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. 2026 Jun 15;3(2):ugag032. doi: 10.1093/narmme/ugag032

A yeast recombination system recapitulates the length polymorphism of a microsatellite implicated in human social behavior

Catie E Kean 1,#, Elina A Radchenko 2,#, Noah M Lindenbaum 3, Grace Materne 4, Sergei M Mirkin 5,✉
PMCID: PMC13294672  PMID: 42367581

Abstract

Arginine Vasopressin (AVP) modulates social and sexual behaviors in mammals through its receptor V1a. Differences in receptor expression and the resulting behavioral changes have been linked to population-level variation in the 5′ flanking region of the AVPR1A gene, specifically, length polymorphism of the composite microsatellite RS3 ((CT)4TT(CT)n(GT)m). However, the source of this length variation is unknown. Here, using a genetically tractable system in Saccharomyces cerevisiae, we recapitulated the full spectrum of the RS3 length polymorphism observed in humans resulting from genetic recombination between two RS3 sequences. Our candidate gene analysis shows that transcription elevates the rate of these recombination events and implicates double-strand break repair via single-strand annealing.

Graphical Abstract

Graphical Abstract.

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Introduction

Short tandem repeats (STRs), otherwise known as microsatellites, consist of 1-to-9 bp repetitive units [1]. These sequences comprise up to 6% of the human genome and are associated with heightened instability [2]. Massive expansions of structure-prone STRs result in over 60 hereditary diseases in humans, collectively termed repeat expansion disorders (REDs) (reviewed in [3]). However, even small variations in repeat length have been shown to alter gene expression by modifying chromatin structure, transcription, or splicing [4].

Beyond pathogenicity, microsatellite length polymorphism has been implicated in divergences in social and sexual behavior [5]. A notable example comes from comparing prairie voles (Microtus ochrogaster) with meadow voles (Microtus montanus). While these species are nearly identical, they exhibit distinct social behaviors. Meadow voles are asocial, non-parental, and promiscuous, while prairie voles are highly affiliative, biparental, and monogamous [6]. This difference has been attributed to decreased expression of the arginine vasopressin receptor V1a (V1aR) in the ventral forebrain of meadow voles relative to prairie voles [7]. Strikingly, this differential expression has been shown to depend on length variation in composite microsatellites located in the 5′ flanking region [6].

Human AVPR1A is nearly identical to the vole homolog; however, the composition of the microsatellite repeats upstream of the promoter is different. In humans, a (GT)n repeat at the -3956 position is followed by a composite microsatellite, RS3 ((CT)4TT(CT)n(GT)m) at -3625, and then a (GATA)n microsatellite, RS1, at –500 [8]. Length variation in the RS3 microsatellite has been linked to significant divergences in mating, aggression, and sexual behaviors [9, 10]. There are at least 16 distinct alleles of AVPR1A in the population due to the RS3 length polymorphism [11, 12]. Alleles with shorter RS3 repeats have been linked to social anxiety, poor pair bonding, autism, and aggression in humans [13–18]. However, the molecular mechanisms underlying the RS3 length polymorphism remain elusive.

The RS3 sequence possesses the intrinsic capability to form alternative DNA structures. The (GT)n component has potential to form left-handed Z-DNA, while its (CT)n repeats could form triplex H-DNA [19]. These alternative structures are known to trigger various forms of genome instability, including large-scale repeat expansions and contractions [20, 21]. Furthermore, slippage of repetitive DNA strands during DNA replication and repair, or misalignment of repetitive DNA strands in mitotic or meiotic recombination, can generate variation in STR length [19]. These events have been observed in simple microsatellites; however, the mechanisms responsible for the length polymorphism of composite microsatellites remain largely unknown.

Here, we sought to determine whether genetic recombination between RS3 microsatellites can recapitulate the length polymorphism observed in the human population. We designed a yeast experimental system to track intramolecular recombination events between the two copies of the RS3 microsatellite. Indeed, we found that misalignment during recombination produces the full spectrum of variability in the human population. Using candidate gene analysis, we found that transcription elevates the rate of intramolecular recombination between the RS3 copies, which likely involves double-strand break (DSB) repair via single-strand annealing (SSA)

Materials and methods

Plasmid construction for the fragility assay

Two plasmids were synthesized for the fragility assays: The PGAL1-UR-RS3-A3 plasmid, which contained the RS3 repeat, and PGAL1-UR-Intron-A3, which lacked the RS3 microsatellite (Supplementary Table S5). The construction procedures for both plasmids are outlined in the Supplementary Data.

Arm loss fragility assay

The plasmids, pYES3-G4G1C1-TET644 (control) and pYES3-G4G1C1-TET644-RS3, were digested with SmiI to integrate into the SMY706 yeast strain on chromosome V proximal to a CAN1 selection marker of the SMY923 strain [22] (Supplementary Table S5). Successful transformation in yeast was confirmed by PCR (Supplementary Table S6). Proper integration and orientation were confirmed with Sanger sequencing. The YER5 strain contained the PGAL1-UR-Intron-A3 cassette, and the YER7 strain contained PGAL1-UR-RS3-A3 (Supplementary Table S7).

Both YER5 and YER7 strains were grown on YPD (6 days) or YPGal (7 days) to produce large colonies. Cultures were diluted and plated on YPD and 5-FOA/canavanine media with galactose (Supplementary Table S8). Selective plates enabled the identification of arm-loss or gross chromosomal rearrangement events, as cells survived only when both CAN1 and URA3 were lost, conferring resistance to both drugs. During the selection phase, YPD plates were incubated for 3 days, while selective plates were incubated for 5 days at 30°C. The frequency of 5-FOA/canavanine-resistant colonies was calculated as the rate of arm-loss (GCR, or Gross Chromosomal Rearrangements) events per cell per generation [23] (Supplementary Table S9).

Plasmid construction for the recombination assay

Two plasmids were synthesized for the fragility assays. The PGAL1-UR-RS3-ADE2-RS3-A3 plasmid, which contained two RS3 repeats, and PGAL1-UR-Control-ADE2-Control-A3, which harbored two control sequences (Supplementary Table S5). The construction procedures for both plasmids are outlined in the supplemental information.

