Characterization of a potent dominant negative mutant variant of Rad51 in Ustilago maydis

DNA Repair 78 (2019) 91–101

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Characterization of a potent dominant negative mutant variant of Rad51 in Ustilago maydis Jeanette H. Sutherland, William K. Holloman

T

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Department of Microbiology and Immunology, Cornell University, Weill Medical College, New York, NY 10065, USA

A R T I C LE I N FO

A B S T R A C T

Keywords: Rad51 BRCA2 Dominant negative DNA repair Homologous recombination

Rad51 serves to maintain and protect integrity of the genome through its actions in DNA repair and replication fork protection. The active form of Rad51 is a nucleoprotein filament consisting of chains of protomer units arranged linearly along single-stranded DNA. In a mutant screen using Ustilago maydis as an experimental system we identified a novel variant of Rad51, in which an amino acid change near the protomer–protomer interaction interface confers a strong trans dominant inhibitory effect on resistance to DNA damaging agents and proficiency in homologous recombination. Modeling studies of the mutated residue D161Y suggested that steric interference with surrounding residues was the likely cause of the inhibitory effect. Changes of two nearby residues, predicted from the modeling to minimize steric clashes, mitigated the inhibition of DNA repair. Direct testing of purified Rad51D161Y protein in defined biochemical reactions revealed it to be devoid of DNA-binding activity itself, but capable of interfering with Rad51WT in formation and maintenance of nucleoprotein filaments on single-stranded DNA and in DNA strand exchange. Rad51D161Y protein appears to be unable to self-associate in solution and defective in forming complexes with the U. maydis BRCA2 ortholog.

1. Introduction The tumor suppressor BRCA2, which plays a central role in cellular systems dedicated to repairing and protecting the genome from exogenous and endogenous clastogens, exerts its activity through regulating its effector Rad51, a pivotal component of the homologous recombination (HR) and replication fork protection systems [1]. Rad51 provides catalytic activity in promoting DNA strand invasion and protective activity in preventing degradation during fork stalling [2]. The active form of Rad51 in these processes is a dynamic array of protomers assembled in helical chains along DNA so as to form a nucleoprotein filament [3,4]. BRCA2 governs Rad51's activities by regulating its ability to initiate and maintain nucleoprotein filaments preferentially on single-stranded (ss) DNA [5]. The DNA strand exchange and fork protection functions rely on the stability of the Rad51 nucleoprotein filament, so cells expressing mutant variants of Rad51 compromised in filament formation, maintenance, or turnover might well be predicted to be defective in these processes. Unlike BRCA2, which exhibits considerable plasticity in size, sequence, number of Rad51-interacting BRC motifs, and composition of DNA binding modules throughout the eukaryotic domain of life [6], Rad51 is much more rigid in domain structure and functional motifs. It

is highly conserved from single-cell eukaryotes to metazoans [7]. Consider, for example, Rad51 from Homo sapiens compared to its ortholog from the yeast-like fungus Ustilago maydis (GenBank accession numbers as follows: HsRAD51CAG38796.1; UmRad51,-AAC61878.1). With the exception of some variation within the first ∼18 N-terminal residues, there is stringent conservation of sequence identity and residue alignment over the entire common length of 339 amino acids (see Fig. 1). Rad51 is an ATP-dependent DNA binding protein and a DNA-stimulated ATPase. Structural features include a globular C-terminal catalytic core domain harboring nucleotide binding Walker motifs as well as DNA-interacting loops, a smaller N-terminal domain, and an interstitial linker containing a β-strand pivotal for protomer–protomer interactions [8–10]. Assembly of protomers into the helical filament is mediated primarily through three interfaces. One is an interface harboring the nucleotide binding pocket of one protomer that is bridged by ATP with the adjacent protomer [8–10]. A second interface is formed by packing of the interstitial linker region β-strand against a central βsheet of the catalytic core domain of the adjoining protomer [11]. A third is formed by packing between an aromatic residue in the Nterminal domain (NTD) of one protomer and a cognate aromatic partner in the catalytic core of the adjacent protomer [8]. The

Abbreviations: DEB, diepoxybutane; HR, homologous recombination; MMS, methyl methanesulfonate; ss, single-strand; UV, ultraviolet light ⁎ Correspondence author at: Weill Cornell Medical College, 1300 York Avenue, New York, NY 10065, USA. E-mail address: [email protected] (W.K. Holloman). https://doi.org/10.1016/j.dnarep.2019.04.003 Received 27 December 2018; Received in revised form 8 March 2019; Accepted 9 April 2019 Available online 11 April 2019 1568-7864/ © 2019 Elsevier B.V. All rights reserved.

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Fig. 1. Screen for dominant negative Rad51 variants. (A) Schematic representation of Rad51 protein domains and functional motifs. All mutated residues that were identified in variants isolated from the screen (bold.face) or that were introduced in mutant variants utilized in this study (regular font) are shown. (B) Alignment of Rad51 orthologs from U. maydis (Um), H. sapiens (Hs), and S. cerevisiae (Sc). Divergent residues at the N-termini are not shown. Identical residues (black background); conserved residues (gray background); divergent residues (white background). (C) Representative plates showing examples of candidates identified in the mutant screen are shown. Individual transformants that were obtained by transformation of wild type cells with the mutagenized library were picked successively onto control and MMS-containing medium supplemented with hygromycin to maintain the plasmids. Controls included wild type strain UCM350 with vector (upper left hand colony) and rad51Δ strain UCM628 with vector (upper right hand colony). Potential candidates are indicated by the arrows. (D) Survival of wild type cells expressing the mutant variants recovered from the screen. Results shown here and in subsequent figures are representative of multiple determinations. At least three independent determinations were performed for each strain.

nucleoprotein filament formed on single-stranded DNA through these interactions is highly flexible and subject to large swings in pitch [8–10,12]. Association of ATP with the protomer subunits governs the pitch of the helical assembly and is therefore essential for functional activity. In the ATP bound state the filament is highly extended and competent for DNA strand transfer. After hydrolysis and conversion to ADP-Pi the filament compacts and inactivates [12,13]. Given that proper Rad51 filament assembly on DNA is key for functional activity, it is obvious that mutations disturbing the interaction interfaces could lead to loss of function. It follows that such mutant variants might also confer interference in a dominant manner by assembly together with wild type Rad51 to form hybrid filaments. This later scenario would be likely to prevail in somatic cells under the

condition of heterozygous expression. A small but growing number of Rad51 mutant variants with dominant interfering capabilities have been identified from tumor sources and from individuals with a subtype of Fanconi anemia [14]. These include variants with mutations D149N, R150Q, or G151D that comprise residues in a Schellman loop [15–18], Q268P or Q272L in DNA-binding loop 2 (L2) [19], F86L in the interstitial linker β-strand, E258A in the NTD/adjacent-protomer interface [20], and T131P [21] or A293T [22] in the nucleotide-binding pocket and cognate protomer acceptor site. While zygosity of the cellular and tissue material was unknown in several instances, there were certain examples where disease correlated with the heterozygous state of Rad51 mutant variant, e.g., [21,22]. Biochemical analysis of these mutant variants supports the conclusion that formation of faulty mixed 92

