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. Author manuscript; available in PMC: 2025 Nov 4.
Published in final edited form as: DNA Repair (Amst). 2025 Oct 14;155:103903. doi: 10.1016/j.dnarep.2025.103903

Overcoming natural replication barriers formed by DNA structures and the role of repositioning to the nuclear periphery

Jan Leendert Boer 1,#, Tyler M Maclay 1,#, Nolan T Caile 1, Catherine Freudenreich 1,*
PMCID: PMC12582535  NIHMSID: NIHMS2120602  PMID: 41175482

Abstract

Endogenous barriers to DNA replication, such as repetitive DNA, non-B DNA structures, and protein barriers present significant challenges to replication. Upon encountering one of these barriers, cells employ a number of strategies to ensure completion of replication. Some of these pathways operate at the stalled replication fork and others occur post-replicatively. These pathways vary both in their timing and the nuclear location in which they occur. Here we review how cells deal with endogenous sources of replication stress, with a focus on structure-forming DNA repeats, and our current understanding of how cells use nuclear positioning to facilitate the repair of natural replication barriers.

Keywords: DNA structure, non-B DNA, nuclear pore complex, replication fork barrier, fork restart, nuclear repositioning

DNA secondary structures as natural replication fork barriers

DNA must be faithfully replicated to maintain genomic stability and viability of cells. There are many impediments that can cause disruption to the canonical replication processes. These impediments can be the result of exogenous sources of DNA damage such as ionizing radiation, ultraviolet radiation, or mutagenic chemicals. Alternatively, they can be endogenous replication barriers such as protein barriers or DNA repeats forming secondary structures. These endogenous sources of replication stress have presented the opportunity to study how cells respond to natural barriers to replication.

Repetitive DNA can form non-B DNA secondary structures such as hairpins, cruciforms, intramolecular triplexes (H-DNA), or G quadruplexes (G4s). These collectively account for ~13% of the human genome (Guiblet et al., 2021; Khristich and Mirkin, 2020; Wang and Vasquez, 2022). The ability of these secondary structures to form depends on both the sequence and environment of the repeat (Mirkin and Mirkin, 2007). In general, the longer the DNA repeat the more likely it is to form a DNA secondary structure. Once formed, these repeat-derived structures can cause a litany of problems including replication fork stalling, interference with transcription, and inhibition of repair (Brown and Freudenreich, 2021; Gadgil et al., 2017; Khristich and Mirkin, 2020; Sinai and Kerem, 2023). If formed in single stranded DNA regions, even imperfect repeats can be the source of chromosomal rearrangements (Ait Saada et al., 2023).

DNA structure-induced replication stalls are most commonly and traditionally measured in vivo by two-dimensional gel electrophoresis (2DGE). Though stronger protein-mediated stalls have recently been detected by sequencing technologies (Claussin et al., 2022; Kara et al., 2021; Muller et al., 2019), to date these technologies haven’t achieved the sensitivity needed to detect DNA structure-mediated stalls in a chromosomal context. These stalls are most easily detected in conditions that favor DNA structure formation such as in negatively supercoiled replicating plasmids or by in vitro replication systems that use single-stranded DNA (ssDNA) templates. However, replication slowing by longer repeat tracts or more stable structures have been detected in a yeast chromosomal context by 2DGE (Kerrest et al., 2009; Kim et al., 2008; Viterbo et al., 2016; Zhang et al., 2013) or on human chromosomes by single molecule approaches (Gerhardt et al., 2016; Gerhardt et al., 2013). An approach using the timing of duplication of two fluorescently tagged loci has been used to measure replication fork slowing created by G4s and replication transcription collisions on a budding yeast chromosome (Dahan et al., 2018; Dovrat et al., 2018; Tsirkas et al., 2022). This technique was recently used to detect a (GAA/TTC)100 triplex-induced replication slowdown (Masnovo et al., 2024). In addition, replication-dependent effects on instability and fragility of chromosomally located expanded repeats are well documented (reviewed in (Leffak et al., 2016; Masnovo et al., 2022)).

The sequence content and length of repeat tracts both influence the strength of the replication fork stall. When visualized by 2DGE the hairpin-forming (CAG/CTG)n repeat, which contains an A/A or T/T interruption within the hairpin, stalls replication relatively weakly, while the more stable (CGG/GCC)n repeats and cruciform forming (AT/TA)n repeats create a stronger stall, and longer inverted repeats and triplex forming (GAA)n and (A2G3)n repeats create more pronounced fork stalls; in all cases tested, the severity of the stall increased with repeat tract length (Chandok et al., 2012; Hisey et al., 2024; Kaushal et al., 2019; Krasilnikova and Mirkin, 2004; Viterbo et al., 2016; Voineagu et al., 2008; Voineagu et al., 2009; Zhang and Freudenreich, 2007).

In contrast, proteins bound to DNA can elicit a particularly strong challenge to replication progression. In the budding yeast S. cerevisiae, the Fob1 protein barrier within the yeast ribosomal DNA (rDNA) promotes a severe replication fork stall and creates a recombination hotspot (Kobayashi and Horiuchi, 1996). In the fission yeast S. pombe, a replication fork barrier (RFB) caused by multiple copies of the replication termination sequence 1 (RTS1) bound by the Rtf1 protein leads to stalling of over 90% of replication forks (Ahn et al., 2005; Lambert et al., 2013; Lambert et al., 2005). The E. coli Tus protein binds to Ter modules, and this protein barrier strongly impedes replication fork progression in a directional manner and increases recombination at the locus (Larsen et al., 2014; Marie and Symington, 2022). The Tus-Ter complex is also able to stall replication forks in human cell culture (Willis et al., 2014).

Mechanisms of fork restart and post-replication repair at DNA structure-induced replication barriers

A replication fork stall needs to be overcome to complete DNA replication. Cells employ mechanisms such as dormant origin firing, translesion DNA synthesis (TLS), barrier bypass mechanisms, and repriming downstream of the barrier followed by reengagement of a new polymerase to complete DNA synthesis (Figure 1). The pathways vary in how error prone they are, their genetic requirements, and utilization at different types of natural replication fork barriers (reviewed in (Ait Saada et al., 2018; Berti et al., 2020; Conti and Smogorzewska, 2020; Kondratick et al., 2021)).

