQ cluster phosphorylation on CSR, which is dependent on RIF1’s ability to protect CSR breaks against resection (Chapman et al., 2013; Di Virgilio et al., 2013; Escribano-Díaz et al., 2013). To this end, we stimulated control and Brca1^mutRif1^S→A cell lines with αCD40, TGFβ, and IL-4, and monitored CSR efficiencies. Whereas Brca1^mutRif1^-/- cells were, as expected, severely impaired in the process, Brca1^mutRif1^S→A cell lines all switched proficiently from expressing IgM to IgA (Figure 2I). This finding indicates that phosphorylation of the conserved IDR-CII SQ motifs is dispensable for physiological levels of CSR.

Finally, we assessed whether phosphorylation of the IDR-CII SQ cluster modulates RIF1’s role in the regulation of DSB resection following ionizing irradiation (IR)-induced DNA damage. To do so, we compared the phosphorylation levels of replication protein A (RPA) in RIF1-proficient, -deficient, and RIF1^S→A-expressing Brca1^mut cells. RPA is a heterotrimeric complex (RPA70, RPA32, and RPA14 subunits) that binds to single-stranded DNA (ssDNA) with high affinity (Maréchal and Zou, 2015). Defects in DSB end protection lead to hyperphosphorylation of the RPA32 subunit on S4/S8 upon IR exposure (Maréchal and Zou, 2015; Noordermeer et al., 2018). As expected, IR induced a marked phosphorylation of RPA32 in Brca1^mutRif1^-/- cells (Figure 2—figure supplement 2H). In contrast, Brca1^mutRif1^S→A cells were as proficient as controls in counteracting RPA32 phosphorylation following IR-induced DSBs (Figure 2—figure supplement 2H).

We concluded that RIF1-mediated DSB end protection activity is not dependent on the phosphorylation of the conserved IDR-CII SQ cluster.

RIF1 has recently been reported to play a genome-protective role under conditions of DNA replication stress (Ray Chaudhuri et al., 2016; Garzón et al., 2019; Mukherjee et al., 2019). RIF1 is recruited to stalled DNA replication forks where it protects nascent DNA from degradation by the DNA2 nuclease in a manner dependent on its interaction with protein phosphatase 1 (PP1) (Ray Chaudhuri et al., 2016; Garzón et al., 2019; Mukherjee et al., 2019). This activity allows for timely restart of stalled forks and prevents genome instability (Garzón et al., 2019; Mukherjee et al., 2019). Given the high proliferative nature of the cellular context where phosphorylation of the conserved IDR-CII SQ motifs was originally detected (activated primary B cells, Figure 1), we asked whether these PTMs could influence RIF1 function during replication stress.

BRCA1 plays a protective role at DNA replication forks that is independent from RIF1 (Ray Chaudhuri et al., 2016; Garzón et al., 2019; Mukherjee et al., 2019; Schlacher et al., 2012). Therefore, to specifically address the contribution of RIF1 phosphorylation to fork protection, we first generated a set of Rif1 knockout and A¹³⁸⁷A¹⁴¹⁶A¹⁵²⁸-bearing phosphomutant cell lines on a BRCA1-proficient background (WT CH12 cells) (Rif1^-/- and Rif1^S→A, Figure 3A, Figure 3—figure supplement 1A–C). As expected, deletion of RIF1 severely impaired CSR (Chapman et al., 2013; Di Virgilio et al., 2013; Escribano-Díaz et al., 2013; Figure 3—figure supplement 1D), whereas, in agreement with the findings from the BRCA1-deficient background (Figure 2I), CSR was not affected in Rif1^S→A cell lines (Figure 3—figure supplement 1D). Furthermore, analogously to what we described in the Brca1^mut setting (Figure 2—figure supplement 2H), Rif1^S→A cells did not display the IR-induced RPA phosphorylation that was detectable in the absence of RIF1 (Figure 3—figure supplement 1E).