Recombination assay

An ade2∆ strain (YER9) was transformed with the PGAL1-UR-Control-ADE2-Control-A3 and PGAL1-UR-RS3-ADE2-RS3-A3 cassettes, generating strains YER109n/3-1 (control) and YER17 and YER94/31 (RS3) (Supplementary Table S7). Successful transformation in yeast was validated with PCR (Supplementary Table S6). Proper integration and orientation were confirmed with Sanger sequencing. The cassettes were positioned downstream of the ARS306 origin in a direct orientation, thus transcription and replication within the construct proceeded co-directionally.

Recombination rates between RS3 microsatellites were quantified using a protocol similar to that used for arm-loss fragility assays. During the non-selective phase, individual colonies were grown on glucose-containing media (YPD) for three days, while colonies on galactose-containing media (YPGal) required four days of growth. For slow-growing strains (hpr1Δ, dna2-H547A), colonies were allowed to grow until reaching approximately 2 mm in diameter. In the selective phase, 12 colonies were suspended in sterile water and serially diluted by measures of 1:10 to achieve a maximum dilution factor of 105. Following dilution, measured volumes from each colony suspension were plated on YPD (non-selective) media and on galactose-containing media lacking uracil (Gal/ura-) for selection (Supplementary Table S8). YPD plates were grown at 30°C for three days and selective plates for five days before obtaining colony counts for both conditions (Supplementary Table S10). Colony counts from selective plates, and corresponding non-selective plates, were entered into the FluCalc program []. This program employs the Ma-Sandri–Sarkar maximum likelihood estimation method. This method is considered one of the most faithful estimators of mutation rate, as it compares across practically infinite ranges of mutation rates and accounts for dilution by incorporating plating efficiency calculations. Differences in recombination rate were considered statistically significant when the 95% confidence intervals did not overlap between strains.

Repeat length analysis with single colony PCR

DNA was extracted from red colonies on selective media for PCR analysis. Genomic DNA was extracted from randomly selected colonies using lyticase (Sigma–Aldrich L4025). The recombined repeat was amplified using primers binding upstream of the first RS3 sequence (Opt1-2_in_s_fwd) and downstream of the second RS3 sequence (RS3_NgoMIV_short_R) (Supplementary Table S6). PCR products were visualized using gel electrophoresis on 1.5%–2% agarose gels until adequate separation was achieved. Bands were categorized as normal length, imperfect, or unclassified. Unclassified fragments underwent additional analysis on 2% agarose gels with wider wells. Control sequences from perfectly recombined RS3 fragments were loaded between unknown samples for comparison. The total frequency of imperfect recombination events was calculated for each strain. The proportions of “perfect” and “variable” recombination events were determined based on the random samples analyzed by PCR and then applied to the complete dataset. The exact sequence of imperfectly recombined RS3 repeats was assessed by Sanger sequencing to characterize the distribution of polymorphisms.

Gene deletion methodology

Candidate gene analysis was conducted by removing nonessential genes from Saccharomyces cerevisiae wild-type strains YER17/1 or YER94/31 and employing either standard gene replacement techniques or CRISPR–Cas9-mediated genome editing, as described by [24]. For conventional gene replacement without DSB induction, a plasmid containing a selectable marker [kanamycin (KAN), nourseothricin acetyl transferase (NAT), or hygromycin resistance (HYG)] was amplified using primers that incorporated homology to the targeted gene (Supplementary Table S5). For CRISPR–Cas9 knockout fragment construction, primers were designed with partially complementary sequences that were half homologous to a region within each target gene and half to the pRCC_N or pRCC_H plasmids [24] (Supplementary Table S5). The Cas9 plasmid backbone was amplified using Thermo Scientific high-fidelity Phusion polymerase with these target-specific primers, producing linear DNA fragments encoding NAT- or HYG-resistance, Cas9, and the gene-targeting sgRNA. Repair template oligos were designed with overlapping sequences from the gene’s flanking regions and assembled via Phusion PCR. Specific information about knockout fragments, oligonucleotides used for synthesis, repair templates, and validation primers is available upon request.

Transformations were performed using a high-efficiency lithium acetate protocol modified from [25]. Donor DNA was introduced into wild-type yeast using a solution containing 0.1 M lithium acetate, 10 mg/ml salmon sperm DNA carrier, and 33% polyethylene glycol. For conventional gene replacement, 1000 ng of the DNA knockout fragment was added, whereas for CRISPR–Cas9 editing, 2 μl of the pRCC plasmid and 1000 ng of the repair template were used. Transformation mixtures were incubated at room temperature for 25 min, followed by 42°C for 25 min. Transformed cells were plated on YPD media supplemented with uracil and adenine (YPDUA) and replica-plated to selective media after 1–2 days, or grown in liquid YPDUA for one hour before direct plating on selective media. After two days, DNA was extracted from selective plate colonies, and successful alteration was confirmed using locus-specific primers. Fragment presence/absence and size were assessed using PCR and gel electrophoresis (Supplementary Table S6). Confirmed colonies were isolated on YPEG media and successful transformants stored at −80°C in liquid YPDUA with 14.2% glycerol. All strains used in this study and their corresponding genotypes are listed in Supplementary Table S7.

Results

The RS3 microsatellite does not promote fragility in yeast

We previously demonstrated that repetitive DNA regions and secondary structures can increase chromosome fragility [26, 27]. Therefore, we first examined whether the RS3 microsatellite induces fragility using Kolodner’s arm-loss assay [27]. To this end, the PGAL1-UR-RS3-A3 cassette or the PGAL1-UR-Control-A3 cassette carrying a non-repetitive sequence in place of RS3 were integrated centromere-proximal to the CAN1 gene on the non-essential left arm of chromosome V (Fig. 1A).

Figure 1.

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A system to detect chromosomal fragility mediated by the RS3 repeat. (A) The PGAL1-UR-RS3-A3 cassette was integrated upstream of the CAN1 gene on the non-essential arm of chromosome V for an arm loss assay. The formation of a double-stranded break leads to loss of the cassette and CAN1, resulting in resistance to 5-FOA and canavanine. (B) Arm-loss rates for strains harboring the PGAL1-UR-RS3-A3 cassette and “control” strains with a non-repetitive sequence in place of RS3. Assays were conducted on both glucose and galactose. Error bars mark 95% confidence intervals.

In the event of DSB formation, the non-essential left arm of chromosome V, including part of the PGAL1-UR-RS3-A3 cassette and CAN1 gene, would be lost. These events could be detected by colony growth on plates containing both canavanine and 5-FOA. As simultaneous CAN1 and URA3 deletions are extremely rare, colony growth on canavanine + 5-FOA media is a proxy for arm loss-mediated DSBs.