Figure 1: Pathways to overcome fork stalling at replication fork barriers.

Figure 1:

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Replication fork progression can be halted by a variety of structures on the DNA, including structure-forming DNA repeat sequences (e.g. a hairpin), proteins bound on the DNA, and DNA damage like interstrand crosslinks. A) Direct helicase unwinding of the DNA structure. B) A switch to a TLS polymerase which can replicate over the lesion “on the fly”. C) Fork reversal followed by recombination-dependent replication (RDR) by polymerase delta. D) After fork uncoupling, repriming downstream of the lesion and resumption of synthesis. E) Post-replication repair (PRR) pathways; alternative helicase unwinding of secondary structure could also occur during PRR. Not pictured: nuclease cleavage and subsequent DSB repair. Made using BioRender.

One of the simplest mechanisms to replicate through a barrier is to directly remove it at the fork. For DNA structures, this can be accomplished by helicase-mediated unwinding or unfolding of the structure (reviewed in (Leon-Ortiz et al., 2014; Polleys and Freudenreich, 2021; Sato and Knipscheer, 2023)). This is usually done by an accessory helicase, not the replicative helicase, and could occur either directly at the fork (Figure 1A) or post-replicatively (not shown). There are multiple helicases known to unwind G4 DNA, including PIF1, RTEL1, BLM, FANCJ and DDX11 (reviewed in (Freudenreich, 2020; Liu et al., 2021; Sauer and Paeschke, 2017)). G4 unwinding could occur at several stages of replication. For example, using an in vivo genetic assay that can detect replication efficiency of a G4 motif, it was shown that DDX11 resolves G4 structures after they are sensed by the fork protection complex located at the leading edge of the replisome (Lerner et al., 2020). This mechanism would be ideal for sensing pre-formed G4s encountered by the replisome. Using a reconstituted budding yeast replication system, it was shown that a G4 structure that forms after CMG unwinding can stall polymerase epsilon, leading to polymerase-helicase uncoupling, and that these G4s occurring between CMG and the polymerase are resolved by Pif1 but not other tested helicases (Williams et al., 2023). The translocase HLTF, which can unfold G4s in double-stranded DNA, suppresses G4 accumulation in all phases of the cell cycle and plays a role in restraining replication forks that encounter G4s to prevent DNA damage (Bai et al., 2024). Different helicases likely work in different contexts to resolve G4 structures and prevent genome instability, which could include in front of the fork, between the helicase and polymerase, or post-replicatively. In cases where the unwinding occurs in the context of a stalled fork this may occur in concert with fork uncoupling, though it isn’t known if this always the case. The mechanism of replication restart after the unwinding could also vary with the situation.

At hairpin-forming structures, the budding yeast Srs2 helicase was found to be needed for unwinding of (CTG) and (CGG) hairpins in vitro (Anand et al., 2012; Dhar and Lahue, 2008; Qiu et al., 2015), and for efficient progression of the replication fork through a (CGG)n repeat in vivo (Anand et al., 2012). A knockdown of the RTEL1 helicase led to an increase in the expansion of (CAG)n repeats in human cells and RTEL1 can compensate for the loss of yeast Srs2 to suppress fragility and expansions at (CAG)n repeats (Frizzell et al., 2014). However, hairpin structures may not always need accessory helicase unwinding. In the reconstituted budding yeast replication system, (CGG)n or (CG)n repeats placed on the leading strand template stalled polymerase epsilon, causing replisome uncoupling, but replication was able to be rescued by polymerase delta (Casas-Delucchi et al., 2022).

Long AT/TA repeats are unusual in that they can form not only hairpin structures in single-stranded DNA, but cruciform structures in double-stranded DNA (Kaushal and Freudenreich, 2019). BLM, WRN and FANCM helicases have all been implicated in unwinding secondary structures formed by AT-rich repeats due to their role in protecting genome instability at loci containing these sequences (see (Sinai and Kerem, 2023) for review). Both BLM and FANCM prevent the formation of double stranded breaks (DSBs) at an (AT/TA)n repeat (Li et al., 2024; Wang et al., 2018a; Wang et al., 2018b). A recent study showed that cells lacking both FANCM and the fork remodeler SMARCAL1 exhibit profound genome instability, with an accumulation of DNA breaks at genomic loci enriched in simple repeats that form secondary structures, including (AT/TA)n repeat tracts (Feng et al., 2024), highlighting the importance of helicases in unwinding secondary structures for cell health. The WRN helicase, which was shown to prevent Mus81-caused DSBs from forming at sites of expanded (AT/TA)n repeats in the human genome (van Wietmarschen et al., 2020), was shown to unfold cruciform structures in vitro (Mengoli et al., 2023).

At persistent replication fork stalls, the use of specialized TLS polymerases can be used to bypass the barrier (Figure 1B) (Maiorano et al., 2021; Paniagua and Jacobs, 2023; Rajpurohit et al., 2025). Specifically, it has been shown that polymerase eta (Pol η) is able to bypass UV-induced lesions at paused replication forks “on-the-fly” (Edmunds et al., 2008), preventing the accumulation of post-replicative gaps (Benureau et al., 2022; Maslowska et al., 2025). Pol η helps maintain genome stability at common fragile sites (CFSs) and can efficiently replicate non-B DNA sequences that are found within these loci (Barnes et al., 2017; Bergoglio et al., 2013). The FRA16D locus shows perturbed replication in Pol η deficient human cells by a DNA fiber assay (Twayana et al., 2021). Pol η has been shown to associate with forks stalled at R-loop stabilized G-quadruplex structures in human cells, assisting with fork bypass (possibly through a repriming mechanism) but leading to micronuclei formation and chromosomal instability (Pepe et al., 2025). Polymerase kappa (Pol κ) can also effectively replicate through non-B DNA sequences derived from FRA16D (Barnes et al., 2017; Mansilla et al., 2016). In vitro, it has been shown that TLS polymerase fidelity is uniquely impacted when synthesizing through G4 DNA with parallel topology (Stein et al., 2022). Polymerase zeta (Pol ζ) can also effectively replicate though several DNA structures (Das et al., 2024; Northam et al., 2014; Shah et al., 2012) (summarized below).