Next, we applied the DNA fiber assay to monitor the degradation of nascent DNA at forks that were stalled via treatment with hydroxyurea (HU) (Figure 3B). HU interferes with DNA synthesis by inhibiting ribonucleotide reductase, the rate-limiting enzyme in dNTP synthesis (Singh and Xu, 2016). Both Rif1^-/- and Brca1^mut genotypes exhibited the expected fork degradation phenotype (Figure 3C), thus indicating that the protective pathways mediated by these factors are active also in CH12 cells. Interestingly, Rif1^S→A clonal derivatives showed increased degradation of stalled forks compared to controls and to the same levels observed in Rif1^-/- cells (Figure 3C), thus suggesting that abrogation of these IDR-CII SQ phosphorylation events prevents RIF1 function at the forks.

RIF1 protective role at stalled forks is dependent on the ability of its interactor PP1 to dephosphorylate and inactivate DNA2, which in turn limits the nuclease-mediated processing of DNA replication forks (Garzón et al., 2019; Mukherjee et al., 2019). To confirm the DNA2 dependency of the fork degradation phenotype observed in cells expressing phosphomutant RIF1 protein, we repeated the DNA fiber assay in the presence of the DNA2 inhibitor NSC-105808 (DNA2i) (Garzón et al., 2019; Kumar et al., 2017). Analogously to the result observed in Rif1^-/- cells (Figure 3D; Garzón et al., 2019; Mukherjee et al., 2019), DNA2i treatment rescued the fork degradation phenotype in all HU-treated Rif1^S→A clonal derivatives (Figure 3D).

We next asked whether the fork degradation phenotype exhibited by Rif1^S→A cells was mediated by the abrogation of phosphorylation at a specific SQ site within the IDR-CII SQ cluster. To answer this question, we generated single SQ mutant CH12 cell lines and assessed their capability to protect nascent DNA at stalled forks via the DNA fiber assay (Figure 3E, Figure 3—figure supplement 1F and G). We found that while Rif1^S1387A and Rif1^S1416A cell lines were proficient in protecting stalled forks from degradation, Rif1^S1528A cells exhibited a reproducible fork degradation defect (Figure 3F). However, the phenotype was modest and did not recapitulate the severe defect of Rif1^-/- and Rif1^S→A cells (Figure 3F). Altogether, this data suggests that phosphorylation of S1528 contributes to, but is not sufficient for, fork protection, and that multiple phosphorylation events within the IDR-CII SQ cluster are responsible for RIF1’s ability to protect nascent DNA under conditions of replication stress.

We concluded that phosphorylation of the conserved IDR-CII SQ cluster enables RIF1-dependent inhibition of DNA2 activity and protection of nascent DNA at stalled replication forks.

To mechanistically dissect how phosphorylation of the conserved IDR cluster contributes to RIF1’s role in protection of stalled DNA replication forks, we first assessed the integrity of RIF1-PP1 interaction via co-immunoprecipitation studies. We found that RIF1^S→A mutant protein retains the ability to interact with PP1, thus indicating that the abrogation of phosphorylation events in the conserved cluster does not have a major impact on RIF1-PP1 association (Figure 4A).

Next, we asked whether phosphorylation of IDR-CII SQ influences RIF1 recruitment to stalled DNA replication forks. To do so, we applied a proximity ligation assay (PLA) that employs flow cytometry measurements to quantitatively assess the localization of RIF1 at sites of EdU incorporation in the presence and absence of HU. As expected (Garzón et al., 2019; Mukherjee et al., 2019), RIF1 and EdU co-localization increased upon HU treatment in control cell lines (Figure 4B). In contrast, the HU-induced RIF1-EdU proximity signal was only modestly affected in Rif1^S→A clonal derivatives (Figure 4B). This data suggests that phosphorylation of the IDR-CII SQ cluster facilitates RIF1 interaction with stalled replication forks.