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filaments with compromised or altered ability to promote reactions was the likely cause of the pathology. Rad51 mutant variants in these studies were identified through DNA sequence analysis of tumor or patient-derived tissue. Here we were interested in identifying additional Rad51 residues with the potential for dominant negative action when mutated to learn more about mechanisms conferring dominance. For practical considerations, the strategy developed could serve as the basis for a means to evaluate expeditiously the status of human RAD51 variants of unknown functional state under the heterozygous condition. Our approach was to use U. maydis as a model system for isolation and functional analysis of Rad51 variants with a focus on residues with latent capacity for trans dominant inhibition. There are a number of advantages for using U. maydis in such a study. Genetic manipulations with U. maydis are facile, engineered genes can be readily introduced and expressed on self-replicating plasmids, and cells can be propagated in haploid or diploid state. Furthermore, since Rad51 is not essential for viability in U. maydis as it is in vertebrates [23], genetic studies involving loss of function are much easier to perform and evaluate. In addition, it could be argued that the homologous recombination system of U. maydis resembles that of human closer than other more mainstream fungal systems such as budding yeast in that U. maydis, unlike budding yeast, relies on a BRCA2 ortholog (Brh2) as a Rad51 mediator [24]. Therefore, insight into mechanisms of interference involving Brh2 could be extended to BRCA2. Finally, the Rad51 sequence of U. maydis is stringently conserved with human, so interpolating findings between organisms should be straightforward. Mutations identified in the U. maydis system would likely represent those that could serve as drivers of genomic instability and tumor progression in human. In this study we developed a screen for dominant negative Rad51 mutant variants, validated the screen by isolating a number of candidates, and characterized a novel variant with a particularly strong phenotype.

Site-Directed Mutagenesis kit (Agilent) and primers containing the desired mutation. A fragment spanning the gap promoter and rad51 allele was then transferred into a self-replicating vector expressing the hph gene (hygromycin resistance) for selection and expression in U. maydis. A vector for replacing the rad51 open reading frame with mutant alleles at the endogenous genomic locus was constructed using a 4 kbp SphI genomic fragment containing the wild type rad51 gene cloned in pUC19. A BamHI fragment with a cassette expressing the hph gene was inserted into the genomic fragment at a BclI site downstream of the rad51 gene. Cloned rad51 alleles to be tested derived from mutagenized cDNA were moved as StuI-NruI fragments from pCM1028 to replace the rad51 StuI-NruI sequence in the genomic open reading frame. As this genomic fragment contains a small intron, successful replacement by a cDNA source was signaled by loss of a PvuII restriction site marking the intron. After digestion with SphI and transfer of the modified genomic fragment into protoplasts prepared from UCM350, transformants were selected on medium containing hygromycin and the rad51 allele confirmed by sequencing after PCR amplification. A library of mutant rad51 alleles was prepared by mutagenic PCR using pCM1028 as a template and the GeneMorph II random mutagenesis kit (Agilent). The mutagenized library of rad51 gene fragments was moved into a self-replicating plasmid under the gap promoter for expression with the hph gene for selection in U. maydis. 2.3. Rad51 protein purification and analysis Rad51WT protein was purified after expression in E. coli as described previously [26,27]. Rad51D161Y protein required a different approach due to insolubility problems. The gene for Rad51D161Y was cloned into pET28b (Novagen) and expressed in E. coli T7 Express cells (New England BioLabs) harboring plasmid pKJE7 co-expressing dnaK, dnaJ, and grpE (Takara). Cell cultures (500 ml) were grown in LB medium supplemented with 50 μg/ml kanamycin, 25 μg/ml chloramphenicol and 1 mg/ml arabinose to A600 0.5, then induced by addition of 0.5 mM isopropyl-β-D-thiogalactoside, and after 4 h at 30 °C cells were harvested by cenrifugation. Cell pellets were resuspended in 30 ml of 50 mM Tris-HCl, pH 7.5, 0.25 M NaCl, 1 mM DTT, 10% glycerol (buffer A) containing 20 mM imidazole and lysed by passage through a French pressure cell. The lysate was centrifuged 1 hr at 30,000 rpm in a Sorvall T-865 rotor, and the clarified supernatant was loaded onto a column (5 ml) of Ni2+-NTA agarose (Qiagen). The column was washed successively with 40 ml buffer A containing 20 mM imidazole and 10 ml of buffer A containing 5 mM ATP and 5 mM MgCl2, and then protein eluted with buffer A plus 0.3 M imidazole. Fractions containing Rad51D161Y protein were pooled and loaded directly onto a column (4 ml) of heparin agarose (Sigma-Aldrich) equilibrated with buffer A. After washing with 40 ml buffer A, the column was eluted with a linear salt gradient (40 ml, 0.25–1.0 M NaCl) in buffer A. Peak fractions were pooled and dialyzed against solid polyethelene glycol 8000 to concentrate. The hexahistidine-affinity leader sequence tag obtained from cloning in pET28b was removed using a Thrombin Cleavage Capture kit (Novagen). Aliquots of 1 mg of protein were digested using I unit biotinylated thrombin at 20 °C for 16 h. Thrombin was then removed by passage of the mixture through a spin column (50 μl) of streptavidin agarose. Co-precipitation of Rad51 with Brh2 was performed by an affinitytag pulldown procedure using maltose binding protein (MBP)-tagged Brh2 purified in complex with Dss1 as described [26,27]. Mixes (30 μl) containing 600 nM MBP-Brh2/Dss1 in 50 mM Tris-HCl, pH 7.5, 250 mM NaCl, 1 mM dithiothreitol, 0.01% NP-40 (Buffer B), and 1.2 μM Rad51 were incubated at 30 °C. After 15 min a slurry (15 μl) of amylose resin beads (New England Biolabs) was added and the mixture left on ice for 10 min with intermittent mixing. Beads were collected by centrifugation and the supernatant saved for analysis of unbound proteins. Beads were washed in buffer (250 μl) and then bound protein eluted with 30 μl buffer B containing 10 mM maltose. Bound and unbound samples