Alternatively, a stalled fork can undergo replication fork reversal, where the annealing of the two daughter strands leads to the formation of a four-way junction (Figure 1C) (Neelsen and Lopes, 2015). Replication forks reverse at the site of the lesion and slow both locally and globally in an ATR-dependent manner, to prevent genome instability (Mutreja et al., 2018). Some structure-forming repeats have direct evidence of causing fork reversal by either 2DGE or electron microscopy, specifically (GAA)n repeats (Follonier et al., 2013; Rastokina et al., 2023), (CAG)n repeats (Fouche et al., 2006; Kerrest et al., 2009; Nguyen et al., 2017), (AAGGG)n repeats (Hisey et al., 2024), and telomeric repeats (Vaurs et al., 2025).

Recombination-dependent replication (RDR) can be used to restart forks stalled by a protein-mediated RFB, often with the consequence of gross chromosomal rearrangements (GCRs) (Ait Saada et al., 2018; Berti et al., 2020; Kondratick et al., 2021). RDR is initiated following resection at a reversed fork, which promotes the loading of RPA and other repair factors onto the exposed single stranded DNA, followed by Rad51-dependent strand invasion into the homologous sequence ahead of the reversed fork (Figure 1C) (Schirmeisen et al., 2021). The restarted fork created by RDR is replicated in a semi-conservative manner, utilizing polymerase delta to replicate both the leading and lagging strands for up to 30 kb (Naiman et al., 2021). When a (CAG/CTG)70 repeat tract was placed in front of an RFB created by RTS1-bound protein Rtf1 in S. pombe cells, replication by the restarted fork exhibited low fidelity through the repeat tract, resulting in a high level of repeat expansions and contractions and revealing that tolerance to replication stress by fork restart comes at the cost of repeat instability (Gold et al., 2021). It is not yet known how often RDR is used as a recovery mechanism for forks stalled by a structure-forming repeat.

The replicative CMG helicase may instead bypass a replication fork barrier and reprime downstream of the barrier while the replicative polymerase remains stalled, a process termed fork uncoupling (Figure 1D) (Semlow and Walter, 2021; Sparks et al., 2019). CMG helicase-mediated bypass is coupled with repriming downstream of the barrier (Berti et al., 2020; Mouron et al., 2013), which requires the Primase-Pol α-Ctf4 complex in budding yeast (Fumasoni et al., 2015) and the DNA Polymerase PrimPol in humans (Conti and Smogorzewska, 2020; González-Acosta et al., 2021; Mellor et al., 2024). PrimPol dependent bypass of non-B DNA structures has been characterized for both G4 forming DNA and H-DNA forming (GAA)n repeats in avian cells (Schiavone et al., 2016; Šviković et al., 2018). In yeast it has been shown that fork uncoupling and repriming is prioritized at bulkier or stronger stalling lesions (Maslowska et al., 2025), therefore it could be particularly relevant at DNA structure barriers. Fork uncoupling and repriming leaves stretches of ssDNA, known as post-replicative gaps, which need to be filled in, either by TLS polymerases or by template switch (Figure 1E) (Chakraborty et al., 2023). Since DNA structures form preferentially in ssDNA, this situation presents an opportunity for their formation or persistence. In some cases, gap repair is delayed until late S/G2 phase and occurs in a PCNA ubiquitination-independent but Rad51-dependent manner; this late recombination repair pathway has been termed salvage recombination (Figure 1E) (Branzei et al., 2008; Joseph et al., 2022; Vanoli et al., 2010).

There is evidence from budding yeast that post-replicative mechanisms are important in repairing gaps occurring due to DNA structures. For (GAA/TTC)n repeats, Rad5-dependent template switch is a mechanism that can cause large-scale expansions (Shishkin et al., 2009) and CAG expansions that occurred in the absence of H4K12 and H4K16 acetylation depended on Rad52, Rad57, and Rad5 and were therefore arising through homology-mediated post-replication repair (PRR) events (House et al., 2014). On the other hand, short CAG expansions were elevated when proteins involved in the error-free branch of PRR were mutated (e.g. in rad18Δ, rad5Δ and PCNAK164R mutants), indicating that proper PRR inhibits some short repeat expansions (Daee et al., 2006). It may be that PRR is necessary to repair DNA structure-induced gaps, but this may come at the cost of instability at the repeat, especially in the context of longer triplet repeats. Direct investigation of this question showed that repair of a (CAG/CTG)70 tract exposed within a large gap resulted in repeat expansions and contractions (Polleys et al., 2023).

Recent studies have illuminated the specialized role of Pol ζ in replicating DNA structures. Following fork uncoupling, Rev1 and Pol ζ mediate repair of post-replicative gaps formed by bulkier UV-induced lesions, such as G-AAF (Guanine-acetylaminofluorene) adducts (Maslowska et al., 2025), which suggests that Pol ζ could play a post-replicative role at DNA structures. Indeed, Pol ζ has been shown to play a role in replication of GAA repeats and G-rich hairpins in S. cerevisiae (Northam et al., 2014; Shah et al., 2012). Recently, Pol ζ has been shown to play an important role in the stability of the Flex1 (AT)34 repeat (Das et al., 2024). Primer extension assays showed that both yeast and human Pol ζ can efficiently replicate through a hairpin formed by an AT repeat while Pol ε and Pol δ were stalled at the hairpin base. Pol ζ was also shown to prevent AT repeat-mediated chromosome breaks in yeast (Das et al., 2024) as well as facilitate replication and prevent breaks within AT-rich human centromeric heterochromatin (Yamin et al., 2021).

Both fork reversal and fork uncoupling have been observed as global responses to broad genotoxic stress via topoisomerase inhibitors, DNA synthesis inhibitors, or DNA damage induction (Zellweger et al., 2015), and both can act at replication forks stalled at barriers. In budding yeast, repriming can help restrain excessive reversed replication forks, which can accumulate in Primase-Pol α-Ctf4-deficient cells (Fumasoni et al., 2015), and in human cells under replication stress, suppression of fork reversal leads to the accumulation of PrimPol-dependent ssDNA gaps (Piberger et al., 2020). Therefore, fork-coupled and fork-uncoupled DNA damage response pathways can compensate for each other and may be promoted under different kinds of replication stress or at different DNA lesions. Both fork uncoupling and fork reversal occur in response to various DNA secondary structures, and both can lead to chromosome breakage or repeat expansions (Casas-Delucchi et al., 2022; Li et al., 2025; Rastokina et al., 2023; Williams et al., 2023).