We performed arm loss assays with the control and RS3 constructs and with growth on both glucose and galactose-containing media. We observed no significant differences in the rate of arm loss between the control sequence and RS3 on both glucose and galactose (Fig. 1B). However, transcription through these cassettes, induced by galactose, caused an 8.5-fold increase in arm loss for the RS3-cassette and a 17.4-fold increase for the non-repetitive control (Supplementary Table S1).

RS3 microsatellites can recombine and generate length variation

We created a new yeast experimental system to measure the rate of intrachromosomal recombination between RS3 microsatellites. In the PGAL1-UR-RS3-ADE2-RS3-A3 cassette, two identical RS3 microsatellites of the base composition (CT)4TT(CT)8(GT)24 were inserted into the artificial intron of URA3, flanking the ADE2 gene, and placed under the control of the galactose promoter (Fig. 2A). Before recombination, this composite intron measured 2824 bp – well above the splicing length limit for yeast [28]. Thus, cells carrying the PGAL1-UR-RS3-ADE2-RS3-A3 cassette are Ura- in the presence of galactose and Ade+, since the intron contains a full copy of the ADE2 gene [29]. Crucially, the two RS3 repeats are the only homologous sequences within this cassette. Recombination excises the ADE2 gene, shortening the intron to 512 bp and restoring URA3 gene function. These recombination events are, therefore, detected as Ura + Ade- (red) colonies growing on galactose-containing media.

Figure 2.

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A system to detect recombination between RS3 repeats. (A) Two RS3 repeats, (CT)4TT(CT)8(GT)24, flanking the ADE2 gene, were artificially inserted into an intron of the URA3 gene under the GAL1 promoter control. The resultant cassette – UR-RS3-ADE2-RS3-A3 – was integrated into the chromosome III adjacent to the ARS306 replication origin. The total length of the intron is 2824 bp. Recombination events between the RS3 sequences excised the ADE2 gene, shortening the intron to 512 bp and enabling its splicing. (B) Recombination rates for strains harboring the construct depicted in (A) (blue) and strains with an identical construct, but in the place of the RS3 microsatellite, a non-repetitive control sequence of the same length was integrated (red). Lighter columns represent cells grown on glucose during the nonselective phase, and dark columns represent cells grown on galactose. Error bars indicate the 95% confidence intervals of each strain.

Fluctuation assays were used to calculate the recombination rate. Individual yeast colonies were first grown on non-selective media containing either glucose or galactose, followed by plating onto the Ura- selective media containing galactose to induce transcription. RS3 microsatellites recombined at a rate of ∼1.3 × 10–6 per cell per generation when the reporter was repressed in the presence of glucose (Fig. 2B and Supplementary Table S2). Transcriptional induction led to a 4-fold increase in recombination rate to ∼5.2 × 10–6 per cell per generation.

Unexpectedly, we observed similar recombination rates for the PGAL1-UR-Control-ADE2-Control-A3 cassette, which contained two identical, non-repetitive DNA sequences of the same length as the RS3 sequence (Fig. 2B and Supplementary Table S2). Thus, we conclude that the RS3 repeat does not serve as a recombinational hot spot, at least during intrachromosomal recombination, regardless of transcriptional status.

We then investigated whether intrachromosomal recombination between RS3 repeats could generate length polymorphism (Fig. 3A). We extracted DNA from Ura+ Ade- colonies, amplified the region containing the RS3 repeat by PCR, and visualized the products by gel electrophoresis. We observed both repeat expansions and contractions and determined that imperfect recombination occurs at a rate of ∼5.2 × 10–7 per cell per generation when transcription is induced by galactose, corresponding to ∼10% of all events. Subsequently, the spectrum of length polymorphism was constructed via Sanger sequencing (Fig. 3C and Supplementary Table S3). Variability was primarily present in the (GT)n segment, with a minimum of 11 and a maximum of 31 units (Fig. 3C and Supplementary Table S3). The (CT)n region demonstrated a smaller range of 7 to 9 units (Supplementary Table S3). Out of the 54 sequenced colonies that displayed variation, two were (CT)7 and one was (CT)9 (Supplementary Table S3). Intramolecular recombination in the presence of glucose generated a similar extent of the RS3 length polymorphism (Fig. 3C). In the (GT)n segment, n varied between 11 and 41 units (Fig. 3C and Supplementary Table S4), while of the 62 sequenced colonies that displayed variation, two were (CT)7 and one was (CT)9 (Supplementary Table S4). Thus, in our experimental system, intramolecular recombination practically fully recapitulated the length polymorphism of the GT segments of the RS3 repeat observed in humans [11].

Figure 3.

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Length polymorphism in RS3 following recombination. (A) Schematic of primer annealing within URA3. Primers Opt1-2_in_s_fwd (top strand) and RS3_NgoMIV_short_R (bottom strand) were used to amplify the RS3 region from Ura+ colonies. (B) PCR products showing the lengths of the RS3 repeat tracts from colonies grown on selective plates. The amplified region is expected to be 120 bp; a 50 bp ladder is used in the distal lanes. Red arrows indicate “out-of-register,” or “imperfect” recombination events. Blue arrows show the target allele (CT)4TT(CT)8(GT)24, or “perfect” recombination events. (C) Length distribution of the (GT)n repeat based on Sanger sequencing from 50 Ura + Ade- colonies grown on galactose (left) and 59 Ura + Ade- colonies grown on glucose (right). Dotted lines represent the consensus sequence of n = 24. Colonies displayed polymorphism according to preliminary gel electrophoresis analysis.

The genetic mechanism of RS3 recombination

Prior to recombination, the RS3 microsatellite in our construct [(CT)4TT(CT)8(GT)24] measures 74 bp. This length is at the lower end for synthesis-dependent strand annealing (SDSA) and double-strand break repair (DSBR)—two key homologous recombination pathways in yeast [30, 31]. Similarly, it satisfies the minimal length requirement for SSA pathway but lies significantly below its optimal length. At the same time, it is much longer than the 5-to-20 bp microhomology required for microhomology-mediated end joining (MMEJ) [32, 33]. Therefore, it was not immediately clear which pathway promoted recombination between RS3 repeats in our system.

