Genetic Regulation of DNA Double-Strand Breaks and Repair Pathways

Abstract

DNA double-strand breaks (DSB) represent one of the most severe forms of genomic damage. Thus, cells have evolved a complex network of DSB repair pathways, including homologous recombination, classical and alternative end joining, and single-strand annealing, which are tightly regulated by genetic and epigenetic factors. The selection and efficiency of these pathways influence genome integrity, oncogenesis, and therapeutic response. This comprehensive review synthesizes recent findings on the genetic regulation of DSB repair, with emphasis on pathway-specific regulators, chromatin context, and post-translational modifications. Moreover, this review integrates primary research from mammalian systems, including CRISPR-based studies, proteomics, and imaging, with a focus on publications from 2020 to 2025. We discuss the role of key players, such as MRE11–RAD50–NBS1 (MRN), ataxia telangiectasia mutated (ATM), mediator tumor suppressor p53-binding protein 1 (53BP1), breast cancer type 1 susceptibility protein (BRCA1), anti-silencing function 1 (ASF1), ring finger protein (RNF)8/168, DNA-dependent protein kinase catalytic subunit (DNA-PKcs), and RAD51 recombinase (RAD51), in orchestrating the associated pathway choice. Epigenetic modifications, RNA-mediated mechanisms, and chromatin remodeling dynamically influence the efficiency and fidelity of repair. Particular attention is provided to emerging regulators, including thyroid hormone receptor interactor 13 (TRIP13), ubiquitin-like with plant homeodomain (PHD) and RING finger domains 1 (UHRF1), Shieldin, and polymerase theta. This review highlights novel insights into transcription-associated DSB repair, the interplay of replication stress with repair pathway engagement, and context-dependent synthetic lethality. We also examine implications for cancer biology, including therapy resistance and biomarker development. Ultimately, understanding the genetic regulation of DSB repair pathways can provide critical insights into genome stability maintenance and reveal new therapeutic opportunities in cancer. Future work should focus on pathway crosstalk, phase-specific regulation, and integrating repair modulation into personalized medicine.

Graphical abstract

Keywords

double-strand breaks / DNA repair / homologous recombination / nonhomologous end-joining / chromatin remodeling / ubiquitination / RNA-DNA hybrid / neoplasms

1. Introduction

Among all DNA lesions, double-strand breaks (DSB) pose the highest threat to genome integrity. Eukaryotic cells rely mainly on two major routes for their resolution: the high-fidelity homologous recombination (HR) and the versatile, ligation-driven classical non-homologous end joining (C-NHEJ) (Fig. 1) [1]. HR is a high-fidelity repair mechanism active during the S and G2 phases of the cell cycle, when a sister chromatid is available as a repair template. In contrast, C-NHEJ functions throughout the cell cycle and directly ligates DNA-ends without the need for homology. More recently, an additional error-prone pathway — alternative end joining (Alt-EJ), also referred to as microhomology-mediated end joining (MMEJ) has been characterized (Fig. 1). This pathway utilizes short microhomologous sequences (typically 5–25 base pairs) flanking the break site to mediate repair [2]. The decision between these pathways is tightly regulated and influenced by multiple factors, including chromatin state at the break locus [3], the extent of 5^′ end resection, and the phase of the cell cycle [1]. In addition, numerous proteins act as key regulators either promoting or inhibiting specific repair routes. The review focuses on the molecular mechanisms underlying pathway choice in DSB repair, emphasizing the roles of DNA-end processing factors, chromatin modifiers, and regulatory proteins that determine the engagement of HR, C-NHEJ, or MMEJ/Alt-EJ.

This review was prepared as a narrative synthesis of peer-reviewed research articles published between 2020 and 2025, identified through comprehensive searches of the PubMed and Scopus databases using defined keywords combinations. Approximately 75% of all cited references originate from this five-year period, ensuring that the review reflects the most recent and authoritative advances in the field. Priority was given to primary research studies in mammalian systems providing mechanistic insights confirmed by biochemical, imaging, or CRISPR-based approaches. Earlier seminal works were included selectively to provide historical or conceptual context.

2. DSB Repair Pathway Selection and Regulation — Factors Involved in DNA Damage Response

While the molecular components of DSB repair have been extensively characterized, the mechanistic hierarchy governing pathway choice remains under active investigation. Emerging evidence suggests that multiple, sometimes conflicting, regulatory layers act simultaneously, underscoring the need for integrated models of DSB repair regulation (Table 1, Ref. [4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47, 48, 49, 50, 51, 52, 53, 54, 55, 56, 57, 58, 59, 60, 61, 62, 63, 64, 65, 66, 67, 68, 69, 70, 71, 72, 73, 74, 75, 76, 77, 78, 79, 80, 81, 82, 83, 84, 85, 86, 87, 88, 89, 90, 91]) [92].

Upon DSB detection, the sensor complex MRE11-RAD50-NBS1 (MRN) recruits and activates the kinase ATM (ataxia-telangiectasia mutated), which initiates a phosphorylation cascade central to the DNA damage response (DDR) [93]. Activated ATM undergoes autophosphorylation and dimer dissociation, enabling phosphorylation of downstream effectors such as breast cancer type 1 susceptibility protein (BRCA1), checkpoint kinase 2 (CHK2), tumor suppressor protein p53 (p53), and the histone variant H2AX, producing the phosphorylated H2AX (\gammaH2AX) mark that scaffolds assembly of repair proteins [94, 95].

Mediator of DNA damage checkpoint protein 1 (MDC1) is subsequently recruited to \gammaH2AX sites to reinforce signal amplification and promote accumulation of repair factors. The ubiquitin ligases ring finger protein (RNF)8 and RNF168 extend this signal by catalyzing K63-linked polyubiquitin chains on histone H1 and H2A [96], providing binding platforms for the mediator p53-binding protein 1 (53BP1), which limits excessive resection of DNA ends and favors C-NHEJ [97, 98, 99]. 53BP1 executes this protective function in cooperation with the Shieldin complex and Rap1-interacting factor 1 (RIF1), thereby preventing nucleolytic degradation and facilitating repair through end joining. Conversely, recruitment of BRCA1 to DSB together with BRCA1-associated RING domain 1 (BARD1) and CtBP-interacting protein (CtIP) promotes resection and drives the shift toward HR [100, 101]. Notably, the Bloom syndrome helicase (BLM) has been shown to function in both HR and C-NHEJ pathways, participating in the recruitment of repair factors during S and G1 phases, respectively, and thus influencing pathway choice based on the cell cycle context [102].

2.1 Classical Non-Homologous End-Joining and Proteins Promoting C-NHEJ Repair

In higher eukaryotic organisms, the C-NHEJ pathway serves as the predominant mechanism for repairing DSB, responsible for correcting approximately 80% of such lesions. The repair process initiates with the Ku70/Ku80 heterodimer, which rapidly associates with the DNA termini, safeguarding them from resection and serving as a scaffold for subsequent recruitment of repair factors (Fig. 1) [4]. DNA-dependent protein kinase (DNA-PK) catalytic subunit (DNA-PKcs) associates with Ku to form the DNA-PK holoenzyme that bridges broken DNA ends and coordinates end-processing enzymes. Upon activation, DNA-PKcs undergoes autophosphorylation and phosphorylates several downstream effectors, including Artemis, polynucleotide kinase phosphatase (PNKP), X-ray repair cross-complementing protein 4 (XRCC4), XRCC4-like factor (XLF), and DNA ligase IV (LIG4), which collectively execute end processing and ligation. The cooperation among these enzymes ensures restoration of DNA continuity while maintaining genome stability [5]. The nucleolytic activities of Artemis, exonuclease 1 (EXO1), tyrosyl-DNA phosphodiesterase 1 (Tdp1), and PNKP prepare DNA-ends, while polymerases from the Pol X family (such as polymerase µ and \lambda) participate in gap-filling synthesis [5]. The final ligation is executed by the XRCC4/XLF/DNA ligase IV complex [6].

Recent cryo-electron microscopy (cryo-EM) structural studies have elucidated the direct interaction interfaces between Ku70/Ku80 and DNA polymerases \lambda and µ (Pol \lambda and Pol µ), detailing the role of their BRCA1 C-terminal (BRCT) domains in mediating efficient gap-filling and C-NHEJ fidelity [7]. It has been recently revealed that glioblastoma stem-like cells (GSCs) bolster their DNA repair proficiency by activating the C-NHEJ pathway through the upregulation of apoptosis antagonizing transcription factor (AATF), thereby contributing to treatment resistance in glioblastoma (GB). AATF directly binds to XRCC4, a critical component of the C-NHEJ machinery, and protects it from ubiquitin-dependent proteasomal degradation. Upon induction of DNA damage, AATF is phosphorylated at serine 189 by ATM kinase, which facilitates its release from XRCC4 and enables the prompt recruitment of XRCC4 to DNA break sites, ensuring efficient NHEJ-mediated repair. Notably, silencing AATF or disrupting its phosphorylation impairs C-NHEJ efficiency in GSCs and sensitizes GB xenograft models to chemoradiotherapy [8].