Nuclease cleavage at structure-forming DNA repeats as a mechanism of recovery from replication stress

An alternative outcome is that the barrier leads to a DNA break, either at the fork or after replication. This outcome may be particularly relevant for DNA structures, which can be targeted by structure-specific endonucleases (SSEs), and which often contain single-stranded regions exposed to nucleases (Kaushal and Freudenreich, 2019). Rare fragile sites usually contain expanded structure-forming repeats (Mirceta et al., 2022). The first recognized case was the expanded CGG/GCC repeat that causes fragile X syndrome, and most recently it was shown that the expanded G4C2 repeat associated with some cases of ALS is a rare fragile site (Mirceta et al., 2024).

In an unbiased screen for breaks that occur in the human genome after ATR inhibition using the breaks identified by TdT labeling (BrITL) method, sequences rich in AT/TA repeats were found to be the most enriched (Shastri et al., 2018). The Mus81-Mms4 nuclease, in conjunction with the Slx1-4 and Rad1-10 (hXPF/ERCC1) nucleases, was found to be responsible for the cleavage of cruciform-forming lengths of the FRA16D-derived Flex1 (AT/TA)n repeat in yeast (Kaushal et al., 2019). This propensity of SSEs to cleave perfect cruciform-forming AT/TA repeats was further described in human cells. MUS81 and SLX4 were found to be responsible for the shattering of the chromosomes in WRN knock out cells that had defects in mismatch repair and had accumulated long AT/TA repeats (van Wietmarschen et al., 2020). Breaks caused by the MUS81-EME1 nuclease were mapped to expanded (AT/TA)n repeat alleles of a length and purity predicted to fold into a cruciform structure (Matos-Rodrigues et al., 2022; van Wietmarschen et al., 2020).

SSEs were also found to play a role at inverted repeats, with Mre11 cleaving hairpins with less than 9 nucleotides in the loop and Mus81 targeting transcribed inverted repeats (Ait Saada et al., 2021). In the absence of WRN-RPA complex binding at G4 DNA, MUS81 can cleave MRE11-extended daughter-strand gaps (Noto et al., 2025). Highlighting the vulnerability of having a DNA structure in a gap, it was shown that the presence of a (CTG)70 hairpin within a gap created a very fragile site, indicating that gap repair mechanisms are crucial to prevent DNA structure-induced breaks and downstream negative consequences (Polleys et al., 2023). The nucleases XPF-ERCC1 and XPG were found to cleave both H-DNA forming sequences (Zhao et al., 2018) and AT/TA repeats (Kaushal et al., 2019; Li et al., 2024). Indeed the XPF-ERCC1 complex has been found to cleave a variety of non-B DNA substrates (reviewed in (Li and Vasquez, 2022)).

In summary, SSEs have been shown to play an important role in the fragility of a variety of non-B DNA structures, and this cleavage may be crucial for recovery from replication stress. However, cleavage at a DNA structure can be especially deleterious because it can lead to hairpin-capped DNA ends that are difficult to repair and prone to rearrangements (Ait Saada et al., 2023; Al-Zain et al., 2023; Deng et al., 2015; Lobachev et al., 2007). Therefore, this mechanism of replication recovery could cause downstream problems and genome instability by the production of breaks that are difficult to repair or repaired by error-prone pathways.

Repositioning to the nuclear periphery of replication barriers and hard-to-repair lesions.

One distinct locus of DNA repair is the Nuclear Pore Complex (NPC). One of the components of the NPC is the Y-shaped Nup84 complex (hsNUP107–160 complex) which is associated with the outer ring of the nuclear pore (Akey et al., 2023). This complex plays a critical role in maintaining genome stability (Bennett et al., 2001); reviewed in (Simon et al., 2024). In yeast, the deletion of Nup84 results in increased sensitivity to hydroxyurea (HU), methyl methanesulfonate, camptothecin, phleomycin, and 4-Nitroquinoline 1-oxide (Chang et al., 2002; Gaillard et al., 2019; Loeillet et al., 2005). In human cells, deletion NUP107–160 complex components also resulted in increased genome instability and sensitivity to DNA damaging agents (Lemaître et al., 2012; Olivieri et al., 2020; Paulsen et al., 2009).

The NPC could play a part in facilitating repair by providing a scaffold for repair factors. Given that key players which facilitate post-translational modifications and DNA repair pathways anchor at the NPC, like the Slx5/Slx8 SUMO-targeted ubiquitin ligase (STUbL) (Nagai et al., 2008) and the SUMO protease Ulp1 (Zhao et al., 2004), the NPC may act as a hub where sites of DNA damage and the factors which can coordinate their repair colocalize. The NPC may also allow normally suppressed repair pathways by relieving constraints on them. It has been shown that the NPC promotes some types of recombination, including the break-induced replication (BIR) pathway or RDR, which can be used to repair one-ended breaks formed during replication (Lee et al., 2024; Mackenroth and Alani, 2020; Schirmeisen et al., 2021; Taddei and Gasser, 2012). Another scaffold at the nuclear periphery (NP) is Mps3 (Taddei and Gasser, 2012), a SUN domain containing nuclear envelope (NE) protein and spindle pole body (SPB) associated protein (Jaspersen et al., 2002). Mps3 is involved in clustering telomeres at the nuclear periphery (NP) (Antoniacci et al., 2007; Bupp et al., 2007) and the assembly of the spindle pole body (Friederichs et al., 2011), the microtubule organizing center in budding yeast. In contrast to the NPC, interaction of DSBs with nuclear envelope protein Mps3 represses ectopic recombination (Chung et al., 2015; Horigome et al., 2014; Oza et al., 2009).