The significant stimulatory role of transcription in intramolecular recombination (Fig. 2B) led us to examine transcription-associated recombination (TAR) factors. DNA damage generated during transcription is often repaired by recombination [34]. Thus, TAR describes a pattern of conditions in which actively transcribed regions are commonly associated with recombinogenic repair [35]. Several mechanisms have been proposed to explain how transcription enhances recombination, including increased exposure of DNA to recombination proteins and DNA-damaging agents, collisions between replication machinery and the transcription apparatus or R-loops, as well as changes in DNA topology [34, 36]. Therefore, we subsequently examined the role of proteins commonly implicated in TAR in our system [33].

The THO complex counteracts TAR by ensuring proper messenger ribonucleoprotein (mRNP) synthesis and export [37]. This complex consists of four core proteins in yeast: Hpr1, Tho2, Mft1, and Thp2. Both Thp2 and Hpr1 are recruited specifically to transcriptionally active chromatin, where they function at the interface between transcription elongation and mRNA export [37–39]. Mutations in either gene result in R-loop accumulation and transcription-dependent hyperrecombination [37]. These THO proteins play a large role in our system, as deletions of THP2 and HPR1 genes resulted in hyperrecombination (Fig. 4 and Supplementary Table S2).

Figure 4.

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Genetic control of intramolecular recombination between transcribed RS3 repeats. Error bars indicate the 95% confidence intervals of each strain. Dashed lines indicate the 95% confidence intervals of the wild-type strain.

To further elucidate the involvement of R-loops, we created strains deficient in both RNase H1 and RNase H2. Both proteins cleave RNA–DNA hybrids but have distinct substrate preferences. Rnh1 requires at least four consecutive ribonucleotides to function effectively, while Rnh201 (the catalytic subunit of RNase H2) can process single ribonucleotides [40]. The double rnh1∆ rnh201∆ mutant completely abolishes RNase H activity resulting in R-loop accumulation [41]. This double mutant displayed hyper-recombination between RS3 repeats, supporting the involvement of R-loops in the pathway (Fig. 4 and Supplementary Table S2).

We also examined other proteins potentially involved in TAR. Topoisomerase I (Top1) resolves topological stress during transcription, preventing stalling and R-loop formation [34]. However, deletion of TOP1, showed no significant effect in our system (Fig. 4 and Supplementary Table S2). We also observed no effect from the deletion of DST1, which encodes the general transcription factor TFIIS, as well as the Rrm3 helicase [42, 43].

Having established transcription as a stimulatory force, we subsequently assessed the precise recombination mechanisms at play. The loss of Rad52—a central effector of recombination in yeast and/or its paralog Rad59 [44–47] resulted in a ∼3-fold reduction in recombination (Fig. 4 and Supplementary Table S2). Loss of Rad51, which catalyzes homologous DNA pairing and strand exchange, along with its cofactor Rad54, resulted in hyper-recombination between the repeats [48–50]. These results strongly argue against the involvement of SDSA or DSBR as both pathways require Rad51-mediated strand invasion. Instead, this data points to the SSA pathway, which depends on Rad52 and/or Rad59 for strand annealing [51, 52].

In SSA, repetitive segments on either side of a double stranded break are annealed following the DSB formation and end-resection [52]. This process generates single-stranded flaps containing the sequences between repetitive regions. The flaps are then removed, and fill-in synthesis performed. The structure-specific endonuclease heterodimer Rad1–Rad10 is essential for the flap removal [53–55]. In agreement with the SSA hypothesis, we observed a substantial 10-fold decrease in recombination in rad1∆ mutants (Fig. 4 and Supplementary Table S2). Furthermore, loss of Slx4, a scaffold protein responsible for recruiting and regulating Rad1–Rad10, caused 3-fold decrease in recombination [56].

In addition to its roles in SSA, the Rad1-Rad10 complex can interface with transcription machinery in transcription-coupled nucleotide excision repair (TC-NER), a specialized DNA repair pathway that targets lesions on the actively transcribed DNA strand [57, 58]. To determine whether the observed Rad1–Rad10 involvement reflects SSA or TC-NER activity, we deleted Rad26, the effector protein responsible for recognizing stalled polymerase complexes and recruiting repair machinery to sites of damage. Loss of Rad26 had no effect on recombination, suggesting TC-NER does not contribute to the repair events observed in our system (Fig. 4 and Supplementary Table S2). Therefore, Rad1–Rad10 activity is likely confined to 3′ flap removal during SSA.

SSA requires extensive 5′- to 3′-end resection to expose internal homologies. Thus, we subsequently examined the influence of end-resection proteins that generate single-stranded overhangs. Mre11 is a component of the MRX complex (Mre11–Rad50–Xrs2) which is essential for the initial phase of DNA end resection during DSB repair [59–62]. Long range resection occurs through two independent mechanisms: Exo1, a 5′→3′ exonuclease or Sgs1–Dna2 helicase-nuclease complex [60]. MRE11 and EXO1 gene knockouts did not affect intramolecular recombination in our system (Fig. 4 and Supplementary Table S2). Since DNA2 is an essential gene, we created a dna2-H547A mutant, which possesses reduced endonuclease activity and impaired end-resection [63]. This mutation had no effect on recombination. However, exo1∆sgs1∆ double mutant showed a significant 5-fold decrease (Fig. 4 and Supplementary Table S2) consistent with redundancy between these two long-range resection mechanisms.

RS3 microsatellites recombined and generated length polymorphism in the absence of transcription, albeit at reduced efficiency. To compare the genetic requirements of recombination depending on transcriptional status, we cultured in glucose to repress the GAL1 promoter and therefore, our cassette.

Loss of HPR1, RNH1, RNH201, and THP2 did result in hyperrecombination, indicating no involvement of TAR under glucose conditions (Supplementary Fig. S1). Furthermore, loss of SGS1 resulted in hyperrecombination only with transcription induction. Sgs1 has been shown to suppress recombination by resolving recombination intermediates [64, 65]. However, the transcription dependence of this effect instead points to Sgs1’s role in resolving co-transcriptional R-loops rather than canonical recombination suppression [66, 67].

Concurrently, the recombination mechanisms under both conditions share clear similarities: recombination is promoted by Rad52, Rad59, Rad1, and Slx4, and counteracted by Rad51. This suggests that SSA is likely responsible for recombination between the two RS3 copies regardless of transcriptional status, with transcription increasing frequency, likely by driving DSB formation within the reporter cassette. This aligns with our observation that transcription greatly elevated chromosomal fragility (Fig 1B). The source of DSBs during transcription repression remains unknown and will be addressed in future work. Unexpectedly, RAD27 deletion resulted in a roughly order-of-magnitude increase in recombination regardless of transcriptional status. We hypothesize that this hyper-recombination phenotype arises from persistent lesions caused by unligated Okazaki fragments [68], though future studies will be needed to verify this hypothesis.