A pivotal factor influencing DSB repair pathway choice is the phosphorylation state of DNA-PKcs [9]. As a central orchestrator of the C-NHEJ process, DNA-PKcs phosphorylates itself and other C-NHEJ-related proteins (Ku, XRCC4, XLF, Artemis), promoting repair fidelity and completion [10]. During G1 phase, DNA-PKcs inhibits ATM signaling through phosphorylation of ATM, thereby favoring C-NHEJ [11]. In contrast, during G2 phase, autophosphorylation of DNA-PKcs leads to its release from DNA-ends, permitting EXO1-mediated resection and subsequent HR activation [12]. During the S phase, the association of BRCA1 with DNA-PKcs counteracts its kinase self-activation, thereby diminishing end-joining efficiency while promoting DNA-end resection and facilitating the transition toward HR [13].

Also, DNA-PKcs exerts indirect control over pathway choice via modulation of RBX1 expression, which targets EXO1 for degradation during G1 phase, suppressing HR [14]. Furthermore, recent reviews underscore the multifaceted role of DNA-PKcs beyond DNA repair, including its regulation of RNA metabolism, transcription machinery interactions, alternative splicing, and noncoding RNA functions in cancer biology [15].

The Ku70/Ku80 heterodimer also plays a crucial role in shielding DNA-ends from nucleolytic degradation, preserving the structural integrity of DSB and enforcing C-NHEJ fidelity [16]. Additionally, the C-terminal SAP domain of Ku70 has been shown to be essential for optimal cellular responses to DNA damage, promoting Ku70 retention at DSB and enhancing DNA ligase IV stability at these sites [17]. Another central regulator is 53BP1, which limits end resection, thereby antagonizing BRCA1-mediated HR and reinforcing C-NHEJ preference [18]. 53BP1 promotes end joining via two primary routes: the replication timing regulatory factor 1 (RIF1)-REV7 (also known as mitotic arrest-deficient protein 2, MAD2L2)-REV7/Shieldin complex and the PTIP-Artemis pathway [19]. Nevertheless, 53BP1 has additional roles in HR under replication stress, including both BLM helicase-dependent and -independent activities [20], and it suppresses HR in BRCA1-deficient contexts by blocking end resection [21]. Importantly, post-translational acetylation of 53BP1 impedes its chromatin recruitment, thus favoring HR [22]. Phosphorylation of 53BP1 by ATM at its N-terminal domain allows for the recruitment of RIF1 to damaged DNA, a process that inhibits end resection during G1 phase. In contrast, during S/G2 phases, CDK-mediated phosphorylation of CtIP activates the BRCA1-CtIP complex, suppressing RIF1 recruitment and thereby enabling HR progression [23].

Acting downstream of RIF1, the Shieldin complex (comprising SHLD1, SHLD2, SHLD3, and REV7) protecs DSB termini from extensive resection [11, 24, 99]. The SHLD2 subunit possesses an RPA70-like oligonucleotide/oligosaccharide-binding fold that anchors to single-stranded DNA, thereby preventing resection nucleases such as EXO1 and DNA2 from accessing DNA ends [24]. Dynein light chain LC8-type 1 (DYNLL1), a 53BP1-interacting protein, further reinforces this blockade by associating with MRE11 - its depletion enhances Shieldin accumulation at break sites [25]. REV7, a vital Shieldin component, suppresses 5^′–3^′ end resection and bolsters C-NHEJ by dimerizing and coordinating interactions with SHLD1 and SHLD3. These combined events recrute the CST complex (CTC1-STN1-TEN1) together with DNA polymerase \alpha, which mediates limited fill-in synthesis to prepare DNA ends for ligation via the C-NHEJ pathway [26]. Notably, even in Shieldin-deficient BRCA1-null cells, an alternative fill-in pathway mediated by CST/Pol\alpha/primase complex can partially substitute the protective function [27]. When REV7 is inactivated, SHLD3 fails to localize to DSBs, allowing BRCA1 to displace 53BP1 and initiate extensive resection, thereby enabling HR [28]. Both RIF1 and PTIP are independently recruited via distinct phosphorylation motifs on 53BP1 and are indispensable for C-NHEJ commitment [29, 30]. PTIP specifically facilitates the recruitment of Artemis, which, via its endonuclease activity, processes overhangs at DSB to render them ligatable, completing the repair through the C-NHEJ pathway [31].

2.2 Homologous Recombination and Proteins Promoting HR Repair

HR is a high-fidelity DNA repair mechanism that restores genomic integrity by exchanging homologous or identical DNA sequences between a damaged strand and an intact homologous template. The process is initiated by the MRN complex (comprising MRE11, RAD50, and NBS1) which detects the break and promotes 5^′ to 3^′ resection at the DSB-ends, generating 3^′ single-stranded DNA (ssDNA) overhangs (Fig. 1). These overhangs are initially coated by replication protein A (RPA), which stabilizes ssDNA and prevents secondary structure formation. RPA is subsequently displaced by the BRCA2-mediated loading of RAD51, leading to the formation of a presynaptic nucleoprotein filament [93]. This filament performs a homology search and invades a homologous duplex DNA, forming a displacement loop (D-loop) that serves as a primer for DNA synthesis.

Following strand invasion and extension, the repair process can proceed through one of three distinct sub-pathways [94]: (a) DSB repair (DSBR), which involves second-end capture and the formation of a double Holliday junction, later resolved by structure-specific endonucleases such as MUS81-EME1, SLX1-SLX4, or flap endonuclease GEN homolog 1 (GEN1), leading to gene conversion events with or without crossover; (b) Synthesis-dependent strand annealing (SDSA), where the extended invading strand dissociates and anneals to the resected second end without forming a Holliday junction, generally resulting in non-crossover outcomes; (c) Break-induced replication (BIR), a less common but highly mutagenic pathway wherein the D-loop is converted into a replication fork, facilitating extensive DNA synthesis to the end of the chromosome and often causing loss of heterozygosity.

RAD54 plays dual roles in HR and chromatin remodeling [32]. It stabilizes the RAD51-ssDNA presynaptic filament, enhancing its strand invasion activity during HR [32]. Beyond the synaptic phase, RAD54 facilitates RAD51 turnover by dissociating it from heteroduplex DNA, thus exerting a postsynaptic regulatory function in vivo [1]. Additionally, RAD54 accelerates joint molecule formation and promotes ATP-dependent branch migration during later stages of HR [1, 32]. Notably, BLM helicase has been shown to augment RAD54-driven chromatin remodeling, contributing to chemotherapy resistance [33]. RAD51, a core HR recombinase, suppresses non-conservative pathways such as SSA and Alt-EJ, while favoring gene conversion (GC). This preference is largely dictated by its loading efficiency onto ssDNA [34].

BRCA1 functions as a central coordinator of HR, engaging at several distinct checkpoints of the repair cascade. Upon the formation of a DSB, BRCA1 associates with CtIP and the MRN complex to initiate limited DNA-end resection, generating stretches of ssDNA that enable RAD51 loading while counteracting 53BP1-mediated end protection. This interplay redirects the balance from end joining toward recombination-based repair [35]. Furthermore, the BRCA1-BARD1 heterodimer enhances the homologous pairing activity of RAD51, thereby supporting recombination fidelity [35, 36].

The human RecQ helicases BLM and WRN also critically influence pathway selection. BLM participates in both DNA end resection and Holliday junction resolution during HR [37, 38], and can stimulate EXO1-mediated resection to promote HR initiation [39]. Additionally, BLM coordinates repair factor recruitment in a cell cycle-dependent manner: in G1, it facilitates XRCC4 loading to support C-NHEJ, while in S phase, it aids RAD51 recruitment to promote HR [102].

WRN functions similarly, participating in end resection via interactions with MRE11, NBS1, EXO1, and DNA2 [40]. Its activity is modulated by posttranslational modifications, including CDK2-dependent phosphorylation at Ser426, which enhances its binding to RPA and stabilizes 3^′ ssDNA intermediates [41]. In early HR stages, WRN also promotes RAD51 loading through CHK1 signaling [42]. Recent work shows that the WRN-RPA complex is crucial for replication fork recovery following replication stress, thereby limiting DSB formation and indirectly supporting HR fidelity [43]. Another important regulator of repair pathway choice is TRIP13 (also known as HPV16E1BP), an ATP-ase that inactivates REV7 by inducing a conformational switch. This process disrupts the REV7-Shieldin complex and promotes HR [44].