Finally, we investigated the dependency of phosphorylation events within the IDR-CII SQ cluster on replication stress. To this end, we have optimized RIF1 pull-downs for the identification of phosphosites in primary B cells and compared the RIF1 peptide composition of mock- versus HU-treated samples in the absence and presence of ATM or ATR inhibitors (Figure 4C). Interestingly, the only SQ site that was identified to be phosphorylated in an HU-dependent manner was indeed one of the three conserved motifs of the IDR-CII SQ cluster, S1416 (Figure 4D). In addition, HU-induced phosphorylation of S1416 was reduced following treatment with ATM and, to a lesser extent ATR, inhibitors (Figure 4D). Although this new dataset did not yield additional phosphorylated SQ motifs, we cannot exclude the likely possibility that peptides containing phospho-S1387 and phospho-S1528 residues might simply be undetectable under the conditions employed for this new set of pull-downs and mass spectrometry analysis. In support of this point, an independent proteomics analysis of hRIF1 isolated from Flp-In T-REx GFP-RIF1-L cells (Watts et al., 2020) identified S1542 (which corresponds to S1528 in mouse RIF1, see Figure 1—source data 2) as an SQ site phosphorylated following treatment with the DNA polymerase inhibitor aphidicolin, which also induces replication stress (Figure 4E). Collectively, these results build on the initial identification of S1416 and S1528 in the RIF1 I-DIRT preparations (Figure 1E and F) and demonstrate that SQ motifs in the conserved cluster are phosphorylated following treatment with replication stress-causing agents.

Altogether, these findings suggest that replication stress induces phosphorylation events within the IDR-CII SQ cluster that promote RIF1 recruitment to stalled replication forks and protection of nascent DNA. The key players and precise molecular mechanism underlying phosphorylation-dependent recruitment of RIF1 to replicated DNA will be the object of future studies.

Rif1^FH/FH (Cornacchia et al., 2012) and Rif1^F/FCd19^Cre/+ (Di Virgilio et al., 2013) mice were previously described and maintained on a C57BL/6 background. Mice were kept in a specific pathogen-free (SPF) barrier facility under standardized conditions (20 ± 2°C temperature; 55% ± 15% humidity) on a 12 hr light/12 hr dark cycle. Animals were maintained in small groups (4–5) or as breeding pairs in individually ventilated cages to ensure optimal habitat condition. Mice of both genders were used for the experiments. All experiments were performed in compliance with the European Union (EU) directive 2010/63/EU, and in agreement with Landesamt für Gesundheit und Soziales directives (LAGeSo, Berlin, Germany).

Primary cell cultures of resting B lymphocytes were isolated from WT, Rif1^FH/FH, and Rif1^F/FCd19^Cre/+ mouse spleens using anti-CD43 MicroBeads (Miltenyi Biotec), and grown in RPMI 1640 medium (Life Technologies) supplemented with 10% fetal bovine serum (FBS), 10 mM HEPES (Life Technologies), 1 mM sodium pyruvate (Life Technologies), 1× Antibiotic Antimycotic (Life Technologies), 2 mM l-glutamine (Life Technologies), and 1× 2-mercaptoethanol (Life Technologies) at 37°C and 5% CO₂ levels. Naïve B cells were activated by addition of 25 μg/ml LPS (Sigma-Aldrich), 5 ng/ml of mouse recombinant IL-4 (Sigma-Aldrich), and 0.5 μg/ml anti-CD180 (RP/14) (BD Biosciences) to the cultures upon isolation.

Primary MEFs (pMEFs) were isolated from WT and Rif1^FH/FH mice as follows. Pregnant mice were sacrificed on day E12.5 by cervical dislocation, and embryos were removed from uterine horns and placed individually in plates containing PBS (Thermo Fisher Scientific). Brain, tail, limbs, and dark red organs were removed and the remaining tissue was transferred into fresh PBS. Tissue was treated with 2 ml of Trypsin-EDTA 0.05% (Gibco) at 37°C for 15 min, and cell suspension was passed through a syringe with 18-gauge needle. Trypsin was neutralized with DMEM medium (Life Technologies) supplemented with 10% FBS, 2 mM l-glutamine, and Penicillin-Streptomycin (Life Technologies). pMEFs from each embryo were expanded in 25 cm plates at 37°C and 5% CO₂ levels to reach 80% confluency, and either used immediately for immortalization (see below) or frozen.

The cell lines employed for this study are CH12 (CH12F3, mouse, Nakamura et al., 1996); Rif1^-/- CH12 (clone 1, mouse, Delgado-Benito et al., 2018); WT (Random clones), Rif1^-/- (clone 2), and Brca1^mut CH12 clonal derivatives (mouse, this paper), as well as RIF1 phosphomutant CH12 cell lines generated on both WT and Brca1^mut backgrounds (mouse, this paper); WT and Rif1^FH/FH immortalized mouse embryonic fibroblasts (iMEFs, this paper). iMEFs were generated by immortalization of the pMEFs cultures described above via retroviral transduction of a construct expressing the SV40 T-antigen.