Discussion

The RS3 microsatellite in the human AVPR1A gene displays length variability across populations with major physiological, behavioral, and evolutionary implications. However, the mechanism(s) responsible for this length polymorphism remain unknown. Our study investigated whether genetic recombination could be responsible for this phenomenon. To address this question, we constructed a genetically tractable system to study intrachromosomal recombination between two RS3 microsatellites in budding yeast.

While RS3 is not a recombination hot spot (Fig. 2B and Supplementary Table S2), length polymorphism was observed in ∼10% of all recombinational events, likely due to repeat misalignment (Fig. 3B). We were able to fully recapitulate the RS3 length polymorphism observed in humans in our yeast experimental system (Fig. 3C).

We observed an increase in recombination upon active transcription of the cassette, pointing to the involvement of TAR [34]. A central cog of this phenomenon is the accumulation of R-loops. Our data show that disruption of R-loop regulation at all levels, from prevention of aberrant elongation (THO complex), to unwinding (Sgs1), and finally, RNA cleavage (RNase H and possibly Rad27) [67, 68] affects recombination between RS3 repeats (Fig. 4 and Supplementary Table S2). Furthermore, we observed that transcription induction greatly increases chromosome fragility (Fig. 1B and Supplementary Table S1), supporting the general observation of transcription-mediated DSB formation.

The presence of R-loops is known to lead to the formation of DNA nicks primarily owing to the exposure of ssDNA on the non-template strand for transcription [57, 67]. For example, cytosine deaminases efficiently target ssDNA, converting cytosine to uracil, which triggers base excision repair, generating nicks [69]. Additionally, structure specific nucleases that recognize R-loops introduce nicks [70]. Unrepaired nicks generated during co-transcriptional R-loops may become recombinogenic DSBs during DNA replication.

It was previously shown that DSB-induced recombination results in microsatellite length polymorphism [71]. We hypothesize, therefore, that transcription through our experimental cassette creates DSBs in between the two RS3 microsatellites ultimately driving intramolecular recombination through the SSA pathway (Fig. 5). In alignment with this hypothesis, we observed substantial involvement of SSA effectors including annealing proteins Rad52 and Rad59 and the Rad1–Rad10 flap cleavage complex, SSA appears to frequently generate length variation owing to improper alignment of RS3 repeats (Fig. 5). This model can also explain the inhibitory role of the Rad51 protein in our system (Fig. 4). When Rad51 is present, DSBs formed during DNA replication are preferentially repaired via sister chromatid exchange (SCE) rather than SSA. Repair via SCE would preserve the general structure of the cassette, i.e. ADE2 gene flanked by RS3 repeats in the URA3 intron—an outcome that our system cannot detect.

Figure 5.

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Transcription modulated genomic instability, either a DSB or a nick converted to a DSB during replication triggers SSA. Here, the damage is depicted in the ADE2 locus. Exo1/Sgs1 perform resection to create single-stranded RS3 homologies. Rad52 aligns the homologies and generates flaps. Rad1–Rad10 cleaves the 3′ flaps. Gaps are filled in and ligated and a single RS3 microsatellite remains. Contractions result from out-of-register alignment of RS3 sequences followed by partial cleavage of RS3 by Rad1–Rad10 (left). Out-of-register annealing can also result in expansion events through incomplete alignment and fill in synthesis (right).

Intriguingly, the SSA pathway appears to be involved in intramolecular recombination in the absence of transcription, where it also drives RS3 length polymorphism. The mechanisms of DSB formation in this case remain unknown.

Overall, we describe a general genetic event, in which ∼70 bp-long sequences on the same chromosome can recombine in a mutagenic and potentially detrimental manner. We found that if these short homologies are intrinsically repetitive, the recombination process produce high levels of length polymorphism. Based on our data, this risk is compounded in actively transcribed regions, i.e. ∼60% of the human genome [72].

We speculate that recombination could also be responsible for the length polymorphism of the human RS3 microsatellite. In humans, recombination would most likely be interchromosomal between RS3 microsatellites on homologous chromosomes in meiosis or sister chromatids in rapidly dividing pre-meiotic germ cells. Alternatively, an intrachromosomal event could potentially occur between the (GT)n repeat ∼300 bp upstream and RS3 [8]. We expect that misalignment of repetitive strands during recombination, similar to what we observed in our system, would generate length polymorphism in humans and potentially manifest as differences in social behavior.

Supplementary Material

ugag032_Supplemental_File
ugag032_supplemental_file.pdf (677.1KB, pdf)

Acknowledgements

We thank Kate Mirkin for suggesting we examine the RS3 length polymorphism. We value the feedback and guidance from members of the Mirkin and Freudenreich labs at Tufts and the Lovett and Haber labs at Brandeis. We are especially grateful to Mitch McVey, Andres Aguilera, Eric Brown, and Shem Scott for their perspectives and thoughtful insights.

Author contributions: All authors have read and agreed to the published version of the manuscript.

Catie E. Kean (Conceptualization [equal], Data curation [equal], Formal analysis [equal], Investigation [equal], Methodology [equal], Validation [equal], Writing—original draft [lead], Writing—review & editing [equal]), Elina A. Radchenko (Conceptualization [equal], Data curation [equal], Formal analysis [equal], Investigation [equal], Methodology [equal], Supervision [supporting], Validation [equal], Writing—original draft [supporting], Writing—review & editing [supporting]), Noah M. Lindenbaum (Data curation [supporting], Formal analysis [supporting], Investigation [supporting], Writing—original draft [supporting]), Grace Materne (Data curation [supporting], Formal analysis [supporting], Investigation [supporting], Visualization [supporting]), and Sergei M. Mirkin (Conceptualization [equal], Formal analysis [lead], Funding acquisition [lead], Investigation [equal], Project administration [lead], Supervision [lead], Validation [equal], Writing—original draft [lead], Writing—review & editing [lead]).

Notes

Present address: Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Boulder, CO 80309, United States

Present address: Monte Rosa Therapeutics, Boston, MA 02118, United States

Present address: Albert Einstein College of Medicine, Bronx, NY 10461, United States

Contributor Information

Catie E Kean, Department of Biology, Tufts University, Medford, MA 02155, United States.