2. Materials and methods 2.1. Ustilago maydis strains and genetic methods Manipulations with U. maydis, culture methods, synthesis of diploids, gene transfer procedures, and survival after DNA damage have been described previously [24,25]. Strains utilized were UCM350 (pan1-1 nar1-6 a1 b1) and UCM520 (met1-2 nar1-1 a2 b2) where met, pan, nar, a1 and b1 indicate auxotrophic requirements for methionine and pantothenate, inability to utilize nitrate, and mating type loci, respectively. UCM628 (rad51Δ pan1-1 nar1-6 a1 b1) was derived from UCM350 by deleting the rad51 gene with a cassette expressing resistance to nourseothricin. Other strains expressing rad51 mutant alleles were derived from UCM350. For survival, cell cultures were diluted to 2 × 107 per ml and then aliquots (10 μl) of serial ten-fold dilutions were spotted on solid medium and irradiated with 254 nm UV light or else were plated on medium containing MMS or DEB. Plates were incubated at 28 °C for 3–5 days until colonies developed. Diploids were prepared by mating UCM350 or strains expressing Rad51 mutant variants derived from UCM350 with UCM520. Heteroallelic recombination at the nar1 locus was measured by formation of Nar+ prototrophs as described previously [24]. In three independent experiments diploid cells (2 × 107) from nine cultures were spread on nitrate minimal medium to select for Nar+ recombinants. Median values were used to calculate the recombination frequency. 2.2. rad51 variant gene and mutant library construction pBluescript II plasmid derivative pCM1028 containing a cDNA copy of the U. maydis rad51 gene driven by the gap promoter was the source used for in vitro mutagenesis and library construction. Point mutations were introduced into the wild type rad51 cDNA using the QuikChange II 93

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From this hunt eighteen different variants of Rad51 with single amino acid replacements were identified [30]. Based on the frequency of identical mutations that turned up, and assuming a Poisson distribution and no bias in the selection of the suppressors, it was deduced that there could be possibly eight additional unidentified residues potentially able to give rise to a dominant negative response to DNA damage. Of these eighteen residues identified in yeast Rad51, their equivalents are all identical in human and U. maydis Rad51 orthologs, with one exception, M307, which is a conserved aliphatic residue (L249) in human and U. maydis orthologs. While in the intervening time period since that pioneering study, additional dominant negative variants of Rad51 have been identified by a variety of other means, we remained intrigued by the notion that other sites with the potential for dominant negative behavior could lie undiscovered. Therefore, we decided to search for additional novel variants, but by a different strategy. Our approach was simply to mutagenize the cloned Rad51 gene, introduce a library of the mutant alleles into a haploid wild type host strain expressing a functional Rad51 allele, and then screen individual transformants for sensitivity to DNA damaging agents. Presumably, any damage-sensitive candidates would represent clones in which both the endogenous wild type and the ectopic mutant Rad51 alleles were being expressed, but with the mutant variant interfering with the function of the wild type. Accordingly, for a pilot study we prepared a Rad51 gene library by a mutagenic PCR procedure that was designed to introduce 13 mutations per 1000 bp of DNA (Rad51 open reading frame is 1017 residues). The library was constructed in a self-replicating, low copynumber plasmid containing a hygromycin resistance gene as a selectable marker and the DNA transformed into a wild type strain. Individual hygromycin resistant transformants were picked and then tested for sensitivity to MMS (Fig. 1C). From about 500 transformants obtained using this procedure, five candidates were identified with a dominant negative phenotype when challenged with UV irradiation or exposure to diepoxybutane, in addition to MMS. Plasmid DNA was extracted from candidates and prepared for sequencing. Sequence determination of the Rad51 gene revealed there were 1-3 missense mutations present in each of the candidates.

were analyzed by SDS-polyacrylamide gel (10%) electrophoresis and stained with SimplyBlue SafeStain (Life Technologies). Density gradient velocity sedimentation was carried out in 15-30% linear gradients of glycerol with 50 mM Tris-HCl, pH 7.5, 0.2 M NaCl, 1 mM DTT, 0.5 mM EDTA using a Sorvall TH-641 rotor spun at 35,000 rpm for 24 h at 4 °C. Fractions were collected from the bottom of the tube. Aliquots of each fraction were analyzed by western blotting after SDS-gel electrophoresis and transfer to PVDF membrane. After incubation of membranes in rabbit anti-U. maydis Rad51 antiserum, Rad51 was visualized using secondary HRP-coupled goat anti-rabbit antibodies and ECL chemiluminescence reagents (GE HealthCare Life Sciences) for detection. Rad51 mutations were modeled with the PyMOL Molecular Graphics System (v1.8.4 Schrödinger) using the human RAD51 structure (NCBI Protein Data Bank PDB ID: 5H1B) as template. Rad51 multiple sequence alignment was performed using the T-Coffee alignment tool (ebi.ac.uk). 2.4. DNA dynamics DNA binding was assayed by electrophoretic mobility shift using 5′IRD800 labeled ss 60mer oligonucleotide (Integrated DNA Technologies) as described previously [28]. The 60mer sequence corresponds to residues 63-122 from bacteriophage ϕX174-(ATTATCTTG ATAAAGCAGGAATTACTACTGCTTGTTTACGAATTAAATCGAAGTGGA CTG). Briefly, reactions (15 μl) were performed with 3.3 nM DNA (as ss60mer oligomer) in 25 mM HEPES, pH 7.5, 50 mM NaCl, 2 mM ATP, and 4 mM CaCl2, or 4 mM MgCl2 where specified. Binding reactions were performed at 30 °C for 15 min, then glutaraldehyde was added to 0.2% to stabilize the protein bound complexes, and after an additional 10 min, 0.1 M Tris-HCl, pH 8.0 was added to quench crosslinking. After electrophoresis in 1% agarose gels cast in 40 mM Tris-acetate, pH 7.6, 1 mM EDTA (TAE), products were examined using the far-infrared fluorescence Odyssey platform (Li-COR Biosciences). DNA strand exchange was assayed by a gel electrophoresis assay using as substrates a ss 89mer derived from pBluescript II residues 13101 (TATAAGGGATTTTGCCGATTTCGGCCTATTGGTTAAAAAATGAG CTGATTTAACAAAAATTTAACGCGAATTTTAACAAAATATTAACG) and a homologous double-strand (ds) 35mer DNA oligomer with the strand complementary in sequence to the 89mer carrying a 5′-IRD800 label ( ATATTTTGTTAAAATTCGCGTTAAATTTTTGTTAA). This was annealed in equimolar amounts with the unlabeled complementary 35mer ( TTAACAAAAATTTAACGCGAATTTTAACAAAATAT) to yield the ds35mer. Reactions (15 μl) were performed by mixing Rad51 in solutions containing 25 mM Tris-HCl, pH 7.5, 50 mM NaCl, 1 mM dithiothreitol, 2 mM ATP, 4 mM CaCl2, 100 μg/ml bovine serum albumin and 26 nM ss 89mer at 37 °C. Mixtures were incubated 15 min, then reaction was initiated by addition of 6.6 nM ds35mer. After incubation for 15 min reactions were quenched by addition of 10 mM EDTA, 0.2% SDS, and 100 μg/ml proteinase K. Following a further incubation of 15 min to enable deproteinization, products were separated by polyacrylamide (10%) gel electrophoresis in TAE buffer and visualized as above.