CH12 cells were grown in RPMI 1640 medium supplemented with 10% FBS, 10 mM HEPES, 1 mM sodium pyruvate, 1× Antibiotic Antimycotic, 2 mM l-glutamine, and 1× 2-mercaptoethanol at 37°C and 5% CO₂ levels. iMEFs were cultured in DMEM medium supplemented with 10% FBS, 2 mM l-glutamine, and Penicillin-Streptomycin at 37°C and 5% CO₂ levels.

Mycoplasma contamination was not detected in any cell line tested in the lab using commercially available mycoplasma detection kits.

RIF1 phosphoresidues were identified via analysis of RIF1 I-DIRT samples (Figure 1E and F) prepared from primary B cell cultures as previously described (Delgado-Benito et al., 2018), with the only difference that preparations with varying concentrations of glutaraldehyde (1–5 mM) were employed. Samples were loaded on NuPAGE Bis-Tris Gels (Thermo Fisher Scientific) and run for a short time to produce gel plugs. The gel samples were subjected to in-gel tryptic digestions. Peptides were extracted, purified, and analyzed by LC-MS using a Thermo Orbitrap Fusion mass spectrometer, with a Thermo Easy-nLC 1000 HPLC and a Thermo Easy-Spray electrospray source. Isotopically labeled proteins were identified by searching against a mouse protein sequence database using the GPM software (Beavis, 2006), which was set to search for tryptic peptides whose lysines and arginines were isotopically labeled and for potential phosphorylation modifications at serines, threonines, and tyrosines.

HU-induced phosphorylation of RIF1 in mouse B lymphocytes (Figure 4C and D) was performed as follows. Splenocytes isolated from WT and Rif1^FH/FH mice were treated at 72 hr post-activation with either 25 nM ATRi (BAY 1895344, Selleckchem), 20 nM ATMi (AZD0156, Selleckchem), or DMSO control for 20 min, followed by 4 mM HU (Sigma-Aldrich) or mock control for 3 hr. Cells were harvested, washed twice with ice-cold 1× PBS, and snap-frozen in liquid nitrogen. Cells were lysed at 4°C for 10 min in lysis buffer (150 mM NaCl, 50 mM Tris-HCl, 1% IGEPAL CA-630 [Sigma-Aldrich], 5% glycerol, 0.5% deoxycholate, and 0.1% sodium dodecyl sulfate) supplemented with MS-SAFE Protease and Phosphatase Inhibitor (Sigma-Aldrich), 5 mM sodium butyrate (HDACs inhibitor, Sigma-Aldrich), 5 mM 2-chloroacetamide (deubiquitinase inhibitor, Sigma-Aldrich), and Benzonase (Sigma-Aldrich). Lysates were clarified at 14,000 rpm for 10 min at 4°C and immediately used for the immunoprecipitation reaction. Protein G Dynabeads (Thermo Fisher) were conjugated with anti-HA antibody (Santa-Cruz, 2 μg/mg of whole-cell extracts) for 1 hr at room temperature (RT). Conjugated beads were washed two times with lysis buffer and incubated with lysates at 4°C for 1 hr. Beads were washed two times with wash buffer (150 mM NaCl, 50 mM Tris-HCl, 0.05% IGEPAL CA-630, 5% glycerol) and two times with wash buffer without IGEPAL CA-630, and snap-frozen in liquid nitrogen.