UHRF1 (Ubiquitin-like with PHD and RING finger domains 1) is an epigenetic modulator that contributes to transcriptional repression via HDAC1 induction [45] and also plays a direct role in DSB repair. Loss of UHRF1 leads to defective DSB resolution, heightened sensitivity to genotoxic stress, chromosomal instability, and increased sister chromatid exchange (SCE) [46]. Phosphorylated UHRF1 promotes HR through its E3 ligase activity at break sites [46, 47], and its methylated form binds PCNA to enhance its polyubiquitination, further stimulating HR [46]. Furthermore, DNMT1 inhibition triggers UHRF1-dependent H3K18ub and concomitant H3K9me3 increases, indicating a chromatin-mediated layer of HR regulation via UHRF1 as shown in 2025 study [48]. Importantly, UHRF1-deficient cells exhibit hypersensitivity to PARP inhibitors, suggesting a pivotal role in DSB repair downstream of BRCA1. Recent pancancer analysis confirms high HR deficiency rates associated with mutations in RAD54L, RAD51C/D, and PALB2, underscoring their functional significance in HR and therapeutic vulnerability [49].

2.3 Microhomology-Mediated End Joining/Alternative End Joining and Its Regulation

The recently identified MMEJ/Alt-EJ, constitutes an error-prone mechanism for DSB repair that can function independently of C-NHEJ [103]. MMEJ/Alt-EJ operates even in cells with intact C-NHEJ components, provided that microhomologous regions (typically 5 nucleotides or longer) are available near the DNA break sites (Fig. 1) [50]. These short homologies are recognized and utilized by the MRN complex in conjunction with PARP1 or potentially other yet-unidentified factors. Following initial recognition, the MRN complex resects DNA ends until complementary microhomologies are exposed on either side of the break. This facilitates strand annealing, while endonucleases such as xeroderma pigmentosum complementation group F–excision repair cross-complementation group 1 complex (XPF)/ERCC1 or flap endonuclease 1 (FEN1) remove non-homologous 3^′ flaps [51]. After reannealing of microhomologous sequences, Pol\theta performs limited DNA synthesis to bridge the remaining gaps, followed by end sealing carried out by the XRCC1-DNA ligase III complex, which restores DNA continuity [52]. As MMEJ/Alt-EJ typically deletes sequences between the microhomology regions, it is highly mutagenic and therefore must be tightly regulated during DSB repair pathway selection [53].

CtIP- and MRE11-driven DNA-end resection initiates the MMEJ/Alt-EJ pathway by revealing microhomologous regions. Notably, depletion of BLM or EXO1 disrupts this process and shifts repair pathway utilization [54, 103]. BRCA1, a central player in HR, acts to suppress MMEJ/Alt-EJ by promoting extended end resection and RAD51-mediated strand invasion [54]. Similarly, replication protein A (RPA) antagonizes MMEJ/Alt-EJ by binding to ssDNA and preventing spontaneous annealing of short homologous sequences close to the break [55].

Further, loss of the tumor suppressor RB1 has been associated with elevated MMEJ/Alt-EJ activity, pointing to its role as another regulatory factor influencing DSB repair pathway choice [56]. Conversely, DNA polymerase theta (Pol\theta), which possesses both polymerase and helicase activity, is a core effector of MMEJ/Alt-EJ and promotes end joining at sites with short microhomology [57, 58]. In HR-deficient tumors, elevated POL\theta expression has been linked to increased reliance on MMEJ/Alt-EJ, suggesting that Pol\theta might serve as a potential therapeutic target in cancers with defective HR machinery, reinforced by recent preclinical data showing that POL\theta inhibition effectively sensitizes HR-deficient cells to DNA damage [59].

2.4 Single-Strand Annealing

Another repair mechanism for DSB, in addition to HR and end-joining pathways, is the error-prone single-strand annealing (SSA) pathway (Fig. 1). Unlike HR, SSA does not require a homologous donor template but instead utilizes direct tandem repeat sequences that flank the break site [60]. Following the generation of DSB, the MRN complex initiates DNA end resection via the exonuclease activity of MRE11, similarly to the HR pathway. However, SSA typically involves more extensive resection than MMEJ/Alt-EJ, requiring long ssDNA regions generated by CtIP and EXO1 to expose homologous repeats on both sides of the break. This mechanism proceeds independently of RAD51 but requires RAD52 to mediate strand annealing between the complementary repeat sequences. Once annealing occurs, ERCC1/XPF endonuclease removes non-homologous 3^′-ssDNA overhangs that result from the resection process [60]. Subsequently, DNA polymerases fill in the remaining gaps, and the DNA ends are ligated by either DNA ligase I or ligase III to restore the strand continuity. However, SSA inevitably results in the deletion of one repeat and the intervening sequence between the repeats, rendering the pathway inherently mutagenic.

Previous studies have proposed that SSA pathway might contribute to precise donor integration by facilitating the annealing of homology arms in the donor construct to complementary genomic sequences [61, 62, 63]. However, to date, no direct experimental evidence has substantiated this hypothesis. Findings from Tei et al. (2025) [64] demonstrate that inhibition of the SSA pathway did not diminish the frequency of precise homology-directed repair (HDR), but rather led to a reduction in asymmetric HDR events. These results imply that SSA may mediate accurate integration at one end of the donor construct, while the opposite end undergoes imprecise resolution via alternative repair mechanisms, ultimately resulting in asymmetrical integration. Therefore, targeted suppression of SSA could reduce erroneous integration events and represents a promising strategy to enhance the efficiency of precise knock-in.

2.5 Outstanding Questions and Future Directions in Canonical DSB Repair

Despite major advances in defining the molecular architecture of canonical DSB repair pathways, several key regulatory principles remain unresolved. One major gap concerns how cells dynamically reconcile competing repair cues from 53BP1-RIF1-Shieldin complexes and BRCA1-CtIP-MRN signaling, particularly during S and G2 phases when both HR and C-NHEJ machineries are accessible. It is not fully understood how post-translational modifications (PTMs) of shared factors (e.g., phosphorylation, acetylation, or SUMOylation of 53BP1, RIF1, or BRCA1) bias repair toward one pathway in different chromatin contexts. Further biochemical and single-molecule studies are needed to clarify whether these modifications act as transient molecular “toggles” or as stable determinants of pathway choice.

Likewise, several recently identified proteins such as AATF and TRIP13 play dual or context-dependent roles, but their in vivo relevance remains only partially defined. AATF has been shown to stabilize XRCC4 during C-NHEJ and confer therapeutic resistance in glioblastoma models, yet whether its phosphorylation by ATM also influences HR efficiency or replication fork recovery is still unclear. TRIP13-mediated conformational regulation of REV7 has emerged as a central switch between Shieldin-dependent C-NHEJ and HR, but the temporal hierarchy and cell-cycle specificity of this process are incompletely mapped. Further in vivo models and time-resolved interactome analyses will be required to delineate these transitions.

Another conceptual uncertainty involves histone modifications. While certain chromatin marks such as H2AK15ub, H4K20me1/2, and H3K79me2 are firmly established as C-NHEJ-promoting features, others (including H3K9me2/3 or H3K36me2) show more correlative associations with pathway bias and remain under debate. Similarly, the histone chaperone ASF1 has been implicated in promoting both HR and C-NHEJ. The determinants of this bifunctionality are not yet known. It is possible that ASF1’s binding partners (RIF1 versus MMS22L-TONSL) or chromatin accessibility around DSB dictate its functional polarity, a hypothesis requiring targeted biochemical validation.

Collectively, these open questions emphasize that DSB repair pathway selection is not governed by linear hierarchies but by highly dynamic, context-dependent signaling networks. Integrating spatial chromatin organization, local transcriptional activity, and real-time post-translational regulation into systems-level models represents a major challenge for the next decade of DSB repair research.

3. Transcription-Coupled DNA Repair and Genomic Instability

Transcription plays a multifaceted role in genome maintenance, exerting both protective and detrimental effects on genomic stability. On the one hand, transcription enhances genome integrity by facilitating the recruitment of lesion-specific DNA repair proteins to transcribed regions, thereby ensuring efficient surveillance and repair of transcriptionally active loci. On the other hand, transcription can contribute to genomic instability under certain conditions, such as when it becomes dysregulated, when transcription machinery encounters replication forks leading to conflicts, or when aberrant formation of (RNA-DNA loops) R-loops occurs. These scenarios can impair genome maintenance and promote mutagenic outcomes.