Certain types of DNA damage and hard to repair lesions have been found to reposition to the NP, summarized in (Table 1) and illustrated in (Figure 2). In budding yeast, this includes persistent DSBs (Nagai et al., 2008; Oza et al., 2009), eroded telomeres in the absence of telomerase (Churikov et al., 2016; Khadaroo et al., 2009), subtelomeric DSBs (Chung et al., 2015), structure-forming CAG/CTG repeat tracts (Su et al., 2015; Whalen et al., 2020), DSBs in rDNA (Horigome et al., 2019), and R-loops in hybrid-accumulating conditions (Penzo et al., 2023). Simultaneous induction of many Cas9-induced DSBs in yeast Ty transposable elements or by zeocin exposure promoted formation of Tel1 (hsATM) foci at the NP, which resolved upon removal of Cas9, implicating the NP as a site of DSB repair (Auboiron et al., 2025). In Drosophila, DSBs induced in heterochromatic regions move to the nuclear periphery (Chiolo et al., 2011; Merigliano et al., 2025; Ryu et al., 2015). In human cells, DSBs induced by a nuclease targeting transcriptionally active regions, the topoisomerase poison etoposide, or a FokI-induced DSB, reposition to the NP (Le Bozec et al., 2024; Shokrollahi et al., 2024). Therefore, the shifting of DNA damage to the NP is conserved between species. In this review, we focus on natural replication barriers and damage resulting from replication stress, comparing to the data on induced DSBs when appropriate. For a recent comprehensive review on repositioning and repair of DSBs, see (Chiolo et al., 2025).

Table 1:

Described relationships between DNA lesion and association with the nuclear periphery

DNA damage that associates with the nuclear periphery Organism Known dependencies for repositioning (specific context as annotated) Source
Prolonged 0.2M HU treatment (≥ 1hour) Budding yeast Not determined (Nagai et al., 2008)
(Oza et al., 2009)
HU + MMS Treatment (0.2 M HU ± 0.033% MMS) Budding yeast Not determined (Nagai et al., 2008)
Eroded telomeres1,2,3 Budding yeast Siz1/Siz2-mediated SUMOylation2
Slx5/Slx82
Rad9/Rad242
Rad593 		1(Khadaroo et al., 2009)
2(Churikov et al., 2016)
3(Charifi et al., 2021)
Persistent DSB at MAT locus induced by HO endonuclease Budding yeast SUMOylation by Nse2 and Siz26
Slx56
Mec14, Tel14
Nup844, Mps35
Rad525, Cdc135 		4(Nagai et al., 2008)
5(Oza et al., 2009)
6(Horigome et al., 2016)
Expanded (CAG)n repeat (70–130 repeat units) Budding yeast Slx5/Slx87
Nse28 SUMOylation of substrates (incl. RPA, Rad52, Rad59, Smc5)8
Long-range resection8
Nup848, Nup19
Activation of the DNA replication checkpoint10.
Phospho-Cep310
DIM extrusion10, Kar310 		7(Su et al., 2015)
8(Whalen et al., 2020)
9(Aguilera et al., 2020)
10(Maclay et al., 2025)
Fob1 protein barrier in rDNA Budding yeast Mec1, Tel1, Tof1
Tof2, Csm1/Lrs4, Sir2
Nup120, Nup84		 (Horigome et al., 2019)
Replication fork barrier (Rtf1-bound RTS1 array) Fission yeast Pli1-mediated SUMOylation11
Rfp1/Rfp2/Slx811
Short-range resection11
Nup6012, Alm112
Rad5211, Rad51 activity11
Short & long-range resection11 		11(Kramarz et al., 2020)
12(Schirmeisen et al., 2024)
Zeocin induced DSBs (0.3 mg/mL zeocin)8,13 Budding yeast Pfa4-mediated Rif1 S-Acylation13 13(Fontana et al., 2019)
8(Whalen et al., 2020)
Damaged telomeres under replication stress Budding yeast Nup1 (Aguilera et al., 2020)
R loops Budding yeast Nse2monoSUMOylation of Rfa1 (Penzo et al., 2023)
Cruciform-forming (AT/TA)n repeats Budding yeast Nse2, Siz1, Siz2
Slx5
Slx4-associated SSEs
Rad51 activity		 (Boer et al., 2025)
DSBs in heterochromatin Drosophila Smc5/6/Nse215, dPIAS15
Dgrn15, Rad6015
ATR14, ATM14
Nup98, Nup88, Sec1317
Nuclear Basket/Outer Ring15
Koi/Spag415
Nuclear F-Actin, Myosin16 		14(Chiolo et al., 2011)
15(Ryu et al., 2015)
16(Caridi et al., 2018)
17(Merigliano et al., 2025)
Expanded (GAA)~310 repeats Human immortalized lymphoblastoid FRDA cells Unknown (Silva et al., 2015)
Aphidicolin stalled replication forks Human U2OS, HT1080 6TG, and IMR90cells ATR
F-Actin
Myosin II
mTOR		 (Lamm et al., 2020)
Replication defects at telomeres caused by POT1 dysfunction Human RPE1 and U2OS cells ATR
F-Actin		 (Pinzaru et al., 2020)
DSBs in transcriptionally active regions Human RPE1 and U2OS cells RNF419
NUP15318, 19, SUN118,19
PER119, PER219
dsbNET formation18 		18(Shokrollahi et al., 2024)
19(Le Bozec et al., 2024)
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Figure 2:

Figure 2:

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Illustration of the process of repositioning of damaged loci to the nuclear periphery. Bottom: Examples of difficult-to-repair DNA lesions which have been shown to reposition. Middle: ssDNA becomes exposed by resection, and SUMOylation, promoted by Smc5/6-associated Nse2/Mms21 and/or Siz1 and Siz2 SUMO ligases, accumulates on targeted substrates at the site of the lesion. SUMOylated proteins bind to the Slx5/Slx8 STUbL, which interacts with Nup84 of the Y complex to promote NPC association. The DNA replication and DNA damage checkpoints also contribute to relocation. Top left: Repair processes hypothesized to occur at the NPC, including homologous recombination (HR), recombination-dependent replication (RDR), and break-induced replication (BIR). Top right: Hypothesized fates of SUMOylated proteins at the nuclear pore complex, including STUbL-mediated ubiquitylation and degradation by the proteasome, or removal of SUMO by SUMO isopeptidases. Homologs (if named differently than in S. cerevisiae): SUMO ligases: scSiz1, scSiz2 = spPli1, dmPIAS, hsPIAS1–4; STUbLs: scSlx5/Slx8 = spRfp1/Slx8, spRfp2/Slx8, dmDgrn, hsRNF4, hsRNF111; SUMO proteases: scUlp1, spUlp1, dmUlp1 = hsSENP1, hsSENP2; Nup84 NPC subcomplex: scNup84 = spNup107, hsNUP107, dmNup107. Created with Biorender.com