Elina A Radchenko, Department of Biology, Tufts University, Medford, MA 02155, United States.

Noah M Lindenbaum, Department of Biology, Tufts University, Medford, MA 02155, United States.

Grace Materne, Department of Biology, Tufts University, Medford, MA 02155, United States.

Sergei M Mirkin, Department of Biology, Tufts University, Medford, MA 02155, United States.

Supplementary data

Supplementary data is available at NAR Molecular Medicine online. 

Conflict of interest

The authors have no relevant financial or non-financial interests to disclose.

Funding

This work was supported by the National Institutes of Health [NIGMS R35GM130322 to S.M.M]; and the National Science Foundation [NSF-BSF 2153071 to S.M.M]. Funding to pay the Open Access publication charges for this article was provided by the National Institutes of Health.

Data availability

All data generated or analyzed during this study are included in this published article and its supplementary information files.

References

  • 1. Liu  Q, Tong  Y, Wang  K. Genome-wide detection of short tandem repeat expansions by long-read sequencing. BMC Bioinf. 2020;21:542. 10.1186/s12859-020-03876-w. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2. Lander  ES, Linton  LM, Birren  B  et al.  Initial sequencing and analysis of the human genome. Nature. 2001;409:860–921. [DOI] [PubMed] [Google Scholar]
  • 3. Chen  Z, Morris  HR, Polke  J  et al.  Repeat expansion disorders. Pract Neurol. 2025;25:204–16. 10.1136/pn-2023-003938. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4. Bagshaw  ATM. Functional mechanisms of microsatellite DNA in eukaryotic genomes. Genome Biol Evol. 2017;9:2428–43. 10.1093/gbe/evx164. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5. Hopkins  WD, Donaldson  ZR, Young  LJ. A polymorphic indel containing the RS3 microsatellite in the 5' flanking region of the vasopressin V1a receptor gene is associated with chimpanzee (Pan troglodytes) personality. Genes Brain Behav. 2012;11:552–8. 10.1111/j.1601-183X.2012.00799.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Young  LJ, Nilsen  R, Waymire  KG  et al.  Increased affiliative response to vasopressin in mice expressing the V1a receptor from a monogamous vole. Nature. 1999;400:766–8. 10.1038/23475. [DOI] [PubMed] [Google Scholar]
  • 7. Hammock  EAD. Functional microsatellite polymorphism associated with divergent social structure in vole species. Mol Biol Evol. 2004;21:1057–63. 10.1093/molbev/msh104. [DOI] [PubMed] [Google Scholar]
  • 8. Donaldson  ZR, Kondrashov  FA, Putnam  A  et al.  Evolution of a behavior-linked microsatellite-containing element in the 5' flanking region of the primate AVPR1A gene. BMC Evol Biol. 2008;8:180. 10.1186/1471-2148-8-180. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9. Donaldson  ZR, Young  LJ. The relative contribution of proximal 5′ flanking sequence and microsatellite variation on brain vasopressin 1a receptor (AVPR1A) gene expression and behavior. PLoS Genet. 2013;9:e1003729. 10.1371/journal.pgen.1003729. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10. Sadino  JM, Donaldson  ZR. Prairie voles as a model for understanding the genetic and epigenetic regulation of attachment behaviors. ACS Chem Neurosci. 2018;9:1939–50. 10.1021/acschemneuro.7b00475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11. Thibonnier  M, Graves  MK, Wagner  MS  et al.  Study of V1-vascular vasopressin receptor gene microsatellite polymorphisms in human essential hypertension. J Mol Cell Cardiol. 2000;32:557–64. 10.1006/jmcc.2000.1108. [DOI] [PubMed] [Google Scholar]
  • 12. Erasmus  JC, Klingenberg  A, Greeff  JM. Allele frequencies of AVPR1A and MAOA in the Afrikaner population. S Afr J Sci. 2015;111:1–6. [Google Scholar]
  • 13. Zhang  Y, Zhu  D, Zhang  P  et al.  Neural mechanisms of AVPR1A RS3-RS1 haplotypes that impact verbal learning and memory. Neuroimage. 2020;222:117283. 10.1016/j.neuroimage.2020.117283. [DOI] [PubMed] [Google Scholar]
  • 14. Meyer-Lindenberg  A, Kolachana  B, Gold  B  et al.  Genetic variants in AVPR1A linked to autism predict amygdala activation and personality traits in healthy humans. Mol Psychiatry. 2009;14:968–75. 10.1038/mp.2008.54. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15. Thompson  RR, Price  D, Burris  D  et al.  Effects of arginine vasopressin on human anxiety and associations with sex, dose, and V1a-receptor genotype. Psychopharmacology (Berl). 2024;241:1177–90. Published online 2024 Feb 15. 10.1007/s00213-024-06551-7. [DOI] [PubMed] [Google Scholar]
  • 16. Walum  H, Westberg  L, Henningsson  S  et al.  Genetic variation in the vasopressin receptor 1a gene (AVPR1A) associates with pair-bonding behavior in humans. Proc Natl Acad Sci USA. 2008;105:14153–6. 10.1073/pnas.0803081105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17. Procyshyn  TL, Hurd  PL, Crespi  BJ. Association testing of vasopressin receptor 1a microsatellite polymorphisms in non-clinical autism spectrum phenotypes. Autism Res. 2017;10:750–6. 10.1002/aur.1716. [DOI] [PubMed] [Google Scholar]
  • 18. Vollebregt  O, Koyama  E, Zai  CC  et al.  Evidence for association of vasopressin receptor 1A promoter region repeat with childhood onset aggression. J Psychiatr Res. 2021;140:522–8. 10.1016/j.jpsychires.2021.05.062. [DOI] [PubMed] [Google Scholar]
  • 19. Khristich  AN, Mirkin  SM. On the wrong DNA track: molecular mechanisms of repeat-mediated genome instability. J Biol Chem. 2020;295:4134–70. 10.1074/jbc.REV119.007678. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20. Shah  KA, Shishkin  AA, Voineagu  I  et al.  Role of DNA polymerases in repeat-mediated genome instability. Cell Rep. 2012;2:1088–95. 10.1016/j.celrep.2012.10.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21. Shah  KA, McGinty  RJ, Egorova  VI  et al.  Coupling transcriptional state to large-scale repeat expansions in yeast. Cell Rep. 2014;9:1594–602. 10.1016/j.celrep.2014.10.048. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22. Aksenova  AY, Greenwell  PW, Dominska  M  et al.  Genome rearrangements caused by interstitial telomeric sequences in yeast. Proc Natl Acad Sci USA. 2013;110:19866–71. 10.1073/pnas.1319313110. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Radchenko  EA, McGinty  RJ, Aksenova  AY. Quantitative analysis of the rates for repeat-mediated genome instability in a yeast experimental system. Methods Mol Biol. 2018;1672:421–38. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24. Generoso  WC, Gottardi  M, Oreb  M  et al.  Simplified CRISPR–Cas genome editing for Saccharomyces cerevisiae. J Microbiol Methods. 2016;127:203–5. 10.1016/j.mimet.2016.06.020. [DOI] [PubMed] [Google Scholar]