3.1 Transcription-Coupled Nucleotide Excision Repair

Genotoxic insults such as UV irradiation induce bulky DNA lesions, which are resolved through two major nucleotide excision repair pathways: global genome NER (GG-NER) and transcription-coupled NER (TC-NER). These pathways differ in the initial lesion recognition steps. GG-NER relies on the detection of DNA damage by two distinct protein complexes: UV-DDB (comprising DDB1 and DDB2) [104] and the XPC-RAD23B-CETN2 complex [105, 106, 107]. In contrast, TC-NER identifies lesions via the stalling of RNA polymerase II (Pol II) at sites of transcription-blocking damage. Despite these different lesion recognition mechanisms, both pathways converge on a shared downstream set of repair proteins, including transcription factor IIH (TFIIH), xeroderma pigmentosum group A protein (XPA), xeroderma pigmentosum group G endonuclease (XPG), RPA, ERCC1-XPF, DNA polymerases, and ligases. A key player in TC-NER is cockayne syndrome protein B (CSB), which is recruited to stalled Pol II, and in turn facilitates the assembly of CSA and UV-stimulated scaffold protein A (UVSSA), initiating the repair cascade by targeting TFIIH to the lesion, halting transcription, and mediating the removal of the stalled Pol II complex [108, 109].

3.2 Transcription-Coupled Homologous Recombination Repair

Although TC-NER has been recognized since the 1980s, only recently has transcription-coupled repair of DNA DSB been described. Chaurasia et al. (2012) [110] reported that DSB in actively transcribed genes are resolved more efficiently than those in transcriptionally silent regions [111]. DSB repair generally occurs via two major pathways: C-NHEJ and HR. C-NHEJ is a rapid but error-prone mechanism that simply ligates broken DNA-ends, whereas HR is a more accurate process requiring a homologous sequence - typically the sister chromatid as a template for repair [112]. The availability of sister chromatids during specific cell cycle phases heavily influences the choice between these two pathways. Furthermore, several studies suggest that HR is preferentially used at sites of active transcription, possibly due to chromatin openness or associated structural features [113, 114]. Factors such as RAD52 and RAD51AP1 have been implicated in facilitating HR specifically at transcribed loci [115].

3.3 RNA Transcription DNA Repair

The preference for HR repair at actively transcribed regions may stem from various factors including open chromatin, specific epigenetic marks, the presence of transcriptional machinery, or the RNA transcripts themselves [116, 117, 118]. Using an inducible DSB model in which transcription can be activated by doxycycline, it was demonstrated that active transcription enhances HR efficiency [119]. Notably, even a tethered RNA transcript homologous to the repair template was sufficient to mimic this effect, indicating a direct functional role of RNA in transcription-coupled HR [119]. This RNA-mediated stimulation of HR depends on sequence homology, suggesting that RNA-DNA hybrid structures form during repair. Several studies in both yeast and mammalian cells have confirmed the presence of RNA-DNA hybrids at DSB sites. The RNA component of these hybrids may originate from various sources including pre-existing transcripts or RNA newly synthesized by Pol II or Pol III [120, 121]. However, genome-wide analyses indicate that pre-existing RNA is the predominant contributor to the RNA-DNA hybrids observed at break sites [122, 123]. In special conditions, such as in yeast lacking both RNase H and reverse transcriptase, RNA can even serve as a direct template for DSB repair [124]. Similarly, in human G0/G1-phase cells where sister chromatids are absent, ROS-induced DSB can be repaired in a manner dependent on RNA-DNA hybrids, implying that RNA templates may also play a role [125]. It remains to be determined whether such repair in G0/G1 represents a form of non-canonical, RNA-templated HR.

3.4 Transcription as a Source of Genomic Instability

Transcription and its associated RNA products can represent significant endogenous sources of genome instability, particularly when transcriptional activity interferes with replication processes or facilitates the formation of aberrant R-loop structures. While pathological R-loops are recognized threats to genome stability, physiological R-loops fulfill critical regulatory roles, including in transcriptional initiation and termination, immunoglobulin class switch recombination, and the maintenance of telomeric and centromeric integrity [126, 127, 128]. Concurrent activity of transcription and replication on the same DNA template can lead to physical encounters between the transcription machinery and the replication fork, resulting in either head-on or co-directional collisions. These events may compromise genome integrity by generating DNA breaks. Much of the foundational understanding of transcription-replication collisions (TRCs) originates from prokaryotic systems, which possess a single replication origin. In E. coli, most highly expressed operons are oriented co-directionally with replication to minimize head-on conflicts, a likely outcome of evolutionary pressure. In contrast, head-on TRCs, which are more mutagenic, may contribute to rapid genomic evolution [129]. A recent study emphasizes the complex role of nascent RNA at transcription-replication conflict (TRC) sites, where the emerging transcript can critically influence conflict outcomes [130].

Reviews now increasingly consider chromatin context and DNA damage signaling in TRC regulation as potential therapeutic targets [131]. In mammalian systems, where thousands of replication origins are activated during each cell cycle [132], and where transcription is tightly regulated in a spatially and temporally complex manner, TRCs are more challenging to characterize. Utilizing an episomal model containing an R-loop-forming sequence and an inducible promoter system, Hamperl et al. (2017) [133] demonstrated that both head-on and co-directional TRCs provoke DNA damage. Notably, these collision orientations elicit distinct types of DNA lesions that are resolved through different DNA repair pathways [134]. Moreover, interference at TRCs has been shown to drive R-loop-dependent DNA damage responses in mammalian cells, corroborating the mechanistically detrimental role of R-loops in genomic instability [135]. Noteworthy, Hartono et al. (2018) [136] provided a critical methodological refinement, revealing that the widely used S9.6 antibody exhibits high affinity not only for DNA-RNA hybrids but also for double-stranded RNA. This discovery demonstrated that previous genome-wide R-loop maps may contain false-positive signals, emphasizing the need for orthogonal validation methods and improved specificity in R-loop detection.

Transcriptional dysregulation is a frequent source of pathological R-loop formation. Oncogene activation, such as by ewing sarcoma RNA-binding protein fused with Friend leukemia integration 1 transcription factor (EWS-FLI1) or harvey rat sarcoma viral oncogene homolog with glycine-to-valine substitution at codon 12 (HRASG12V), has been shown to enhance R-loop accumulation at target genes regulated by these factors [137, 138]. This phenomenon is thought to arise from an overabundance of RNA transcripts available for hybridization with the template DNA. Interestingly, reduced transcriptional output due to RNA polymerase II (Pol II) pausing can also increase R-loop formation, likely due to prolonged retention of nascent transcripts within the transcription bubble [139, 140]. R-loop formation is tightly coupled to co-transcriptional RNA processing, including splicing, editing, nuclear export, and RNA-binding protein interactions. Impairment in these processes can induce transcriptional stress and facilitate aberrant R-loop accumulation, thereby linking RNA metabolism to transcription-associated genomic instability [141, 142]. Interestingly, in Leishmania major, replication timing and chromosome stability correlate with formation of R-loops and RNase H1 distribution, highlighting how R-loop resolution factors interface with chromatin accessibility and replication programming [143].

To maintain genome integrity, pathological R-loops must be efficiently resolved. The RNA moiety of an R-loop can be specifically degraded by RNase H enzymes. Alternatively, RNA-DNA helicases such as SETX, aquarius intron-binding spliceosomal factor (AQR), DEAD-box helicase 5 (DDX5), and DEAH-box helicase 9 (DHX9) can unwind the hybrid structure. Additionally, nucleases including CtIP, FEN1, XPF, and XPG are implicated in excising R-loops formed during transcription [144, 145], whereas enzymes like MUS81 and MRE11 resolve DNA structures arising from R-loop-replication fork collisions [146, 147]. Furthermore, DNA repair proteins such as BRCA1, BRCA2, and the FANCD2-FANCI complex have been shown to suppress R-loop formation in diverse biological contexts [148, 149]. Errors introduced during transcription, defined as sequence mismatches between nascent RNA transcripts and the DNA template, may stem from the inherent infidelity of RNA polymerases or be triggered by DNA damage and unfavorable chromatin environments. Although typically transient, these errors can exert meaningful effects on genome stability, especially under specific conditions or in repetitive genomic contexts [150, 151].