Tryptic on-bead digestion was carried out following essentially the protocol from Hubner et al., 2010. Proteins were digested from the beads in the presence of 2 M urea, 50 mM Tris pH 7.5, 1 mM dithiothreitol (DTT), and 5 μg/ml trypsin (Promega) at 25°C for 1 hr. Eluted bead-free pre-digested material was reduced with 4 mM DTT at 25°C for 30 min followed by an alkylation step with 10 mM iodoacetamide at 25°C for 30 min. Main digest occurred by addition of 1 μg trypsin at 25°C overnight. Samples were acidified by adding 1% (v/v) formic acid and then desalted on stage tips (Rappsilber et al., 2007). Eluted peptides were subjected to a modified SP3 procedure for an additional cleanup on peptide level (Hughes et al., 2019). Specifically, peptides were precipitated on 1 mg SP3 bead mix (Sera-Mag A and Sera-Mag B beads, GE Healthcare) by adding acetonitrile to a final concentration of ≥98% (v/v). After incubation for 20 min and three washes with pure acetonitrile, samples were eluted twice with 50 μl water. After lyophilization, samples were dissolved in MS sample buffer (3% [v/v] acetonitrile, 0.1% [v/v] formic acid). LC-MS measurements were carried out on an orbitrap Exploris 480 mass spectrometer (Thermo Fisher Scientific) coupled to an EASY-nLC 1200 system (Thermo Fisher Scientific) applying a 110 min gradient in data-dependent MS2-mode. MS1 resolution was set to 60,000 for a scan range of 300–1800, MS2 resolution was specified to 15,000 while the maximum injection time for MS2 was set to 100 ms with an isolation width of 1.3 m/z.

Analysis was done in MaxQuant (version 1.5.2.8; Cox and Mann, 2008) applying an Andromeda search against a UniProt mouse database (2018) plus common contaminants and a false discovery rate of 0.01 on peptide as well as site level while using the match-between-runs feature. RIF1 was identified with an overall sequence coverage of 65.6%. Phosphorylation on serine, threonine and tyrosine, acetylation on protein N-termini, as well as oxidized methionine were set as variable modifications. Carbamidomethylation on cysteine was set as fixed modification. The number of maximum missed cleavages was set to 5, and the number of allowed variable modifications specified to 4. Phosphosite intensities were normalized to the bait protein. A requirement of at least 60% valid values (across all samples except the WT control) was used to filter for phospho-STY sites for quantitation and further normalized by median. For two-sample moderated t-testing (limma R package; Ritchie et al., 2015) across all sites, imputation was applied by replicate using a randomized Gaussian distribution with a width of 0.2 and a downshift of 1.8. Significance calling on sites was done after multiple comparison correction by calculating adjusted p-values with the Benjamini–Hochberg method.

Analysis of aphidicolin-induced phosphorylation of hRIF1 (Figure 4E) was performed using Flp-In T-REx GFP-RIF1-L cells (HEK293-derived Flp-In T-REx 293 cells expressing GFP-hRIF1-L, Watts et al., 2020) as follows. Flp-In T-REx GFP-RIF1-L cells were cultivated in DMEM medium and induced for GFP-RIF1-L expression by addition of 1 µg/ml doxycycline (Sigma-Aldrich) 48 hr before harvesting. Cells were treated with 1 µM aphidicolin (Abcam) or DMSO (for mock control) for the final 24 hr. Cells were gently washed in dishes with 1× ice-cold Tris-buffered saline (TBS), lysed, and gently scraped off in ice-cold TBS IP buffer (1× TBS supplemented with 1% CHAPS, 1× Halt protease and phosphatase inhibitor cocktail [Thermo Fisher], and 1 mM phenylmethylsulfonyl fluoride [PMSF]) supplemented with 3 mM MgCl₂ and Benzonase. Lysates were incubated for 30 min at 4°C with gentle agitation. The lysate was spun at 20,000 for 10 min, and the supernatant then used for immunoprecipitation using GFP-Trap Magnetic-agarose beads (Chromotek). Immunoprecipitation was carried out according to the manufacturer’s instruction but in Tris IP buffer. Beads were further washed with 100 mM ammonium bicarbonate and on-beads trypsin digestion was performed essentially as described (Garzón et al., 2019) but without Cys alkylation. Peptides were analyzed using an Orbitrap Q Exactive Plus mass spectrometer equipped with nano-LC C18 liquid chromatography over 60 min elution gradient. The raw MS datasets were analyzed using MaxQuant software (version 1.6.2.3). MS1 intensity of each phosphorylation site was normalized by iBAQ value of RIF1 in each sample. The normalized phospho/RIF1 values between samples were compared.