3.2. Heterozygous expression of the Rad51D161Y allele sensitizes cells to DNA damage Several candidates contained mutations in or near motifs known to be in functionally important regions, such as the K133R example in the Walker A box that is required for ATP hydrolysis, F86S in the β-zip stretch that forms part of the interface scaffolding supporting protomer–protomer docking, and Q242L immediately adjacent to the DNA binding L1 loop (Fig. 1A, B & D). One candidate with multiple mutations, all in conserved resides (V82A E128G Y228P), was not explored further due to the likelihood of complications in interpretation, but instead our attention turned to the candidate with the strongest phenotype – Rad51S34LD161Y. As the S34 residue is not stringently conserved (Fig. 1B) we surmised that this change was probably of lesser consequence, and that the pivotal residue contributing to the dominant negative activity was D161, which is highly conserved across species. Since this residue is located in a region outside of known functional motifs, we judged it would be informative to investigate this Rad51 variant in more detail to understand the basis for the potent dominant negative effect on DNA repair. Therefore, we constructed the same mutation in Rad51 by site-directed mutagenesis and confirmed that the D161Y change did confer a similarly strong dominant negative effect on resistance to MMS or DEB when expressed on the same self-replicating plasmid in wild type cells. We did not pursue analysis of a variant with the S34L change. To gauge the potency of the interference imposed on DNA repair, we compared the DNA damage sensitivity of cells expressing Rad51D161Y to that of a small collection of Rad51 variants of particular interest to us in our studies of fork protection following replication fork stalling [31].

3. Results 3.1. A screen for dominant negative variants of Rad51 Searches for dominant negative mutant variants of Rad51 have been conducted previously in budding yeast. A particularly fruitful mutant hunt [29] was based on culturing homozygous srs2Δ diploids in the presence of lethal levels of methylmethane sulfonate (MMS). As the lethality, in this case, results from formation of toxic Rad51 filaments, it would be expected that mutations that disturb filament stability should suppress the toxicity. Viable papillae that emerged were suppressors that contained mutations exclusively in one of the two Rad51 alleles. 94

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Fig. 2. DNA repair and recombination proficiency supported by dominant negative Rad51 variants. (A) Survival of wild type haploid cells expressing different mutant variants from a self-replicating vector. (B) Survival of heterozygous diploids synthesized by mating wild type and mutant parental strains expressing the indicated Rad51 variant. Three independent determinations were performed for each strain in A and B. (C) Recombination at the nar1 locus was measured in heterozygous diploids expressing the indicated Rad51 mutant variant. Error bars indicate standard deviations of the median value from nine independent isolates (n = 3).

These included Rad51T131P, recently identified as a monoallelic mutation causing Fanconi anemia [21]; Rad51K133R, the well-established dominant negative variant able to bind, but not dissociate from DNA [32–34]; Rad51F232A, defective in the primary DNA-binding L1 loop [9,30,35]; and Rad51II3A, equivalent to the S. cerevisiae Rad51 DNAbinding-site-II triple mutant shown able to form nucleoprotein filaments but defective in promoting DNA strand transfer due to failure to

interact with a second DNA molecule [36]. When the alleles were expressed from a constitutive promoter on self-replicating plasmids in wild type haploid cells, or in heterozygous configuration under the natural promoter at the endogenous locus with the wild type allele in diploids, the results were similar (Fig. 2A & B). The D161Y variant along with the F232A variant were the most potent in inhibiting resistance to MMS or DEB compared to the others tested. We also 95

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sheet β2 and borders a pocket that articulates with the protruding aromatic ring on the β-zip hairpin from an adjacent protomer at the oligomerization interface. To gain understanding of the impact of mutational change of D161Y on Rad51 activity, we examined a series of Rad51 variants with a range of substitutions of D161 designed to probe whether the dysfunction results from a change in charge, size, polarity, etc. (Fig. 3A). For this end, we changed aspartate 161 to alanine, asparagine, lysine, phenylalanine, or tryptophan. There was no loss of DNA repair proficiency associated with the D161A or D161N changes as measured by ability to complement the damage sensitivity of the rad51Δ mutant (Fig. 3B). This was surprising since it might have been predicted that D161 would be part of an ion pair forming a salt bridge or a hydrogen bond donor/acceptor, but the substitution of alanine, whose side chain is small and nonpolar, without loss of function suggests a different role. On the other hand, while D161K, D161F, and D161W all caused loss of function, only D161F resulted in a comparable dominant negative inhibition (Fig. 3B). So, the inhibitory effect would appear to stem from a particular feature of the phenol or benzene side chain. This feature does not appear to be simply bulk or aromaticity since substitution of tryptophan did not cause dominant inhibition. Using the human Rad51 structure as a template and guide, we inspected the molecular architecture of residues surrounding 161 using the visualization system PyMOL and the PyMOLProbity plugin and noticed that the change from aspartate to tyrosine at residue 161 resulted in a configuration of the hydroxyphenyl ring subject to numerous van der Waals steric clashes (note position of side chains of D161 and Y161 with respect to the transverse loop in Fig. 3A). Those same clashes were not evident with tryptophan substitution, because the indole ring of tryptophan was oriented at a right angle to that of tyrosine. Curious as to whether diminishing these contacts with the hydroxyphenyl ring by altering other residues in the vicinity might mitigate the dominant effect of the D161Y alteration, we manually changed several surrounding residues and empirically evaluated steric clashes using PyMOL. It appeared that by altering both E163 and T165 to Y residues the molecular environment around Y161 would be opened so as to lessen the steric clashes imposed (Fig. 4A). We tested this prediction in vivo by making double- and triple-mutant changes in combination with D161Y. As with the Rad51D161Y variant, expression of Rad51T165Y from a plasmid in wild type conferred a strong dominant negative phenotype, although expression of Rad51E163Y did not. The double mutant Rad51D161YT165Y was slightly reduced in its dominant inhibitory activity, but the triple mutant Rad51D161YE163YT165Y was substantially suppressed in accord with the prediction from our modeling studies (Fig. 4B). These investigations suggest that the strong dominant inhibitory DNA repair phenotype resulting from the D161Y mutation arises from structural changes impacting the Rad51 oligomerization interface.