3.5 Outstanding Questions and Future Directions in Transcription-Coupled DNA Repair

Although the relationship between transcription and genome maintenance has been intensively studied, the mechanistic and regulatory landscape of transcription-coupled DNA repair (TCR) remains only partially understood. One unresolved issue concerns how cells prioritize between RNA polymerase removal, lesion bypass, and DSB repair at sites of transcriptional blockage. The temporal coordination between TC-NER, HR, and C-NHEJ in active chromatin is still poorly defined, particularly in non-dividing or terminally differentiated cells where repair template availability is limited.

Another major question relates to the functional role of RNA transcripts and R-loops in the repair of transcription-associated DSB. While accumulating evidence supports a positive contribution of RNA templates to HR through DR-loop formation, the molecular determinants that distinguish physiological R-loops from pathological structures remain elusive. It is also unclear which RNA species (pre-existing transcripts, antisense RNAs, or small noncoding RNAs) serve as bona fide repair templates in mammalian cells. Identifying the factors that mediate RNA-directed repair without triggering deleterious R-loop-dependent genome instability will be an important direction for future work. In addition, TRCs represent an emerging source of DSBs, but the molecular sensors and signaling cascades that determine whether these lesions are resolved by HR, C-NHEJ, or replication restart mechanisms are still being elucidated. Understanding how transcription factors, chromatin remodelers, and topoisomerases cooperate to prevent or resolve TRCs in human cells remains a key open question. Furthermore, how the phase separation of transcription and repair factors at damage sites affects repair pathway selection and chromatin restoration is an area only recently being explored.

Finally, many questions remain regarding the integration of TCR with epigenetic and metabolic signaling. The influence of histone modifications, RNA modifications (such as m6A), and chromatin topology on TCR efficiency and fidelity is still largely correlative. Future studies combining live-cell imaging, nascent RNA sequencing, and high-resolution proteomics are needed to map these interdependent layers and to define how transcriptional programs shape DNA damage responses at a systems level.

These challenges underscore that transcription-associated DNA repair represents a dynamic and multilayered regulatory network. Deciphering how transcriptional activity, chromatin context, and RNA metabolism jointly determine repair outcome will be essential for understanding genome stability maintenance and its perturbation in cancer and neurodegenerative disease.

4. Posttranslational Regulation of Repair Pathway Choice and Chromatin Modifications

Posttranslational modifications (PTM) orchestrated by chromatin-bound E3 ubiquitin ligases RNF8 and RNF168 critically influence the selection of DSB repair pathways and, ultimately, cell fate [65]. The RNF8-RNF168 ubiquitin ligase cascade constitutes a pivotal signaling module that shapes the outcome of DSB repair. Although the RNF168-ubiquitin-53BP1 branch predominantly favors end joining, and the RNF8-K63-linked ubiquitin-BRCA1/RAD51 route facilitates recombination, both ligases can participate in either pathway depending on the chromatin and cell-cycle context. RNF8 acts as the initial ubiquitin E3 ligase at DNA damage sites, assembling K63-linked chains on histone substrates that recruit both 53BP1 and BRCA1 adaptors. Subsequently, RNF168 amplifies this ubiquitin signal and extends histone modification spreading around the lesion, with its expression levels and chromatin occupancy influencing the balance between C-NHEJ and HR engagement. Evidence from RIDDLE syndrome cells harboring RNF168 mutations and from Rnf168-knockout mice demonstrates that insufficient RNF168 impairs 53BP1 recruitment but not BRCA1 or RAD51, suggesting that higher RNF168 levels favor C-NHEJ, while reduced levels shift preference toward HR [66, 67]. Notably, RNF8 and RNF168 do not act in a mutually exclusive manner. Rather than functioning in isolation, RNF8 and RNF168 establish an interconnected ubiquitin-dependent signaling framework that allows the repair machinery to dynamically choose between pathways. This network supports the rapid but error-prone C-NHEJ response as an initial attempt, with the potential to shift toward HR during the S and G2 phases, when sister chromatids provide an available template. Their coordinated ubiquitin activity also influences the behavior of the BLM helicase: RNF8/RNF168-dependent ubiquitylation facilitates BLM recruitment to stalled replication forks, an event linked to the transient attenuation of recombination activity [65]. The principle remains strongly supported by recent investigations. For instance, Chauhan et al. (2025) [68] uncovered a SUMOPIN1-dependent mechanism limiting RNF168 chromatin retention and preventing excessive ubiquitin signaling and 53BP1 over-recruitment. Furthermore, Yang et al. (2024) [69] described how RNF168 directs ubiquitin-dependent recruitment of DDR factors including 53BP1, reflecting its central role in tuning repair pathway choice.

ASF1, along with its paralogs ASF1A and ASF1B, functions as a histone chaperone facilitating the transfer of H3-H4 dimers to CAF1 for nucleosome assembly. In response to DNA damage, DNA-PKcs-mediated phosphorylation of ASF1A at Ser192 enhances its interaction with histones and promotes HR by supporting MMS22L-TONSL-driven RAD51 loading onto ssDNA [70]. Conversely, phosphorylation of ASF1A at Ser166 by CHK1 (activated via ATM signaling) facilitates C-NHEJ by promoting ASF1A-MDC1 interaction, which enhances MDC1 phosphorylation by ATM, leads to histone ubiquitination, and facilitates 53BP1 recruitment through the H2AX-MDC1-ATM-RNF8 pathway [71]. ASF1, together with RIF1, also promotes C-NHEJ by inducing chromatin compaction around DSBs, thus influencing pathway selection [72]. ASF1’s role in repair pathway preference depends on chromatin accessibility and protein recruitment. In heterochromatin, ASF1 associates with 53BP1 and RIF1 to promote C-NHEJ. When resected DNA is present and C-NHEJ is compromised, ASF1 may instead facilitate RAD51 loading and activate HR. Beyond its chaperone activity, ASF1 may exert additional regulatory functions, and its paralogs likely exhibit context-dependent roles in DNA repair.

Specific histone marks and variant incorporation strongly influence which DSB repair pathway becomes engaged. The replacement of canonical histones variants: H3.3 (by CHD2) or H2A.Z (by p400) facilitates the recruitment of proteins associated with end joining [73]. Additional post-translational changes: H4Y51 phosphorylation by TIE2, conversion of H2BK120ub to acetylated H2BK120 via the SAGA complex, and H3K36 dimethylation catalyzed by SETMAR have also been linked to promoting C-NHEJ. Modifications including H2AK15ub (mediated by RNF168), H4K20me1/2, H3K416me1, and H3K79me2 contribute to the recruitment of anti-resection proteins such as 53BP1, thus stabilizing DNA ends and favoring C-NHEJ [74, 75]. Furthermore, SETD1A-BOD1L-dependent methylation of H3K4 supports RIF1 recruitment and biases repair toward C-NHEJ [73]. In contrast, HR is favored under chromatin states enriched in macroH2AX and depleted of marks such as H2A.Z, H2BK120ub, H3K4me3, H3K79me2, or H4K12ac near DSB. H3K9 dimethylation promotes the accumulation of BRCA1 and BRAD1, thereby stimulating HR activity. During the G2 phase, acetylation of H2AK15 and H4K16 interferes with 53BP1 binding and facilitates DNA-end resection through BRCA1- and BRAD1-dependent mechanisms [73].

Despite major progress in understanding how PTM and chromatin remodeling influence DSB repair, several mechanistic aspects remain unresolved. It is still unclear how sequential ubiquitination and deubiquitination events mediated by RNF8, RNF168, BRCC36, and USP48 are coordinated to balance 53BP1 versus BRCA1 recruitment. The temporal coupling between histone modification and chromatin accessibility during repair progression also requires clarification, particularly regarding how histone chaperones such as ASF1 and remodelers like INO80 or SMARCA5 interpret and reset specific histone marks. In addition, growing evidence suggests that metabolic intermediates directly modulate chromatin-based repair regulation. Nuclear acetyl-CoA availability shapes histone acetylation landscapes and impacts damage signaling [76, 77] while NAD⁺ functions as a critical substrate for sirtuins and PARPs involved in chromatin regulation and DSB repair [78]. How cells locally regulate the availability of these metabolites at DNA lesions, and whether metabolic fluctuations bias repair pathway selection toward HR or C-NHEJ, remains an open and important question for future research. Addressing these interconnected issues will be critical for developing a unified model of how PTM networks, chromatin dynamics, and cellular metabolism collectively determine genome stability.