CH12 cells were stimulated to undergo CSR to IgA by treatment with 1–5 μg/ml αCD40 (BioLegend), 5 ng/ml TGFβ (R&D Systems), and 5 ng/ml of mouse recombinant IL-4 for 48 hr. For class switching analysis, cell suspensions were stained with fluorochrome-conjugated anti-IgA (Southern Biotech) and samples were acquired on an LSRFortessa cell analyzer (BD Biosciences).

Targeting of Brca1 and Rif1 loci for generation of indel-bearing clonal derivatives was performed with two gRNA pairs per gene (gGene-N1a and gGene-N1b for Nickase 1, and gGene-N2a and gGene-N2b for Nickase 2) cloned into tandem U6 cassettes in a version of pX330 plasmid (pX330-U6-Chimeric_BB-CBh-hSpCas9, Addgene #42230) mutated to express Cas9^D10A-T2A-GFP (Nickase-1/2). The Nickase-1/2 constructs were individually transfected into CH12 via electroporation with Neon Transfection System (Thermo Fisher Scientific).

For the generation of Rif1^S→A, Rif1^S1387A, Rif1^S1416A, Rif1^S1528A, and Brca1^mut Rif1^S→A cell lines, gRNAs targeting Rif1 exon 30 (gRNA-5′ and gRNA-3′) were cloned into tandem U6 cassettes in a variant of the original pX330 plasmid (pX330-U6-Chimeric_BB-CBh-hSpCas9, Addgene #42230) modified to express Cas9^WT-T2A-GFP (pX330-Cas9^WT-T2A-GFP, kind gift from Van Trung Chu, MDC). CH12 cells (parental WT and Brca1^mut-1) were co-electroporated with the gRNA-5′/3′-expressing construct and a circular donor plasmid carrying the synthesized knock-in template (GeneArt Invitrogen). The template was purchased containing all three phosphosites mutated to alanines (A1387, A1416, and A1528) and used for the generation of Rif1^S→A and Brca1^mut Rif1^S→A cluster mutant cell lines. For the generation of Rif1^S1387A, Rif1^S1416A, and Rif1^S1528A cell lines, the donor plasmids carrying the individual SQ mutations were individually produced via two rounds of site-directed mutagenesis starting from the original synthesized knock-in template to eventually revert the other two AQ sites back to SQ motifs.

For the generation of both indel- and knock-in-bearing clonal derivatives, single GFP-positive cells were sorted in 96-well plates 40 hr after electroporation and allowed to grow for ca. 12 days before expansion of selected clones. Clonal cell lines were validated at the level of genomic scar (all clonal derivatives), protein level (RIF1 in Brca1^mutRif1^-/-, Brca1^mut Rif1^S→A, Rif1^-/-, Rif1^S→A, Rif1^S1387A, Rif1^S1416A, and Rif1^S1528A), and phenotypic consequences (BRCA1-deficiency-driven genome instability and lethality in Brca1^mut). Random control cell lines were generated with gRNAs against random sequences not present in the mouse genome (random gRNAs pairs-Cas9^D10A constructs).

For in-bulk targeting of Brca1^mut-1 cells in the rescue-of-viability assay, gRNAs against random sequences, 53bp1, Rif1, and Rev7 genes were cloned into the U6 cassette of pX330-Cas9^WT-T2A-GFP. Brca1^mut-1 cells were transfected with the Cas9-gRNAs expressing constructs via electroporation, sorted for GFP-positive cells after 40 hr, left to recover for 72 hr, and then treated with 1 μM PARPi for 72 hr before assessment of cell viability.

The sequences of the gRNAs, genotyping, and mutagenesis primers employed in this study are listed in Table 1.

Western blot analysis of protein levels was performed on whole-cell lysates prepared by lysis in RIPA buffer (Sigma-Aldrich) supplemented with 1 mM DTT (Sigma-Aldrich), cOmplete EDTA-free Protease Inhibitor Cocktail (Roche), and Pierce Phosphatase Inhibitor Mini Tablets (Thermo Fisher). For assessment of RPA phosphorylation, CH12 were seeded at a density of 10⁵ cells/ml and irradiated 24 hr later with 25 Gy, followed by 3 h of recovery time.