Some types of replication fork impediments relocate to the NP. In yeasts, these include expanded CAG repeats (Maclay et al., 2025; Su et al., 2015; Whalen et al., 2020), stalls within structure forming telomere sequences (Aguilera et al., 2020), and RFBs like the Fob1-bound rDNA sequence in S. cerevisiae and the Rtf1-bound RTS1 sequence in S. pombe (RTS1-RFB) (Horigome et al., 2019; Kramarz et al., 2020; Schirmeisen et al., 2024). A ssDNA binding-defective POT1, a protein normally required for the stability of telomeres in human cells (Zade and Khattar, 2023), triggers mitotic DNA synthesis (MiDAS) at telomeres and causes them to reposition to the NPC (Pinzaru et al., 2020). Most recently, our group found that cruciform-forming AT/TA repeats of a size known to interfere with replication reposition to the nuclear periphery in budding yeast (Boer et al., 2025).

Replication fork inhibition by drugs that cause fork collapse or replisome dissociation can also result in relocation of loci to the nuclear periphery. In yeast, longer treatments of HU or HU+MMS that result in replication fork collapse result in repositioning to the NPC (Nagai et al., 2008; Whalen et al., 2020), and aphidicolin results in increased association of the replication fork with the NP in human cells (Lamm et al., 2020). In contrast, low doses of HU or MMS, which cause transient fork stalling or limited base damage do not cause repositioning (Nagai et al., 2008; Whalen et al., 2020; Wong et al., 2020). This indicates that not all types of replication barriers trigger a spatial shift, and replication forks may need to collapse or generate a secondary lesion to initiate repositioning to the NP.

Mechanism of repositioning of damaged loci to the nuclear periphery.

The repositioning of damaged loci to the nuclear periphery is a highly regulated process. Many lesions require the activity of checkpoint proteins to promote repositioning. HO-induced DSBs and Fob1-associated protein mediated fork stalls require activation of the Mec1 (hsATR) and Tel1 (hsATM) damage-sensing kinases to promote repositioning (Horigome et al., 2019; Nagai et al., 2008) (Figure 2). Similarly, heterochromatic DSBs require ATM and ATR for their repositioning (Ryu et al., 2015; Tsouroula et al., 2016). DSBs in human cells in transcriptionally active regions that reposition to the nuclear pore complex are clustered together in an ATM-dependent manner to create a clustered DSB “D compartment” (Arnould et al., 2023).

Aphidicolin-induced fork stalls require ATR to relocate (Lamm et al., 2020). In contrast, for (CAG)130 repeats, neither deletion of the ATR homolog Mec1 nor the ATM homolog Tel1 eliminated relocation (Maclay et al., 2025; Su et al., 2015). A recent study from our laboratory resolved this conundrum, showing that activation of the DNA replication checkpoint mediator Mrc1 is the crucial initial signal for (CAG)130 repeat repositioning, but that the Mec1 and Tel1 checkpoints can act redundantly to facilitate the process (Maclay et al., 2025). Similarly, when combined, defects in Rad9 (hs53BP1) and Rad24 (a component of the checkpoint clamp loader that facilitates recruitment of Mec1) suppressed repositioning of eroded telomeres and (CAG)n repeats (Churikov et al., 2016; Maclay et al., 2025). Thus, there is significant involvement of the checkpoint in sensing these difficult-to-repair lesions that reposition to the nuclear pore.

In addition to the phosphorylation mediated checkpoint response, SUMOylation cascades have been shown to be vital to promote lesion repositioning in all systems where this was studied (Table 1, Figure 2). For DSBs, homologous SUMO ligases respond across these different systems. For replication barriers caused by DNA structures, the Nse2/Mms21 SUMO ligase associated with the Smc5/6 complex, which is known to be recruited to and stabilize stalled forks (Tanasie et al., 2022) becomes more critical (Whalen et al., 2020). Though the full complement of SUMOylated targets important for relocation has yet to be determined, RPA, Rad52, Rad59, and Smc5 are known to be among them, acting in a partially redundant manner (Whalen, 2020). In fission yeast, Ulp1-mediated deconjugation of SUMO from pore-associated substrates has been shown to be critical to clear poly-SUMO chains to facilitate fork restart at the RTS1-RFB, describing an additional role for SUMO in regulating the events that occur after NPC association (Schirmeisen et al., 2024). SUMOylation has also been shown to be essential to promote the viability of human cells with POT1 mutations (Pinzaru et al., 2020), implying that SUMOylation may be required for damaged locus repositioning in human cells as well. In most cases where SUMOylation has been shown to be important for repositioning, a SUMO-targeted ubiquitin ligase has been shown to also be required (Kramarz et al., 2020; Le Bozec et al., 2024; Ryu et al., 2015; Su et al., 2015; Whalen et al., 2020).

These checkpoint and SUMOylation pathways are the best-described regulatory mechanisms that cells use to control the repositioning of damaged loci to the NPC. It is tempting to presume that these pathways interact to somehow facilitate the spatial shift, but that remains to be determined. Additionally, the requirement of SUMO-targeted ubiquitin ligases suggests that ubiquitination at these loci plays a role in signaling or promoting repositioning to the periphery, but how ubiquitin is involved in this pathway remains unclear.

How lesions physically move to the nuclear periphery is an area of active investigation (Chiolo et al., 2025; Shokrollahi and Mekhail, 2021). There appears to be significant involvement of the cytoskeleton in the movement of loci through the nucleus. Nuclear F-actin driven movement was observed for heterochromatic DSBs in Drosophila (Caridi et al., 2018; Dialynas et al., 2019). Similarly, in response to aphidicolin-induced replication fork stalling in human cells, nuclear F-actin and myosin II were shown to promote mobility and repositioning of the resultant fork stall in an ATR-dependent manner (Lamm et al., 2020). Recently, the actin-related Arp2/3 complex and type I myosins have been shown to promote DSB mobility in yeast, though the connection to repositioning remains unclear (Zhou et al., 2025).