5. Dysregulation of DNA Damage Response and its Implications in Tumorigenesis

Disruptions in DDR mechanisms are key contributors to oncogenesis and malignant transformation. Cells with defective DNA repair systems accumulate genomic instability, which in turn accelerates cancer progression. In addition to chromatin modifications, the stability and activity of DNA repair proteins can be modulated through epigenetic mechanisms, contributing to homologous recombination deficiency (HRD). Epigenetic silencing of BRCA1 through promoter hypermethylation occurs in a variety of malignancies. Most notably breast and ovarian cancer, but also in uterine, gastric, colorectal, pancreatic, and non-small cell lung tumors [79].

The FANC-BRCA repair axis is frequently compromised in ovarian cancers due to CpG island methylation of HR-related genes; for example, BRCA1 promoter hypermethylation has been directly associated with poor prognosis in high-grade serous ovarian carcinoma [80]. Somatic or germline mutations in several HR components have been implicated in diverse malignancies. Alterations in CtIP and RAD54 have been reported in non-Hodgkin’s lymphoma and colorectal cancer, whereas RAD51B mutations are recurrent in benign mesenchymal tumors such as lipomas and uterine leiomyomas. Changes affecting RAD51C are frequently detected in breast and ovarian cancers, while PALB2 inactivation is observed in gastro-intestinal tumors. Moreover, defects in the RecQ helicases BLM and WRN have been broadly linked to genomic instability across multiple cancer types.

Paradoxically, upregulated expression of RAD51 has been observed in several tumor types; this trend is reinforced by a recent report indicating that RAD51 upregulation correlates with metastasis and therapy resistance across solid tumors [81]. This overexpression is frequently driven by promoter hyperactivation [82, 83] and mutant KRAS signaling [84], resulting in elevated DSB repair capacity. However, excessive RAD51 levels are associated with poor clinical outcomes and chemoresistance [81, 85], prompting the development of RAD51-targeted inhibitors as a therapeutic strategy [86]. Moreover, comprehensive functional screening such as the “HR eye” assay not only allows ultra-rapid detection of HR activity but has already uncovered previously unrecognized mutations in HR genes like RAD51D [87].

Enhanced activity of the C-NHEJ pathway also promotes malignant transformation. Aberrations in XRCC4, LIG4, and XLF have been correlated with worse prognosis and therapy resistance [88], which has justified the development of DNA ligase IV inhibitors [88, 89]. A recent study by Hildebrand et al. (2025) [90] further exemplifies this approach by testing SCR130, a ligand IV inhibitor, in HNSCC cell lines, where it demonstrated moderate radiosensitizing effects. On the other hand, reduced expression of 53BP1, XRCC5, and DNA-PKcs has been shown to favor MMEJ/Alt-EJ, potentially increasing genomic instability through frequent chromosomal rearrangements [2]. This is supported by a study from 2025 showing that the senataxin helicase and DNA-PKcs act redundantly to promote C-NHEJ, underscoring the critical role of DNA-PKcs in maintaining DNA repair fidelity and suggesting that loss of DNA-PKcs function could shift repair toward more error-prone MMEJ/Alt-EJ pathways [91]. Loss of HR capacity forces tumor cells to exploit MMEJ/Alt-EJ as a compensatory repair route. This synthetic vulnerability forms the mechanistic basis for the clinical efficacy of PARP inhibitors in HR-deficient malignancies [2].

Despite growing understanding of how defective DNA repair drives tumorigenesis, several key questions remain unresolved. It is not fully clear how the balance between HR and C-NHEJ is dynamically rewired during malignant transformation or therapy resistance, nor which signaling networks determine whether enhanced repair confers survival advantage or triggers lethal genomic instability. The causal hierarchy among epigenetic events such as BRCA1 promoter hypermethylation, histone mark alterations, and chromatin compaction also remains to be established. Furthermore, the molecular mechanisms linking oncogenic signaling pathways (e.g., KRAS, MYC) to the upregulation of RAD51 and other repair factors are incompletely understood. How tumors fine-tune partial HR deficiency to achieve “repair plasticity” that supports survival under genotoxic stress is an emerging concept that warrants further study. Addressing these questions will be critical for identifying vulnerabilities that can be exploited therapeutically through synthetic lethality and pathway-specific inhibitors.

6. Pathway Choice and Its Impact on Genomic Stability

The selection among DSB repair pathways - C-NHEJ, MMEJ, or HR is a dynamic process that critically influences genome stability. The selection of a repair pathway is influenced by multiple parameters such as the cell-cycle phase, the nature and complexity of the DNA break, the chromatin landscape surrounding the lesion, and the local concentration of repair factors. Central to this regulation is DNA end resection and its tight coordination with cell cycle cues. C-NHEJ predominates in G1-phase, offering rapid, albeit error-prone repair in the absence of a sister chromatid. The S and G2 phases of the cell cycle favor HR, as sister chromatids act as faithful templates for DNA synthesis-based repair. Under conditions where C-NHEJ or HR are defective or saturated, the cell can engage the MMEJ/Alt-EJ mechanism, which relies on short microhomologies and frequently introduces small insertions or deletions.

The pathway selected for repair directly impacts genomic integrity. For instance, during S/G2, ionizing radiation-induced DSB are resolved through both C-NHEJ and HR [10], whereas in G1, C-NHEJ may process resected ends in a way that promotes chromosomal translocations [152, 153]. C-NHEJ, while efficient, may introduce small insertions or deletions, and excessive reliance on it can cause gross chromosomal abnormalities. Although HR tends to restore DNA faithfully, alternative processing routes like SSA or MMEJ/Alt-EJ following end resection can lead to deletions and genomic rearrangements [98, 114, 154]. MMEJ/Alt-EJ, being intrinsically inaccurate, is a known driver of such mutagenic outcomes.

Hence, while the plasticity in repair pathway choice supports cellular adaptability, its dysregulation (frequent in cancer) exacerbates genomic instability, fostering malignancy and therapeutic resistance. Understanding the molecular control of DNA repair provides the foundation for precision treatments that leverage cancer-specific repair weaknesses.

7. Conclusions

This review has summarized the complex regulatory networks that govern the cellular preference between error-prone DNA DSB repair pathways such as C-NHEJ and MMEJ/Alt-EJ and the high-fidelity HR pathway. We have outlined how key molecular players (ranging from chromatin modifiers and posttranslational regulators to transcription-coupled and metabolic factors) coordinate the dynamic “switch” between these routes. In each thematic section, we have highlighted outstanding questions that remain unresolved, underscoring the multifaceted nature of DSB repair regulation and its context dependence.

Given the central role of these pathways in maintaining genomic integrity, many of their core regulators have emerged as therapeutic targets, and several inhibitors are currently being evaluated for the treatment of repair-deficient cancers [155, 156, 157]. Advances in functional genomics, particularly CRISPR/Cas9-based screening technologies, continue to uncover novel repair components and to identify synthetic lethal interactions among HR, C-NHEJ, and accessory genes [158, 159, 160, 161]. Nevertheless, a major clinical obstacle remains the development of resistance to existing inhibitors — most notably PARP, ATR, ATM, WEE1, and PRMT inhibitors — often through the restoration of HR activity [162, 163]. Consequently, strategies aimed at achieving complete HR suppression, or at dual inhibition of HR and C-NHEJ, may offer improved therapeutic efficacy in specific tumor contexts [164, 165, 166]. The integration of mechanistic insight, functional screening, and targeted therapy design will be essential for translating our growing understanding of DSB repair into precision oncology. By systematically addressing open questions across each level of DSB regulation, future research can bridge molecular mechanisms with clinical applications, ultimately enabling rational combination therapies that exploit the vulnerabilities of repair-deficient cancers.

References

[1] Oh JM, Myung K. Crosstalk between different DNA repair pathways for DNA double strand break repairs. Mutation Research. Genetic Toxicology and Environmental Mutagenesis. 2022; 873: 503438. https://doi.org/10.1016/j.mrgentox.2021.503438.

[2] Caracciolo D, Riillo C, Di Martino MT, Tagliaferri P, Tassone P. Alternative Non-Homologous End-Joining: Error-Prone DNA Repair as Cancer’s Achilles’ Heel. Cancers. 2021; 13: 1392. https://doi.org/10.3390/cancers13061392.

[3] Dinant C, Houtsmuller AB, Vermeulen W. Chromatin structure and DNA damage repair. Epigenetics & Chromatin. 2008; 1: 9. https://doi.org/10.1186/1756-8935-1-9.

[4] Britton S, Coates J, Jackson SP. A new method for high-resolution imaging of Ku foci to decipher mechanisms of DNA double-strand break repair. The Journal of Cell Biology. 2013; 202: 579–595. https://doi.org/10.1083/jcb.201303073.

[5] Ghosh D, Raghavan SC. Nonhomologous end joining: new accessory factors fine tune the machinery. Trends in Genetics: TIG. 2021; 37: 582–599. https://doi.org/10.1016/j.tig.2021.03.001.