Fig. 3. DNA repair proficiency of Rad51 variants with amino acid changes at residue 161. (A) Residue 161 with different amino acid substitutions. PyMOL visualization of the region around the tip of beta sheet β4 and contiguous loop. The wild type aspartate residue is at the top. (B) Survival of wild type and rad51Δ mutant cells expressing the Rad51 variants from a self-replicating plasmid. Three independent determinations were performed for each strain.

3.4. The Rad51D161Y variant impairs formation and destabilizes the Rad51 nucleoprotein filament The genetic and structural studies suggested that the molecular mechanism by which the Rad51D161Y variant operates to confer the dominant effect is through disturbance of Rad51WT nucleofilament assembly. To test this hypothesis, we investigated metrics of the nucleofilament using purified proteins (Fig. 5A) and DNA oligonucleotides in defined in vitro biochemical reactions. Since our working model was that the Rad51D161Y variant interfered with formation of Rad51WT filaments, likely by forming co-complexes that stunted polymerization, we reasoned that a straightforward test would be to show that the Rad51D161Y variant could inhibit formation of Rad51WT filaments on single-stranded (ss) DNA. Rad51 filament formation on DNA has been intensively studied by numerous means ranging from simple bulk phase procedures to highly sophisticated single molecule methods using microfluidic devices [12,13,37–39]. It has been concluded that Rad51 associates with DNA,

investigated the influence of the Rad51 variants in inhibiting mitotic recombination in diploids heterozygous for the Rad51 gene. Here recombination at the nar1 locus was measured by gene conversion to yield prototrophs able to utilize nitrate as a source of nitrogen. Spontaneous recombination was reduced substantially by expression of the Rad51D161Y allele, to about the same level as seen with F232A, more so compared to the other alleles (Fig. 2C). Together these findings support the conclusion that Rad51D161Y expressed in a heterozygous configuration with the wild type Rad51 protein (Rad51WT) exerts strong inhibition on DNA repair and recombination processes. 3.3. Structural context of the D161Y mutation Residue D161 is located near the surface of Rad51 at the end of beta 96

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Fig. 4. Suppression of the dominant negative phenotype of Rad51D161Y by altering the atomic environment. (A) Residue D161 altered to Y (left panel) and with surrounding residues changed to Y (right panel) as visualized using PyMOL. (B) Survival of wild type cells expressing Rad51 variants from a self-replicating plasmid. Three independent determinations were performed for each strain.

Fig. 5. Biochemical characterization of Rad51D161Y. (A) SDS polyacrylamide (10%) gel of Rad51 proteins used in the analysis after staining with SimplyBlue. (B) DNA binding activity of Rad51WT and Rad51D161Y. Reactions with 2 mM ATP and 4 mM CaCl2 contained ss60mer DNA oligonucleotide and increasing levels of Rad51WT or Rad51D161Y (left or right panels) as follows: lane 1, no protein; lane 2, 47 nM; lane 3, 94 nM; lane 4, 188 nM; lane 5, 375 nM; lane 6, 750 nM; lane 7, 1.5 μM; lane 8, 2 μM. Rad51D161Y interferes with Rad51WT DNA binding. DNA binding reactions were set up as in B with ss60mer DNA oligonucleotide and a constant level of Rad51WT at 750 nM but with varying levels of Rad51D161Y (lanes 4–7). Rad51WT, Rad51D161Y, and mixtures were preincubated in buffer at 20 °C for 16 h before addition of DNA (lanes 2–7). Molar ratios of Rad51D161Y to Rad51WT as follows: 0.25:1 (lane 4); 0.5:1 (lane 5); 0.75:1 (lane 6); 1:1 (lane 7). (D) Rad51D161Y destabilizes Rad51WT filaments. Filaments were formed in a reaction mix containing ss60mer DNA oligonucleotide, Rad51WT, 2 mM ATP and 4 mM MgCl2. After incubation for 30 min at 30 °C an aliquot was removed to assess filament formation (lane 1), then the reaction mix was split into two fractions and Rad51D161Y was added to one fraction at a molar ratio of 1:1 with Rad51WT. Aliquots were removed after 2, 4, and 6 h of additional incubation and complex formation was assessed (lanes 2–7). (E) Rad51D161Y interferes with Rad51WT DNA strand exchange. DNA strand exchange reactions (substrate and product in schematic on the left) were set up with ss89mer and IRD800 ds35mer oligonucleotide substrates and Rad51as described in Section 2. Mixtures of Rad51D161Y and Rad51WT were preincubated together at 20 °C for 16 h before adding to ss89mer (lanes 4–7). No protein control, 0 min incubation (lane 1); No protein control, 15 min incubation (lane 2); Rad51WT at 1 μM (lanes 3–7); Rad51D161Y at 0.25 μM (lanes 4, 5, 8, 9); 0.5 μM (lanes 6, 10); 1 μM (lanes 7, 11). Rad51D161Y at 1 μM, heat-inactivated at 100 °C, 5 min (lane 8). Vertical bars indicate splice points from different gels in the composite image. The figures shown in C, D and E are representative of three independent determinations. 97

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for the reactions under study. In a third approach, we examined Rad51D161Y in DNA strand exchange by a gel electrophoresis assay using as substrates an ss89mer and a homologous ds35mer labeled on the strand complementary to the ss89mer. Strand exchange promoted by Rad51WT was signaled by a shift in mobility of the label from the faster moving ds35mer homoduplex to the slower moving 35mer/89mer heteroduplex (Fig. 5E). As expected, based on the inability of Rad51D161Y to form complexes with ssDNA, the mutant variant was unable to promote strand exchange. Mixing Rad51wt with increasing levels of Rad51D161Y resulted in inhibition of strand exchange. The level of inhibition appeared to correlate generally with the degree of inhibition observed in filament formation (Fig. 5C), but it should be noted that the assays differed in the concentrations and lengths of the single-stranded oligonucleotides and in the levels of Rad51WT employed. Overall, these findings are in accord with a model that Rad51D161Y variant operates to confer the dominant effect by interfering with assembly and stability of Rad51WT nucleofilaments.