  • 25. Gietz  RD, Schiestl  RH. High-efficiency yeast transformation using the LiAc/SS carrier DNA/PEG method. Nat Protoc. 2007;2:31–4. 10.1038/nprot.2007.13. [DOI] [PubMed] [Google Scholar]
  • 26. Kim  H‐M, Narayanan  V, Mieczkowski  PA  et al.  Chromosome fragility at GAA tracts in yeast depends on repeat orientation and requires mismatch repair. EMBO J. 2008;27:2896–906. 10.1038/emboj.2008.205. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27. McGinty  RJ, Rubinstein  RG, Neil  AJ  et al.  Nanopore sequencing of complex genomic rearrangements in yeast reveals mechanisms of repeat-mediated double-strand break repair. Genome Res. 2017;27:2072–82. 10.1101/gr.228148.117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28. Shishkin  AA, Voineagu  I, Matera  R  et al.  Large-scale expansions of Friedreich’s ataxia GAA repeats in yeast. Mol Cell. 2009;35:82–92. 10.1016/j.molcel.2009.06.017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Hooks  KB, Delneri  D, Griffiths-Jones  S. Intron evolution in saccharomycetaceae. Genome Biol Evol. 2014;6:2543–56. 10.1093/gbe/evu196. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30. Pâques  F, Haber  JE. Multiple pathways of recombination induced by double-strand breaks in Saccharomyces cerevisiae. Microbiol Mol Biol Rev. 1999;63:349–404. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Al-Zain  AM, Symington  LS. The dark side of homology-directed repair. DNA Repair (Amst). 2021;106:103181. 10.1016/j.dnarep.2021.103181. [DOI] [PubMed] [Google Scholar]
  • 32. McVey  M, Lee  SE. MMEJ repair of double-strand breaks (director’s cut): deleted sequences and alternative endings. Trends Genet. 2008;24:529–38. 10.1016/j.tig.2008.08.007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Villarreal  DD, Lee  K, Deem  A  et al.  Microhomology directs diverse DNA break repair pathways and chromosomal translocations. PLoS Genet. 2012;8:e1003026. 10.1371/journal.pgen.1003026. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34. Gottipati  P, Helleday  T. Transcription-associated recombination in eukaryotes: link between transcription, replication and recombination. Mutagenesis. 2009;24:203–10. 10.1093/mutage/gen072. [DOI] [PubMed] [Google Scholar]
  • 35. Aguilera  A. The connection between transcription and genomic instability. EMBO J. 2002;21:195–201. 10.1093/emboj/21.3.195. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36. Browning  KR, Merrikh  H. Replication–transcription conflicts: a perpetual war on the chromosome. Annu Rev Biochem. 2024;93:21–46. 10.1146/annurev-biochem-030222-115809. [DOI] [PubMed] [Google Scholar]
  • 37. Chávez  S, Beilharz  T, Rondón  AG  et al.  A protein complex containing Tho2, Hpr1, Mft1 and a novel protein, Thp2, connects transcription elongation with mitotic recombination in Saccharomyces cerevisiae. EMBO J. 2000;19:5824–34. 10.1093/emboj/19.21.5824. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38. Chávez  S, Aguilera  A. The yeast HPR1 gene has a functional role in transcriptional elongation that uncovers a novel source of genome instability. Genes Dev. 1997;11:3459–70. 10.1101/gad.11.24.3459. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39. García-Rubio  M, Chávez  S, Huertas  P  et al.  Different physiological relevance of yeast THO/TREX subunits in gene expression and genome integrity. Mol Genet Genomics. 2008;279:123–32. 10.1007/s00438-007-0301-6. [DOI] [PubMed] [Google Scholar]
  • 40. Lockhart  A, Pires  VB, Bento  F  et al.  RNase H1 and H2 are differentially regulated to process RNA–DNA hybrids. Cell Rep. 2019;29:2890–900. 10.1016/j.celrep.2019.10.108. [DOI] [PubMed] [Google Scholar]
  • 41. Amon  JD, Koshland  D. RNase H enables efficient repair of R-loop-induced DNA damage. eLife. 2016;5:e20533. 10.7554/eLife.20533. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42. Clark  AB, Dykstra  CC, Sugino  A. Isolation, DNA sequence, and regulation of a Saccharomyces cerevisiae gene that encodes DNA strand transfer protein alpha. Mol Cell Biol. 1991;11:2576–82. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43. Ivessa  AS, Lenzmeier  BA, Bessler  JB  et al.  The Saccharomyces cerevisiae helicase Rrm3p facilitates replication past nonhistone protein-DNA complexes. Mol Cell. 2003;12:1525–36. 10.1016/S1097-2765(03)00456-8. [DOI] [PubMed] [Google Scholar]
  • 44. Mortensen  UH, Bendixen  C, Sunjevaric  I  et al.  DNA strand annealing is promoted by the yeast Rad52 protein. Proc Natl Acad Sci USA. 1996;93:10729–34. 10.1073/pnas.93.20.10729. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45. Shinohara  A, Shinohara  M, Ohta  T  et al.  Rad52 forms ring structures and co-operates with RPA in single-stranded DNA annealing. Genes Cells. 1998;3:145–56. 10.1046/j.1365-2443.1998.00176.x. [DOI] [PubMed] [Google Scholar]
  • 46. Sugawara  N, Ira  G, Haber  JE. DNA length dependence of the single-strand annealing pathway and the role of saccharomyces cerevisiae RAD59 in double-strand break repair. Mol Cell Biol. 2000;20:5300–9. 10.1128/MCB.20.14.5300-5309.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. Davis  AP, Symington  LS. The yeast recombinational repair protein Rad59 interacts with Rad52 and stimulates single-strand annealing. Genetics. 2001;159:515–25. 10.1093/genetics/159.2.515. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48. Ogawa  T, Yu  X, Shinohara  A  et al.  Similarity of the yeast RAD51 filament to the bacterial RecA filament. Science. 1993;259:1896–9. 10.1126/science.8456314. [DOI] [PubMed] [Google Scholar]