[6] Lieber MR. The mechanism of double-strand DNA break repair by the nonhomologous DNA end-joining pathway. Annual Review of Biochemistry. 2010; 79: 181–211. https://doi.org/10.1146/annurev.biochem.052308.093131.

[7] Frit P, Amin H, Zahid S, Barboule N, Hall C, Matharu G, et al. Structural and functional insights into the interaction between Ku70/80 and Pol X family polymerases in NHEJ. Nature Communications. 2025; 16: 4208. https://doi.org/10.1038/s41467-025-59133-2.

[8] Mi L, Cai Y, Qi J, Chen L, Li Y, Zhang S, et al. Elevated nonhomologous end-joining by AATF enables efficient DNA damage repair and therapeutic resistance in glioblastoma. Nature Communications. 2025; 16: 4941. https://doi.org/10.1038/s41467-025-60228-z.

[9] Neal JA, Dang V, Douglas P, Wold MS, Lees-Miller SP, Meek K. Inhibition of homologous recombination by DNA-dependent protein kinase requires kinase activity, is titratable, and is modulated by autophosphorylation. Molecular and Cellular Biology. 2011; 31: 1719–1733. https://doi.org/10.1128/MCB.01298-10.

[10] Drouet J, Frit P, Delteil C, de Villartay JP, Salles B, Calsou P. Interplay between Ku, Artemis, and the DNA-dependent protein kinase catalytic subunit at DNA ends. The Journal of Biological Chemistry. 2006; 281: 27784–27793. https://doi.org/10.1074/jbc.M603047200.

[11] Zhou Y, Lee JH, Jiang W, Crowe JL, Zha S, Paull TT. Regulation of the DNA Damage Response by DNA-PKcs Inhibitory Phosphorylation of ATM. Molecular Cell. 2017; 65: 91–104. https://doi.org/10.1016/j.molcel.2016.11.004.

[12] Shibata A, Conrad S, Birraux J, Geuting V, Barton O, Ismail A, et al. Factors determining DNA double-strand break repair pathway choice in G2 phase. The EMBO Journal. 2011; 30: 1079–1092. https://doi.org/10.1038/emboj.2011.27.

[13] Davis AJ, Chi L, So S, Lee KJ, Mori E, Fattah K, et al. BRCA1 modulates the autophosphorylation status of DNA-PKcs in S phase of the cell cycle. Nucleic Acids Research. 2014; 42: 11487–11501. https://doi.org/10.1093/nar/gku824.

[14] Xie Y, Liu YK, Guo ZP, Guan H, Liu XD, Xie DF, et al. RBX1 prompts degradation of EXO1 to limit the homologous recombination pathway of DNA double-strand break repair in G1 phase. Cell Death and Differentiation. 2020; 27: 1383–1397. https://doi.org/10.1038/s41418-019-0424-4.

[15] Hou J, Lu M, Guo J, Wu J, Wang C, Zhou PK, et al. DNA-PKcs, a player winding and dancing with RNA metabolism and diseases. Cellular & Molecular Biology Letters. 2025; 30: 25. https://doi.org/10.1186/s11658-025-00703-z.

[16] Pierce AJ, Hu P, Han M, Ellis N, Jasin M. Ku DNA end-binding protein modulates homologous repair of double-strand breaks in mammalian cells. Genes & Development. 2001; 15: 3237–3242. https://doi.org/10.1101/gad.946401.

[17] Wang Y, Czap MS, Kim H, Masaka PM, Wang H, Beg MA, et al. The mammalian Ku70 C-terminus SAP domain is required to repair DNA damage. Nucleic Acids Research. 2025; 53: gkaf499. https://doi.org/10.1093/nar/gkaf499.

[18] Lei T, Du S, Peng Z, Chen L. Multifaceted regulation and functions of 53BP1 in NHEJ mediated DSB repair (Review). International Journal of Molecular Medicine. 2022; 50: 90. https://doi.org/10.3892/ijmm.2022.5145.

[19] Rass E, Willaume S, Bertrand P. 53BP1: Keeping It under Control, Even at a Distance from DNA Damage. Genes. 2022; 13: 2390. https://doi.org/10.3390/genes13122390.

[20] Tripathi V, Nagarjuna T, Sengupta S. BLM helicase-dependent and -independent roles of 53BP1 during replication stress-mediated homologous recombination. The Journal of Cell Biology. 2007; 178: 9–14. https://doi.org/10.1083/jcb.200610051.

[21] Bunting SF, Callén E, Wong N, Chen HT, Polato F, Gunn A, et al. 53BP1 inhibits homologous recombination in Brca1-deficient cells by blocking resection of DNA breaks. Cell. 2010; 141: 243–254. https://doi.org/10.1016/j.cell.2010.03.012.

[22] Sivanand S, Rhoades S, Jiang Q, Lee JV, Benci J, Zhang J, et al. Nuclear Acetyl-CoA Production by ACLY Promotes Homologous Recombination. Molecular Cell. 2017; 67: 252–265.e6. https://doi.org/10.1016/j.molcel.2017.06.008.

[23] Escribano-Díaz C, Orthwein A, Fradet-Turcotte A, Xing M, Young JTF, Tkáč J, et al. A cell cycle-dependent regulatory circuit composed of 53BP1-RIF1 and BRCA1-CtIP controls DNA repair pathway choice. Molecular Cell. 2013; 49: 872–883. https://doi.org/10.1016/j.molcel.2013.01.001.

[24] Gupta R, Somyajit K, Narita T, Maskey E, Stanlie A, Kremer M, et al. DNA Repair Network Analysis Reveals Shieldin as a Key Regulator of NHEJ and PARP Inhibitor Sensitivity. Cell. 2018; 173: 972–988.e23. https://doi.org/10.1016/j.cell.2018.03.050.

[25] Swift ML, Zhou R, Syed A, Moreau LA, Tomasik B, Tainer JA, et al. Dynamics of the DYNLL1-MRE11 complex regulate DNA end resection and recruitment of Shieldin to DSBs. Nature Structural & Molecular Biology. 2023; 30: 1456–1467. https://doi.org/10.1038/s41594-023-01074-9.

[26] Mirman Z, Lottersberger F, Takai H, Kibe T, Gong Y, Takai K, et al. 53BP1-RIF1-shieldin counteracts DSB resection through CST- and Polα-dependent fill-in. Nature. 2018; 560: 112–116. https://doi.org/10.1038/s41586-018-0324-7.

[27] Mirman Z, Sasi NK, King A, Chapman JR, de Lange T. 53BP1-shieldin-dependent DSB processing in BRCA1-deficient cells requires CST-Polα-primase fill-in synthesis. Nature Cell Biology. 2022; 24: 51–61. https://doi.org/10.1038/s41556-021-00812-9.

[28] Clairmont CS, D’Andrea AD. REV7 directs DNA repair pathway choice. Trends in Cell Biology. 2021; 31: 965–978. https://doi.org/10.1016/j.tcb.2021.05.009.

[29] Zimmermann M, de Lange T. 53BP1: pro choice in DNA repair. Trends in Cell Biology. 2014; 24: 108–117. https://doi.org/10.1016/j.tcb.2013.09.003.

[30] Callen E, Di Virgilio M, Kruhlak MJ, Nieto-Soler M, Wong N, Chen HT, et al. 53BP1 mediates productive and mutagenic DNA repair through distinct phosphoprotein interactions. Cell. 2013; 153: 1266–1280. https://doi.org/10.1016/j.cell.2013.05.023.

[31] Wang J, Aroumougame A, Lobrich M, Li Y, Chen D, Chen J, et al. PTIP associates with Artemis to dictate DNA repair pathway choice. Genes & Development. 2014; 28: 2693–2698. https://doi.org/10.1101/gad.252478.114.

[32] Crickard JB. Discrete roles for Rad54 and Rdh54 during homologous recombination. Current Opinion in Genetics & Development. 2021; 71: 48–54. https://doi.org/10.1016/j.gde.2021.06.013.

[33] Kaur E, Agrawal R, Arun R, Madhavan V, Srivastava V, Kumar D, et al. Small molecules that disrupt RAD54-BLM interaction hamper tumor proliferation in colon cancer chemoresistance models. The Journal of Clinical Investigation. 2024; 134: e161941. https://doi.org/10.1172/JCI161941.

[34] So A, Dardillac E, Muhammad A, Chailleux C, Sesma-Sanz L, Ragu S, et al. RAD51 protects against nonconservative DNA double-strand break repair through a nonenzymatic function. Nucleic Acids Research. 2022; 50: 2651–2666. https://doi.org/10.1093/nar/gkac073.