not in monomer units, but as multimeric assemblies. Estimates range from assemblies comprised of 2-3 or 5-6 monomers depending on the laboratory and methodology employed [13,38,40]. To assess Rad51 filament integrity we used a basic agarose gel electrophoresis mobility-shift assay to measure complexes formed between purified Rad51 and a fluorescently tagged ss60mer as a substrate (Fig. 5B). Since Rad51 binds weakly to ssDNA [41], we added glutaraldehyde following the binding reaction mixtures to crosslink and stabilize protein-DNA complexes enabling visualization. In previous studies using this assay we noted [42], as have others previously, that complex formation did not increase linearly with increasing Rad51, but rather required addition of a certain threshold level of Rad51 before complex formation became apparent, consistent with a multimeric and cooperative mode of assembly. With a single-stranded oligonucleotide 60 residues in length (ss60mer), it could be calculated that at complete saturation Rad51 nucleoprotein filaments would be composed of chains of about 20 monomer units, which would constitute filaments of about 3 helical turns (6.4 Rad51 monomers per turn [10]). It would also be expected that with lower levels of Rad51, ss60mers bound with shorter chains could also form complexes detectable by the gel shift assay. It is not known what would be the minimum number of Rad51 monomers associated to enable an electrophoretic gel shift, but presumably this would be the basic assembly unit. If the mechanism of inhibition imposed by Rad51D161Y variant is to interfere with Rad51WT oligomerization, then it would seem reasonable to expect that interference with formation of competent Rad51WT multimer units would lead to decreased DNA binding as measured by the gel shift assay. Thus, an assumption for our analysis is that Rad51 co-complexes with fewer than the minimum number of contiguous Rad51WT monomer subunits in an assembly unit would likely be deficient in binding DNA. Rad51D161Y variant by itself was defective in binding to the ss60mer as determined by the gel shift assay (Fig. 5B), in line with the loss of biological function as measured by failure to complement the DNA damage sensitivity of the rad51Δ mutant. Little DNA binding activity was evident as apparent by the absence of well-defined complexes seen with Rad51WT. When the variant was premixed with Rad51WT binding to DNA was inhibited (Fig. 5C). At a Rad51D161Y/Rad51WT molar ratio of 0.25/1 and with enough Rad51WT to completely saturate the ss60mer there was ∼50% inhibition of binding, and nearly complete inhibition at a ratio of 1/1. This suggests that Rad51D161Y interferes with formation of competent Rad51WT multimer assemblies and inhibits DNA binding. We note that to observe inhibition, it was necessary to premix Rad51D161Y with Rad51WT for several hours. We observed little inhibition of Rad51WT function if Rad51D161Y and Rad51WT were added separately, but simultaneously, to DNA binding reactions. Rad51 exists in heterogeneous polymeric form in solution, so mixed polymer formation likely requires a suitable time period for dissociation and reassortment of the different subunits to achieve a new distribution of protomers into polymeric form. We have not performed a systematic study detailing the impact of Rad51D161Y on Rad51WT DNA binding potential over time, but routinely premixed proteins for 16 h at 22 °C to allow for redistribution of polymer subunits. It should be noted that in two related, earlier studies on other dominant negative variants of human RAD51, no premixing was performed, yet loss of RAD51WT activity in DNA binding or strand exchange was evident [21,22]. We also tested whether Rad51D161Y could destabilize preformed nucleoprotein filaments of Rad51WT and ss60mer oligonucleotide (Fig. 5D). When Rad51D161Y was added to preformed nucleoprotein filaments with Rad51WT at a ratio of Rad51D161Y/Rad51WT of 1/1, there was dissociation as observed by release of free ss60mer over time. It should be noted that Ca2+ was used as cofactor in the study on formation of Rad51-ssDNA complexes as it does not support ATP hydrolysis and locks the filament in the high affinity state. Mg2+ was used as cofactor in the destabilization study since it supports ATP hydrolysis and enables Rad51 release from complexes [13]. ATP and divalent ion concentrations were titrated to determine empirically the optimal level

3.5. Rad51D161Y intermolecular protein interactions The interface formed by docking the interdomain linker-region βhairpin with the hydrophobic pocket of its cognate partner-molecule during conjunction of Rad51 protomers can be thought of as a ball-andsocket joint. Since our structural analysis suggests that the Rad51D161Y variant is defective in the acceptor face or socket, then it could be predicted that Rad51D161Y would not self-associate in solution. It could also be predicted that if Rad51D161Y variant is mixed with Rad51WT, then the latter's capacity for self-polymerization would be diminished due to formation of co-complexes with Rad51D161Y, which would contribute a defective interaction surface thereby capping further polymerization. We tested these predictions by determining the size of the different Rad51 variants by velocity sedimentation in glycerol gradients (Fig. 6A). Rad51WT was found to sediment in a heterodisperse manner with the bulk of the protein near the bottom of the gradient, consistent with formation of polymeric complexes. Rad51D161Y sedimented near the top of the gradient in a peak almost coincident with that of an ovalbumin standard (Mw = 43 kDa), supporting the notion that Rad51D161Y is present in solution as a monomer (Mw = 37 kDa). When a mixture of Rad51WT and Rad51D161Y was examined after a preincubation period of 16 h, there was an apparent shift in distribution of the Rad51D161Y peak with concomitant accumulation of material at an intermediate position in the gradient. This is indicative of formation of mixed polymers smaller in mass than those formed with Rad51WT, but larger than monomeric Rad51D161Y. These findings support the notion that Rad51D161Y is defective in the oligomerization surface and can limit Rad51WT polymerization. Further studies using more sophisticated methodology, e.g., dynamic light scattering, might be able to provide better resolution and more insight into the kinetics of the redistribution of wild type and mutant Rad51 protomers. We also examined the interplay between Rad51D161Yand Brh2, which contains a single BRC element mimicking the Rad51 interdomain β-hairpin [24]. To determine interaction, we used a pulldown procedure to co-precipitate MBP-tagged Brh2 and therefore capture any associated Rad51 (Fig. 6B). As was apparent, Brh2 was active in forming complexes with Rad51WT, but not Rad51D161Y. These findings are consistent with the notion that Rad51D161Y variant has a defective βhairpin acceptor interface. 4. Discussion Here we identified a novel variant of Rad51 in which a specific single amino acid change at a residue situated near the protomer–protomer interaction interface confers a strong trans-acting dominant negative phenotype in resistance to DNA damaging agents 98

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Fig. 6. Rad51D161Y protein interactions. (A) Glycerol gradient centrifugation. Rad51WT (2 μM) and Rad51D161Y (1 μM) protein samples in 0.3 ml buffer A, separately or in a mixture pre-incubated at 20 °C for 16 h, were layered on top of polyallomer centrifuge tubes containing glycerol gradients (11 ml) and spun as described in Section 2. Fractions (∼0.5 ml) were collected from the bottom of each tube and aliquots (40 μl) removed for western blot analysis. Ovalbumin (500 μg) used as a size marker in a parallel gradient. It was detected by staining the gel with Simply Blue. (B) Pulldown analysis of Rad51D161Y. MBP-tagged Brh2 in complex with Dss1 was mixed with a 2-fold molar excess of either Rad51WT or Rad51D161Y variant as described in Section 2. After incubation, amylose resin was added to capture Brh2 and associated Rad51, and then the composition of the amylose-bound (bnd) and unbound (unbnd) fractions was determined by SDS-gel electrophoresis. Lane 1, Brh2 control; lane 2, Rad51WT control; lanes 3, 5, Brh2 + Rad51WT; lanes 4, 6, Rad51WT; lanes 7, 9, Brh2 + Rad51D161Y; lanes 8, 10, Rad51D161Y.