Microtubules also play a role in damaged locus repositioning. In yeast, some types of DNA damage promote microtubule extrusion from the SPB. These were termed damage-induced microtubules (DIMs), and have been found associated with HO endonuclease or zeocin-induced DSBs, damage from HU+MMS or from a (CAG)130 tract (Maclay et al., 2025; Oshidari et al., 2018). Extrusion of the DIM from the SPB required the activity of the Rad9-mediated checkpoint for DSBs repairing by BIR (Oshidari et al., 2018) or the Mrc1-mediated checkpoint for uncoupled forks arising from a (CAG)130 tract (Maclay et al., 2025). The Mrc1-mediated checkpoint signal results in the phosphorylation of the kinetochore protein Cep3 leading to centromere release and formation of damage-inducible microtubules (DIMs) which promote relocation (Maclay et al., 2025). For other sytems, the mechanism by which the checkpoint facilitates relocation is unknown; it could be worthwhile to investigate if release of tethering constraints on chromosomes is similarly a critical step. Interestingly, these DIMs are sites of Rad52 liquid/liquid phase separated domain formation (Oshidari et al., 2020). In human cells, nuclear envelope tubules are induced in response to induced double strand breaks (dsbNETs) (Shokrollahi et al., 2024). These cytoplasmic microtubules associate with the LINC complex and NPCs embedded in the nuclear lamina to promote the invagination of the NE to “capture” DSBs (Shokrollahi et al., 2024). It is interesting that across eukaryotes, there seems to be significant involvement of the cytoskeleton in repositioning of damaged loci, though the specific details of which cytoskeleton components are involved vary.

Role of nuclear positioning in recovery from replication stress and repair: why relocate to the nuclear pore complex?

Many types of DNA damage, including transient fork stalling caused by hydroxyurea or base modifications, or DSBs that have a readily available template for repair, do not cause repositioning to the NPC (Nagai et al., 2008; Whalen et al., 2020; Wong et al., 2020), and these are repaired by a variety of well-studied pathways. However, for types of lesions that do shift their position, preventing the relocation pathway results in increased genome instability, indicating that it plays a genome protective role. With hard to repair DSBs, mutants involved with repositioning such as nup84Δ, and slx5Δ or slx8Δ showed an increased rate, by ~10-fold, of GCRs and gene conversion events (Nagai et al., 2008). Genome instability within a (CAG)130 repeat tract also increased in mutants that disrupted repositioning to the NPC: an increase in chromosome fragility (as measured by repeat-dependent yeast artificial chromosome end loss) and instability (as measured by contractions and expansions of the (CAG)130 repeat) were both observed (Su et al., 2015; Whalen et al., 2020). In S. pombe increased replication fork slippage events occurred when RTS1-RFB repositioning was impeded (Kramarz et al., 2020). These data all support a positive genome-protective role for NPC association.

What types of repair are promoted at the NPC? Using repair assays that score for repair of an induced DSB by BIR, decreased frequencies were observed in mutants defective for relocation (nup84, kar3, nup60, slx8) (Chung et al., 2015; Horigome et al., 2016), suggesting that the NPC location promotes BIR at a persistent DSB. However, for replication fork stalls, relocation to the NPC may lead to a different repair outcome. The relocation of the S. pombe RTS1-RFB was shown to be important for initiation of RDR fork restart (Kramarz et al., 2020; Schirmeisen et al., 2024). Forks stalled within telomeric repeats also relocate to the NPC, and molecular analysis of telomeres suggested that association with the NPC facilitates a fork restart mechanism that prevents breaks and unequal sister chromatid recombination (Aguilera et al., 2020; Pinzaru et al., 2020). For a long CAG/CTG tract, repeat expansions and contractions were increased in a Rad52-dependent manner in conditions where relocation did not occur (for example in the absence of Nup84 anchoring) (Su et al., 2015). This suggests that the NPC promotes Rad51-dependent recombination and prevents Rad52-mediated HR, a conclusion supported by the fact that there is greater Rad51 association with the repeat locus at the NPC (Whalen et al., 2020). Combined, these results suggest that at a fork stall, Rad51-dependent recombination may be initially constrained. However, if the damage persists, leading to accumulation of SUMOylated repair proteins, relocation to the NPC allows Rad51-dependent HR repair pathways to proceed.

Whether the damaged locus anchored to the NPC is a DSB or stalled fork, if these HR-dependent alternatives such as BIR and RDR are promoted, they would heal or prevent chromosome breaks. Though not necessarily error-free, these pathways may be preferable to alternatives such as Rad52-dependent annealing events which could lead to deletions or rearrangements. Indeed, recent results from our lab show that AT/TA repeats that are subject to Slx4 complex-mediated cleavage also relocate to the nuclear periphery, and this prevents Rad52-dependent single-strand annealing events that result in deletions (Boer et al., 2025).

What is the DNA damage signal that triggers relocation of replication barriers? As a wide variety of different types of DNA damage reposition, it appears that there is not one single structure that initiates the pathway. A stalled fork can undergo fork reversal followed by resection, which leaves ssDNA that can be coated by repair proteins that can be SUMOylated. This is what was hypothesized to occur at (CAG)130 repeats (Whalen and Freudenreich, 2020), since long CAG tracts are known to cause fork reversal (Kerrest et al., 2009) and resection proteins including Exo1 were required for the shift to the NPC. For the S. pombe RFB, only MRN-Ctp1-mediated short-range resection was required, along with Rad51 strand exchange activity, implicating a remodeled fork (Kramarz et al., 2020; Schirmeisen et al., 2024). We recently showed that the relocation of a (CAG)130 repeat is dependent on the phosphorylation of Mrc1, which occurs upon fork uncoupling, implicating an uncoupled fork as the initial signal (Maclay et al., 2025).