For RIF1-PP1 co-immunoprecipitation analysis, exponentially growing CH12 cells were treated with 4 mM HU (Sigma-Aldrich) for 3 hr. Cells were harvested, washed twice with ice-cold 1× PBS, and snap-frozen in liquid nitrogen. Cells were lysed at 4°C for 10 min in lysis buffer (150 mM NaCl, 20 mM Tris-HCl, 0.5% IGEPAL CA-630, 1.5 mM MgCl₂ [Sigma-Aldrich]) supplemented with EDTA-free Protease Inhibitor Cocktail, Pepstatin A (Sigma-Aldrich), PMSF (Sigma-Aldrich), phosphatase inhibitors (PhosSTOP, Roche/Sigma-Aldrich), and Benzonase. Lysates were clarified at 14,000 rpm for 10 min at 4°C and immediately used for the immunoprecipitation reactions. Protein A Dynabeads (Thermo Fisher) were conjugated with either anti-RIF1 antibody (4 μg/mg of whole-cell extracts for anti-RIF IP) or equal volume of pre-immune serum (Pre IP control) for 1 hr at RT. Conjugated beads were washed three times with lysis buffer and incubated with lysates at 4°C for 1 hr. Beads were washed five times with lysis buffer, and proteins were eluted by incubation at 72°C for 10 min in NuPAGE LDS sample buffer supplemented with 45 mM DTT.

The antibodies used for co-IP and WB analysis are anti-FLAG M2 (Sigma-Aldrich), FLAG-M2 peroxidase (HRP conjugated, Sigma-Aldrich), anti-HA (Santa-Cruz), pre-immune serum and anti-RIF1 (Di Virgilio et al., 2013), anti-PP1 (PPP1A/PPP1CA, Abcam), anti-phospho-RPA32 (S4/S8) (Bethyl Laboratories), anti-RPA32 (Millipore), anti-phospho-CHK1 (S345) (Cell Signaling), anti-CHK1 (Cell Signaling), anti-γH2AX (S139) (Cell Signaling), anti-H2AX (Novus Biologicals), anti-tubulin (Abcam), anti-β-actin (Sigma-Aldrich), and anti-vinculin (Sigma-Aldrich).

For assessment of cell viability, CH12 cells were either mock-treated (DMSO, Carl Roth) or incubated with 1 μM PARPi (Olaparib – AZD2281, Selleckchem) for 72 hr. Residual viability was expressed as percentage of cell viability of PARPi- over DMSO-treated cultures.

For genomic instability analysis, exponentially growing cells were treated with DMSO or 1 μM PARPi for 24 hr followed by 45 min incubation at 37°C with Colcemid (Roche). Metaphase preparation and aberration analysis were performed as follows. Cell pellets were resuspended in 0.075 M KCl at 37°C for 15 min to perform a hypotonic shock, and washed/fixed with 3:1 methanol (VWR)/glacial-acetic acid (Carl Roth) solution for 30 min at RT. Metaphase spreads were prepared by dropping fixed cells on humidified microscope slides, which were air-dried and placed at 42°C for 1 hr. Giemsa staining was performed by using KaryoMAX Giemsa Stain Solution and 1× Gurr Buffer (tablets, Gibco). Metaphases were acquired with the Automated Metaphase Finder System Metafer4 (MetaSystems).

Degradation of nascent DNA at stalled forks was assessed as follows. Exponentially growing CH12 cells were sequentially pulse-labeled with 40 μM of idoxuridine (IdU) (Sigma-Aldrich) and 400 μM of 5-chloro-2′-deoxyuridine thymidine (CldU) (Sigma-Aldrich) for exactly 20 min each, washed once with 1× PBS, and treated with 4 mM HU for 3 hr. Cells were collected and resuspended in 1× PBS at a concentration of 3.5 × 10⁵ cells/ml. 3 μl of cell suspension was diluted with 10 μl of lysis buffer (200 mM Tris-HCl pH 7.5, 50 mM EDTA, and 0.5% [w/v] SDS) on a glass slide and incubated for 2 min at RT. The slides were titled at 15–60°, air-dried, and fixed with 3:1 methanol/acetic acid for 10 min. Slides were denatured with 2.5 M HCl for 80 min, washed with 1× PBS, and blocked with 5% BSA (Carl Roth) in PBS for 40 min. The newly replicated CldU and IdU tracks were labeled for 1.5 hr with anti-BrdU antibodies recognizing CldU (1:500, Abcam) and IdU (1:50, BD Biosciences), followed by 45 min incubation with secondary antibodies anti-mouse Alexa Fluor 488 (1:500, Invitrogen) and anti-rat Alexa Fluor 546 (1:500, Invitrogen). The incubations were performed in the dark in a humidified chamber. DNA fibers were visualized using a Carl Zeiss LSM800 confocal microscope at a 40× objective magnification, and images were analyzed using ImageJ software.