  • 49. Sung  P. Catalysis of ATP-dependent homologous DNA pairing and strand exchange by yeast Rad51 protein. Science. 1994;265:1241–3. 10.1126/science.8066464. [DOI] [PubMed] [Google Scholar]
  • 50. Wright  WD, Heyer  WD. Rad54 functions as a heteroduplex DNA pump modulated by its DNA substrates and Rad51 during D loop formation. Mol Cell. 2014;53:420–32. 10.1016/j.molcel.2013.12.027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51. Mott  C, Symington  LS. RAD51-independent inverted-repeat recombination by a strand-annealing mechanism. DNA Repair. 2011;10:408–15. 10.1016/j.dnarep.2011.01.007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52. Symington  LS, Rothstein  R, Lisby  M. Mechanisms and regulation of mitotic recombination in Saccharomyces cerevisiae. Genetics. 2014;198:795–835. 10.1534/genetics.114.166140. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53. Fishman-Lobell  J, Haber  JE. Removal of nonhomologous DNA ends in double-strand break recombination: the role of the yeast ultraviolet repair gene RAD1. Science. 1992;258:480–4. 10.1126/science.1411547. [DOI] [PubMed] [Google Scholar]
  • 54. Ivanov  EL, Haber  JE. RAD1 and RAD10, but not other excision repair genes, are required for double-strand break-induced recombination in Saccharomyces cerevisiae. Mol Cell Biol. 1995;15:2245–51. 10.1128/MCB.15.4.2245. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55. Mazón  G, Lam  AF, Ho  CK  et al.  The Rad1–Rad10 nuclease promotes chromosome translocations between dispersed repeats. Nat Struct Mol Biol. 2012;19:964–71. 10.1038/nsmb.2359. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56. Flott  S, Alabert  C, Toh  GW  et al.  Phosphorylation of Slx4 by Mec1 and Tel1 regulates the single-strand annealing mode of DNA repair in budding yeast. Mol Cell Biol. 2007;27:6433–45. 10.1128/MCB.00135-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 57. Petermann  E, Lan  L, Zou  L. Sources, resolution and physiological relevance of R-loops and RNA–DNA hybrids. Nat Rev Mol Cell Biol. 2022;23:521–40. 10.1038/s41580-022-00474-x. [DOI] [PubMed] [Google Scholar]
  • 58. Duan  M, Selvam  K, Wyrick  JJ  et al.  Genome-wide role of Rad26 in promoting transcription-coupled nucleotide excision repair in yeast chromatin. Proc Natl Acad Sci USA. 2020;117:18608–16. 10.1073/pnas.2003868117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59. Mimitou  EP, Symington  LS. Sae2, Exo1 and Sgs1 collaborate in DNA double-strand break processing. Nature. 2008;455:770–4. 10.1038/nature07312. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60. Zhu  Z, Chung  W-H, Shim  EY  et al.  Sgs1 helicase and two nucleases Dna2 and Exo1 resect DNA double strand break ends. Cell. 2008;134:981–94. 10.1016/j.cell.2008.08.037. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 61. Shim  EY, Chung  W‐H, Nicolette  ML  et al.  Saccharomyces cerevisiae Mre11/Rad50/Xrs2 and ku proteins regulate association of Exo1 and Dna2 with DNA breaks. EMBO J. 2010;29:3370–80. 10.1038/emboj.2010.219. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 62. Symington  LS. DNA repair: making the cut. Nature. 2014;514:39–40. 10.1038/nature13751. [DOI] [PubMed] [Google Scholar]
  • 63. Park  S, Karatayeva  N, Demin  AA  et al.  The secondary-structured DNA-binding activity of Dna2 endonuclease/helicase is critical to cell growth under replication stress. FEBS J. 2021;288:1224–42. 10.1111/febs.15475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 64. Sugawara  N, Goldfarb  T, Studamire  B  et al.  Heteroduplex rejection during single-strand annealing requires Sgs1 helicase and mismatch repair proteins Msh2 and Msh6 but not Pms1. Proc Natl Acad Sci USA. 2004;101:9315–20. 10.1073/pnas.0305749101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 65. Schmidt  KH, Pennaneach  V, Putnam  CD  et al.  Analysis of gross-chromosomal rearrangements in Saccharomyces cerevisiae. Methods Enzymol. 2006;409:462–76. [DOI] [PubMed] [Google Scholar]
  • 66. Chang  EY-C, Novoa  CA, Aristizabal  MJ  et al.  RECQ-like helicases Sgs1 and BLM regulate R-loop-associated genome instability. J Cell Biol. 2017;216:3991–4005. 10.1083/jcb.201703168. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 67. Aguilera  A, García-Muse  T. R loops: from transcription byproducts to threats to genome stability. Mol Cell. 2012;46:115–24. 10.1016/j.molcel.2012.04.009. [DOI] [PubMed] [Google Scholar]
  • 68. Liu  C-C, Chan  H-R, Su  G-C  et al.  Flap endonuclease Rad27 cleaves the RNA of R-loop structures to suppress telomere recombination. Nucleic Acids Res. 2023;51:4398–414. 10.1093/nar/gkad236. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 69. Lee  SY, Miller  KM, Kim  JJ. Clinical and mechanistic implications of R-loops in human leukemias. Int J Mol Sci. 2023;24:5966. 10.3390/ijms24065966. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 70. Su  X, Freudenreich  C. Cytosine deamination and base excision repair cause R-loop–induced CAG repeat fragility and instability in Saccharomyces cerevisiae. Proc Natl Acad Sci USA. 2017;114:E8392–401. 10.1073/pnas.1711283114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 71. Pâques  F, Richard  GF, Haber  JE. Expansions and contractions in 36-bp minisatellites by gene conversion in yeast. Genetics. 2001;158:155–66. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 72. ENCODE Project Consortium . An integrated encyclopedia of DNA elements in the human genome. Nature. 2012;489:57–74. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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Supplementary Materials

ugag032_Supplemental_File
ugag032_supplemental_file.pdf (677.1KB, pdf)

Data Availability Statement

All data generated or analyzed during this study are included in this published article and its supplementary information files.

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