Since the active form of Rad51 in DNA repair is a polymer of end-toend assemblies of monomer units along ssDNA, it is easy to envision how mixing in a mutant variant that disturbs the assembly could impose an adverse effect. Those residues involved in docking at the protomer–protomer interface, those in the nucleotide binding pocket of one protomer making contact with the ATPase domain of the next protomer, those in allosteric regulation, in sensing nucleotide binding and in coupling hydrolysis with overall filament conformation and DNA binding might all have the potential for rendering the protein into a dominant negative variant when mutated. Numerous instances of dominant negative Rad51 variants have been reported and examples representative of these different mechanistic classes of inhibitory proteins have been documented. The Rad51D161Y variant reported here that appears to be deranged in protomer–protomer docking is particularly potent in interfering with activity of the Rad51WT as evidenced by the extreme sensitivity to MMS or DEB conferred on cells expressing both alleles. It is instructive to compare the properties of Rad51D161Y with other dominant negative variants. Rad51D161Y is as inhibitory as the variant Rad51F232A, which has an alteration of the important aromatic residue within the L1 DNA binding loop and strongly diminishes DNA binding ability [9,35]. Both of these variants are more inhibitory than Rad51K133R, which is able to form a nucleoprotein filament that is incapable of turning over due to absence of ATP hydrolytic activity [32], and far more inhibitory than Rad51T131P, found causative of disease in a Fanconi anemia individual [21], and which is abnormal in ATP hydrolysis and DNA binding and able to diminish DNA strand exchange of Rad51WT. Besides the more obvious conclusion that Rad51 variants with mutations in different functional domains can confer a spectrum of defects to different

and in homologous recombination. Simple molecular modeling studies suggested that steric interference with surrounding residues resulting from the change of aspartate 161 to tyrosine was the mainspring of the inhibitory effect and that secondary changes of two nearby residues predicted to minimize steric clashes could mitigate the inhibition. Restoration of DNA repair proficiency in cells expressing the Rad51 mutant variant with the secondary changes supported the prediction. Direct testing of purified Rad51D161Y protein in defined biochemical reactions revealed it to be devoid of DNA-binding and strand exchange activity and capable of interfering with Rad51WT in formation and maintenance of nucleoprotein filaments on single-stranded DNA. The purified Rad51D161Y protein did not form polymers and was defective in forming complexes with Brh2. The recent recognition from whole genome sequencing studies that HR deficient cancers constitute a distinct entity in the spectrum of human cancers has awakened new interest in the components of the HR system beyond BRCA1 and BRCA2 that could contribute [43]. HR-deficiency includes up to 10% of all cancers, 25% of breast cancers and 50% of ovarian cancers suggesting that it is a large population of cancers [44]. While it would be expected that biallelic mutation in an HR gene would be the primary mode triggering loss of HR proficiency and setting the stage for tumorigenic transformation, the recent reports documenting monoallelic mutation of Rad51 in certain tumor sources [17] and in individuals with a subtype of Fanconi anemia [21,22,45], which overlaps mechanistically with HR in maintaining genomic integrity, supports the notion that generation of dominant negative variants of HR components provides an additional route towards tumorigenesis. These developments piqued our curiosity in identifying additional dominant negative variants of Rad51. 99

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Conflicts of interest

degrees, the observations also demonstrate that similarly severe dominant negative impacts on biological activity from Rad51 variants can arise from mutations in different functional domains. While the Rad51D161Y variant is a potent inhibitor of DNA repair in the context of reducing survival of diploid cells treated with clastogens by several orders of magnitude, it is remarkable to note the inhibition of homologous recombination activity in unstressed cells amounts only to a four- to five-fold reduction. This apparent discrepancy in the magnitude of responses seems paradoxical. However, there are two aspects of this conundrum that might rationalize the findings. The first is that clastogen treatment probably causes widespread damage to the genome resulting in massive replication-associated DNA breakage, which would require large scale mobilization and usage of Rad51. If a toxic mutant variant were present in the Rad51 pool when the demand is high, the variant would probably also be incorporated, but with deadly consequences. On the other hand, in unstressed cells the demand for Rad51 at sites of replication fork collapse would likely be much lower, and also relatively more Rad51WT could be available for repair, if there were a mechanism for selectively utilizing it. As we showed here, Brh2 fails to interact with Rad51D161Y, and so it could act as a filter to selectively mediate loading of only the Rad51WT variant at sites requiring repair. This might account for the seemingly high residual level of spontaneous allelic recombination in heterozygotes expressing both the wild type and mutant alleles. By the same token, it could be that when the demand for Rad51 is elevated as a result of widespread damage, there is less discrimination in utilization of Rad51, and as a result defective filaments are formed with dire consequences on survival. It is interesting to consider our findings in the context of Rad51 mutant variants associated with human cancers that have been annotated in the COSMIC (Catalogue of Somatic Mutations in Cancer [https://cancer.sanger.ac.uk/cosmic]) database. The number of mutated Rad51 samples is small (108 listed in COSMIC v89). Nevertheless, from among the several dominant negative variants that came from our screen, an example is evident in COSMIC, i.e., a variant with mutation at F86 (F86L, breast carcinoma, TCGA-BH-A1FN-01). This phenylalanine is well established from modeling and structural studies as an essential residue of the interstitial linker region β-strand that is key for establishing protomer–protomer interaction by docking into a pocket in the adjoining protomer [46]. And it is well known that a phenylalanine residue within the BRC motif of BRCA2 mimics this interaction interface enabling interplay with RAD51 [11,46,47]. Based upon the potential for disruption of Rad51 polymer assembly, it would certainly be predicted that mutation of F86 could be pathogenic. While an example of a tumor with the D161Y mutation is not yet in the database, our analysis would suggest that this change would be a candidate for triggering tumorigenic transformation, especially given that as demonstrated here the Rad51D161Y exhibits a more severe phenotype in DNA repair than Rad51F86S. A final aspect of the work here that deserves mention is question of whether there could be some utility to Rad51D161Y or other dominant negative variants. It is perhaps unorthodox, but the argument could be made that while Rad51D161Y has lost function, it could serve as a specific and potent large molecule inhibitor of HR by precisely targeting Rad51. Similarly, it could be imagined that by expressing less potent dominant negative variants, HR proficiency could be attenuated or “dialed down.” A collection of Rad51 variants calibrated for potency as measured by effects on survival of wild type cells such as described in this study could be utilized as a set of specific reagents.