Whenever indicated, the DNA2 inhibitor NSC-105808 (Kumar et al., 2017) was added at a final concentration of 0.3 μM for 24 hr prior to HU addition.

Exponentially growing CH12 cells were incubated with 10  μM of 5-ethynyl-2′-deoxyuridine (EdU) (Merck) for 15  min. For each sample, cells were washed once with 1× PBS and split into two aliquots, one of which was incubated with fresh media containing 4 mM HU for 3 hr. The other aliquot was incubated with fresh media without HU and processed for PLA in parallel (untreated condition). Cells were washed once with 1× PBS and fixed with 4% paraformaldehyde (Sigma-Aldrich) for 10 min at RT. Cells were washed twice with 1× PBS and then permeabilized for 5 min at RT using 0.2% Triton-X-100 (Roth). Cells were washed twice with 1× PBS and incubated for 30 min in the dark at RT with Click-iT Cell Reaction Buffer Kit (Thermo Fisher) supplemented with 25 μM biotin-azide (Thermo Fisher) according to the manufacturer’s instructions to conjugate incorporated EdU with biotin. Specifically, 500 μl of the Click-iT reaction was used for 6 × 10⁶ cells. After the click reaction, cells were washed with 1× PBS and then blocked with 3% BSA in 1× PBS for 1 hr at 37°C in a humidified chamber. The blocking solution was removed and cells were incubated overnight at 4°C with primary antibodies against RIF1 (1:1000) and biotin (1:1000) in blocking solution. The following day the PLA was performed using the Duolink flowPLA Detection Kit – FarRed (Sigma-Aldrich) according to the manufacturer’s instructions. Specifically, 40 μl of reaction mix per sample were used at each step, and all incubations were performed at 37°C in a humidified chamber. Cells were washed twice with Duolink wash buffer and incubated for 1 hr with the Duolink PLA probes anti-mouse plus (for biotin) and anti-rabbit minus (for RIF1) diluted 1:5 in blocking solution. Cells were then washed twice with Duolink wash buffer and incubated with Duolink ligation mix prepared by diluting ligation buffer 1:5 and ligase 1:40 in high-purity water for 30 min. Cells were washed twice and incubated for 100 min with the Duolink amplification mix prepared by diluting amplification buffer 1:5 and rolling circle polymerase 1:80 in high-purity water. Cells were washed twice and then incubated for 30 min with Duolink detection solution prepared by diluting detection buffer 1:5 in high-purity water. The detection solution was washed off, and cells were resuspended in 1× PBS containing 3% BSA. Samples were acquired on a BD LSRFortessa cell analyzer. To control for EdU incorporation, PLA values were expressed as the ratio of the mean fluorescent intensity of the HU-treated versus untreated conditions, which derived from the same EdU-incubated sample, as indicated above.

The sequence alignment of RIF1 orthologs was performed simultaneously on the full-length proteins from all 18 species listed in Figure 1—source data 1 using the multiple sequence alignment program Clustal Omega (clustalo version 1.2.4, https://www.ebi.ac.uk/Tools/msa/clustalo/).

The disorder profile plots were determined by the Protein DisOrder prediction System (PrDOS) server (https://prdos.hgc.jp/cgi-bin/top.cgi; Ishida and Kinoshita, 2007) using the template-based prediction option and with the prediction false-positive rate set to 5.0%.

Information about the statistical analysis of the mass spectrometry datasets is included in the section ‘Identification of RIF1 phosphoresidues’ above. For all other data presented in this study, the statistical significance of differences between groups/datasets was determined by the Mann–Whitney U-test. Statistical details of experiments can be found in the figure legends.