Alternatively, the replication fork can be reprimed downstream of the stalled polymerase, leaving a ssDNA gap behind the replication fork. New data from (Boer et al., 2025), found that an (AT)34 repeat shifts to the nuclear periphery in a replication-dependent manner, however, NP association occurred in late S or G2 phase, peaking 40–50 minutes after the replication fork is predicted to reach the repeat tract. If the fork were stalled for this long period of time the replication profile of the region should be altered, but this was not observed. Furthermore, Mus81-Mms4/Slx1-Slx4-mediated cleavage were required for repositioning. Together, this implicates a post-replicative DSB as the signal for relocation of an (AT)34 repeat. A commonality of all examples of fork barriers that relocate is the presence of exposed ssDNA. Single-stranded DNA provides a template for proteins to bind and be SUMOylated to trigger recruitment to the NP, and also serves as a signal to initiate the DNA damage checkpoint.

In addition to RDR and DSB repair pathways, is gap filling taking place at the nuclear periphery? A gap is one possible outcome of an uncoupled fork. DNA structures such as the hairpin-forming expanded CAG/CTG tracts can stall and reverse forks, but also cause difficult to repair gaps (reviewed in (Polleys and Freudenreich, 2021); see also (Polleys et al., 2023)). Additionally, DSB-capturing nuclear envelope tubules have been shown to be recruited by treatment of BRCA1-deficient cells with PARP inhibitors (Shokrollahi et al., 2024), which have been shown to induce formation of ssDNA gaps (Cong and Cantor, 2022). However, other types of damage that produce ssDNA gaps, such as MMS damage (Krokan and Bjoras, 2013) or HU stalled forks (Ercilla et al., 2019) do not cause a shift to the NPC. The Ulrich laboratory specifically tested the location of MMS-induced damage bypass events in the nucleus by tracking RPA foci, and found they occur at post-replicative repair territories (PORTs) in the nuclear interior (Wong et al., 2020). The key differences that drive one type of lesion to shift to the nuclear periphery while the other remains in the nuclear interior, even though both accumulate ssDNA, remain to be elucidated.

Connections and remaining questions

One outstanding question is the relationship between nuclear filaments, movement to the NP, and fork restart. In humans, myosin VI recruits WRNIP to protect stalled forks from DNA2-mediated degradation, requiring the ubiquitin binding domain of myosin VI (Shi et al., 2023). It has been previously shown that WRN and DNA2 cooperate to facilitate fork restart after fork reversal (Thangavel et al., 2015). This suggests a relationship between movement along actin filaments and fork restart. Additionally, F-actin filaments have been shown to restrain PrimPol activity (Palumbieri et al., 2023) and heterochromatin markers appear at sites of persistent stalling and serve to antagonize PrimPol recruitment (Gaggioli et al., 2023). Together, these works suggest that F-actin association, and potentially mobilization of damaged loci within the nucleus, are regulatory mechanisms for protecting and restarting perturbed replication.

There is an interesting connection between recruitment of repair factors to phase-separated domains, and mobilization of damaged loci (Chiolo et al., 2025; Spegg et al., 2023) SUMOylation drives formation of phase-separated domains in response to stress (Cheng, 2023), and repositioning of expanded CAG/CTG repeats to the nuclear pore complex requires SUMOylation of repair proteins, such as RPA and Rad52 (Whalen et al., 2020). Additionally, DIMs that form in budding yeast have been shown to serve as nucleation sites for Rad52 droplet formation (Oshidari et al., 2020), and expanded CAG/CTG repeats associate with both Rad52 and DIMs prior to relocation (Maclay et al., 2025; Su et al., 2015; Whalen et al., 2020). Formation of a phase separated domain could be facilitating the relocation event and could also be influencing the repair pathway. Indeed, in Drosophila, association of heterochromatic DSBs with the nucleoporin Nup98 creates a condensate that excludes Rad51(Merigliano et al., 2025).

Another question is what types of protein modifications or turnover processes occur at the NPC to facilitate recovery from replication stress? In the best understood case to date, it was shown that polySUMOylation of repair proteins at the RTS1-RFB stalled fork prevents fork restart by RDR, and these SUMO chains must be removed by the NPC-associated Ulp1 SENP protease in order for RDR to initiate (Kramarz et al., 2020; Schirmeisen et al., 2024). Interestingly, the proteasome is required for processivity of the restarted fork (but not initiation of the process), implicating a different role for the ubiquitin pathway (Schirmeisen et al., 2024). The regulation of the repair processes that occur at the NPC is not as well understood for the structure-induced replication barriers, or for that matter any of the other types of DNA damage that relocate. Differences in the signal between RFBs and repeat stalled replication forks such as SUMO requirements, resection nuclease requirements and the role of Rad51 hint that there are multiple repair or fork restart pathways that may be facilitated by the peripheral location.

Spatial relocation to a different nuclear compartment clearly plays a genome protective role, but the specific repair processes that occur at the nuclear periphery or at the nuclear pore complex remain to be fully elucidated. While there are many substrates that reposition to the nuclear periphery, they have distinct requirements for doing so (Table 1, Figure 2). These different requirements for different lesions suggest distinct pathways for repair at the nuclear periphery exist, and the specific characteristics of these pathways remain open questions.

In summary, repositioning to the nuclear periphery is an important component of recovery from some types of DNA replication stress. This pathway appears to exist, with minor differences, in many organisms across the tree of life. This suggests that it has been an effective and therefore conserved strategy for the response to hard to repair DNA lesions, including those resulting from replication barriers and persistent DSBs. Not all DNA lesions shift to the nuclear periphery, but those that do present unique challenges for efficient fork restart or repair. The repositioning requires not only processing at the site of damage but also a number of post-translational modifications of proteins that bind to resected DNA, suggesting that this spatial shift is a highly regulated and organized process. There are, however, still significant gaps in our understanding of the underlying molecular cues, modified protein targets, and functional outcomes. In particular, the suite of repair or fork recovery pathways that occur at the periphery is not yet fully known, nor the balance between error-free repair and error-prone pathways. It may be that the main purpose of relocation to the NPC is to promote viability in response to a severe genomic insult. This highlights the need for further research into this mechanism to answer these important questions.

Acknowledgements

The authors would like to thank Chiara Masnovo for her thoughtful comments on this manuscript.

Funding Statement

Institute of General Medical Sciences of the National Institutes of Health under Award Number R35 GM144215 to CHF. Beckman Scholars Grant to NC.

Footnotes

Declaration of Competing Interest

The authors declare no conflicting interests

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