The authors confirm that there is no conflict of interest, financial or otherwise in this work. Acknowledgments The work was supported in part by grant 18-A8-104 from the STARR Cancer Consortium. We are grateful to the following individuals: Maria Jasin (Memorial Sloan Kettering Cancer Center), Agata Smogorzewska (Rockefeller University) for encouragement and support during inception of this work; Steve Kowalczykowski (UC Davis), Claire Wyman (Erasmus University) for Rad51 advice; George Khelashvili, Neal Lue (Weill Cornell Medical College) for help with molecular modeling and strain construction; Lorraine Symington (Columbia University) for discussion and comments on the manuscript; Qingwen Zhou (this laboratory) for sharing reagents and protocols. References [1] C.C. Chen, W. Feng, P.X. Lim, E.M. Kass, M. Jasin, Homology-directed repair and the role of BRCA1, BRCA2, and related proteins in genome integrity and cancer, Annu. Rev. Cancer Biol. 2 (2018) 313–336. [2] K.P. Bhat, D. Cortez, RPA and RAD51: fork reversal, fork protection, and genome stability, Nat. Struct. Mol. Biol. 25 (2018) 446–453. [3] J.E. Haber, DNA Repair: the search for homology, Bioessays 40 (2018) e1700229. [4] S.C. Kowalczykowski, An overview of the molecular mechanisms of recombinational DNA repair, Cold Spring Harb. Perspect. Biol. 7 (2015). [5] M.K. Shivji, S.R. Mukund, E. Rajendra, S. Chen, J.M. Short, J. Savill, D. Klenerman, A.R. Venkitaraman, The BRC repeats of human BRCA2 differentially regulate RAD51 binding on single- versus double-stranded DNA to stimulate strand exchange, Proc. Natl. Acad. Sci. U.S.A. 106 (2009) 13254–13259. [6] H. Saeki, N. Siaud, N. Christ, W.W. Wiegant, P.P. van Buul, M. Han, M.Z. Zdzienicka, J.M. Stark, M. Jasin, Suppression of the DNA repair defects of BRCA2-deficient cells with heterologous protein fusions, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 8768–8773. [7] D.S. Shin, L. Pellegrini, D.S. Daniels, B. Yelent, L. Craig, D. Bates, D.S. Yu, M.K. Shivji, C. Hitomi, A.S. Arvai, N. Volkmann, H. Tsuruta, T.L. Blundell, A.R. Venkitaraman, J.A. Tainer, Full-length archaeal Rad51 structure and mutants: mechanisms for RAD51 assembly and control by BRCA2, EMBO J. 22 (2003) 4566–4576. [8] A.B. Conway, T.W. Lynch, Y. Zhang, G.S. Fortin, C.W. Fung, L.S. Symington, P.A. Rice, Crystal structure of a Rad51 filament, Nat. Struct. Mol. Biol. 11 (2004) 791–796. [9] J.M. Short, Y. Liu, S. Chen, N. Soni, M.S. Madhusudhan, M.K. Shivji, A.R. Venkitaraman, High-resolution structure of the presynaptic RAD51 filament on single-stranded DNA by electron cryo-microscopy, Nucleic Acids Res. 44 (2016) 9017–9030. [10] J. Xu, L. Zhao, Y. Xu, W. Zhao, P. Sung, H.W. Wang, Cryo-EM structures of human RAD51 recombinase filaments during catalysis of DNA-strand exchange, Nat. Struct. Mol. Biol. 24 (2017) 40–46. [11] L. Pellegrini, D.S. Yu, T. Lo, S. Anand, M. Lee, T.L. Blundell, A.R. Venkitaraman, Insights into DNA recombination from the structure of a RAD51-BRCA2 complex, Nature 420 (2002) 287–293. [12] I. Brouwer, T. Moschetti, A. Candelli, E.B. Garcin, M. Modesti, L. Pellegrini, G.J. Wuite, E.J. Peterman, Two distinct conformational states define the interaction of human RAD51-ATP with single-stranded DNA, EMBO J. 37 (2018). [13] T. van der Heijden, R. Seidel, M. Modesti, R. Kanaar, C. Wyman, C. Dekker, Realtime assembly and disassembly of human RAD51 filaments on individual DNA molecules, Nucleic Acids Res. 35 (2007) 5646–5657. [14] N.L. van der Zon, R. Kanaar, C. Wyman, Variation in RAD51 details a hub of functions: opportunities to advance cancer diagnosis and therapy, F1000Res. 7 (2018). [15] J. Chen, M.D. Morrical, K.A. Donigan, J.B. Weidhaas, J.B. Sweasy, A.M. Averill, J.A. Tomczak, S.W. Morrical, Tumor-associated mutations in a conserved structural motif alter physical and biochemical properties of human RAD51 recombinase, Nucleic Acids Res. 43 (2015) 1098–1111. [16] T. Ishida, Y. Takizawa, I. Sakane, H. Kurumizaka, Altered DNA binding by the human Rad51-R150Q mutant found in breast cancer patients, Biol. Pharm. Bull. 30 (2007) 1374–1378. [17] M. Kato, K. Yano, F. Matsuo, H. Saito, T. Katagiri, H. Kurumizaka, M. Yoshimoto, F. Kasumi, F. Akiyama, G. Sakamoto, H. Nagawa, Y. Nakamura, Y. Miki, Identification of Rad51 alteration in patients with bilateral breast cancer, J. Hum. Genet. 45 (2000) 133–137. [18] C.G. Marsden, R.B. Jensen, J. Zagelbaum, E. Rothenberg, S.W. Morrical, S.S. Wallace, J.B. Sweasy, The tumor-associated variant RAD51 G151D induces a hyper-recombination phenotype, PLoS Genet. 12 (2016) e1006208. [19] M.C. Silva, M.D. Morrical, K.E. Bryan, A.M. Averill, J. Dragon, J.P. Bond, S.W. Morrical, RAD51 variant proteins from human lung and kidney tumors exhibit DNA strand exchange defects, DNA Repair (Amst.) 42 (2016) 44–55.

Authors’ contributions JS and WKH conceived the study. JS conducted experimentation. JS and WKH analyzed data and interpreted results. JS and WKH wrote the article.

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