[35] Zhong AX, Chen Y, Chen PL. BRCA1 the Versatile Defender: Molecular to Environmental Perspectives. International Journal of Molecular Sciences. 2023; 24: 14276. https://doi.org/10.3390/ijms241814276.

[36] Wang M, Li W, Tomimatsu N, Yu CH, Ji JH, Alejo S, et al. Crucial roles of the BRCA1-BARD1 E3 ubiquitin ligase activity in homology-directed DNA repair. Molecular Cell. 2023; 83: 3679–3691.e8. https://doi.org/10.1016/j.molcel.2023.09.015.

[37] Hodson C, Low JKK, van Twest S, Jones SE, Swuec P, Murphy V, et al. Mechanism of Bloom syndrome complex assembly required for double Holliday junction dissolution and genome stability. Proceedings of the National Academy of Sciences of the United States of America. 2022; 119: e2109093119. https://doi.org/10.1073/pnas.2109093119.

[38] Croteau DL, Popuri V, Opresko PL, Bohr VA. Human RecQ helicases in DNA repair, recombination, and replication. Annual Review of Biochemistry. 2014; 83: 519–552. https://doi.org/10.1146/annurev-biochem-060713-035428.

[39] Kaur E, Agrawal R, Sengupta S. Functions of BLM Helicase in Cells: Is It Acting Like a Double-Edged Sword? Frontiers in Genetics. 2021; 12: 634789. https://doi.org/10.3389/fgene.2021.634789.

[40] Thakkar MK, Lee J, Meyer S, Chang VY. RecQ Helicase Somatic Alterations in Cancer. Frontiers in Molecular Biosciences. 2022; 9: 887758. https://doi.org/10.3389/fmolb.2022.887758.

[41] Lee JH, Shamanna RA, Kulikowicz T, Borhan Fakouri N, Kim EW, Christiansen LS, et al. CDK2 phosphorylation of Werner protein (WRN) contributes to WRN’s DNA double-strand break repair pathway choice. Aging Cell. 2021; 20: e13484. https://doi.org/10.1111/acel.13484.

[42] Gupta P, Saha B, Chattopadhyay S, Patro BS. Pharmacological targeting of differential DNA repair, radio-sensitizes WRN-deficient cancer cells in vitro and in vivo. Biochemical Pharmacology. 2021; 186: 114450. https://doi.org/10.1016/j.bcp.2021.114450.

[43] Noto A, Valenzisi P, Di Feo F, Fratini F, Kulikowicz T, Sommers JA, et al. Phosphorylation-dependent WRN-RPA interaction promotes recovery of stalled forks at secondary DNA structure. Nature Communications. 2025; 16: 997. https://doi.org/10.1038/s41467-025-55958-z.

[44] Clairmont CS, Sarangi P, Ponnienselvan K, Galli LD, Csete I, Moreau L, et al. TRIP13 regulates DNA repair pathway choice through REV7 conformational change. Nature Cell Biology. 2020; 22: 87–96. https://doi.org/10.1038/s41556-019-0442-y.

[45] Unoki M, Sasaki H. The UHRF protein family in epigenetics, development, and carcinogenesis. Proceedings of the Japan Academy. Series B, Physical and Biological Sciences. 2022; 98: 401–415. https://doi.org/10.2183/pjab.98.021.

[46] Ashraf W, Ahmad T, Reynoird N, Hamiche A, Mély Y, Bronner C, et al. Natural and Synthetic Anticancer Epidrugs Targeting the Epigenetic Integrator UHRF1. Molecules (Basel, Switzerland). 2023; 28: 5997. https://doi.org/10.3390/molecules28165997.

[47] Zhang H, Liu H, Chen Y, Yang X, Wang P, Liu T, et al. A cell cycle-dependent BRCA1-UHRF1 cascade regulates DNA double-strand break repair pathway choice. Nature Communications. 2016; 7: 10201. https://doi.org/10.1038/ncomms10201.

[48] Liu Y, Hrit JA, Chomiak AA, Stransky S, Hoffman JR, Tiedemann RL, et al. DNA hypomethylation promotes UHRF1-and SUV39H1/H2-dependent crosstalk between H3K18ub and H3K9me3 to reinforce heterochromatin states. Molecular Cell. 2025; 85: 394–412.e12. https://doi.org/10.1016/j.molcel.2024.11.009.

[49] Li Y, Yang X, Cai H, Wang F. Homologous recombination deficiency among patients with germline or somatic non-BRCA1/2 homologous recombination repair gene variations. NPJ Precision Oncology. 2025; 9: 192. https://doi.org/10.1038/s41698-025-00999-2.

[50] Sharma S, Javadekar SM, Pandey M, Srivastava M, Kumari R, Raghavan SC. Homology and enzymatic requirements of microhomology-dependent alternative end joining. Cell Death & Disease. 2015; 6: e1697. https://doi.org/10.1038/cddis.2015.58.

[51] Bennardo N, Cheng A, Huang N, Stark JM. Alternative-NHEJ is a mechanistically distinct pathway of mammalian chromosome break repair. PLoS Genetics. 2008; 4: e1000110. https://doi.org/10.1371/journal.pgen.1000110.

[52] Della-Maria J, Zhou Y, Tsai MS, Kuhnlein J, Carney JP, Paull TT, et al. Human Mre11/human Rad50/Nbs1 and DNA ligase IIIalpha/XRCC1 protein complexes act together in an alternative nonhomologous end joining pathway. The Journal of Biological Chemistry. 2011; 286: 33845–33853. https://doi.org/10.1074/jbc.M111.274159.

[53] Wang H, Gao B, Wu Z, Wang H, Dong M. Alteration of serum adropin level in preeclampsia. Pregnancy Hypertension. 2017; 8: 6–8. https://doi.org/10.1016/j.preghy.2017.02.003.

[54] Ahrabi S, Sarkar S, Pfister SX, Pirovano G, Higgins GS, Porter ACG, et al. A role for human homologous recombination factors in suppressing microhomology-mediated end joining. Nucleic Acids Research. 2016; 44: 5743–5757. https://doi.org/10.1093/nar/gkw326.

[55] Deng SK, Gibb B, de Almeida MJ, Greene EC, Symington LS. RPA antagonizes microhomology-mediated repair of DNA double-strand breaks. Nature Structural & Molecular Biology. 2014; 21: 405–412. https://doi.org/10.1038/nsmb.2786.

[56] Jiang Y, Yam JC, Tham CC, Pang CP, Chu WK. RB Regulates DNA Double Strand Break Repair Pathway Choice by Mediating CtIP Dependent End Resection. International Journal of Molecular Sciences. 2020; 21: 9176. https://doi.org/10.3390/ijms21239176.

[57] Kent T, Chandramouly G, McDevitt SM, Ozdemir AY, Pomerantz RT. Mechanism of microhomology-mediated end-joining promoted by human DNA polymerase θ. Nature Structural & Molecular Biology. 2015; 22: 230–237. https://doi.org/10.1038/nsmb.2961.

[58] Ito F, Li Z, Minakhin L, Khant HA, Pomerantz RT, Chen XS. Structural basis for Polθ-helicase DNA binding and microhomology-mediated end-joining. Nature Communications. 2025; 16: 3725. https://doi.org/10.1038/s41467-025-58441-x.

[59] Teicher BA, Dexheimer TS, Chen L, Silvers T, Jones EM, Coussens NP, et al. Polymerase Ѳ inhibitors combinations with approved and investigational agents in patient-derived tumor multi-cell type (mct) spheroids. Experimental and Molecular Pathology. 2025; 143: 104978. https://doi.org/10.1016/j.yexmp.2025.104978.

[60] Bhargava R, Onyango DO, Stark JM. Regulation of Single-Strand Annealing and its Role in Genome Maintenance. Trends in Genetics: TIG. 2016; 32: 566–575. https://doi.org/10.1016/j.tig.2016.06.007.

[61] Zhang JP, Li XL, Li GH, Chen W, Arakaki C, Botimer GD, et al. Efficient precise knockin with a double cut HDR donor after CRISPR/Cas9-mediated double-stranded DNA cleavage. Genome Biology. 2017; 18: 35. https://doi.org/10.1186/s13059-017-1164-8.

[62] Yao X, Wang X, Hu X, Liu Z, Liu J, Zhou H, et al. Homology-mediated end joining-based targeted integration using CRISPR/Cas9. Cell Research. 2017; 27: 801–814. https://doi.org/10.1038/cr.2017.76.

[63] Han W, Li Z, Guo Y, He K, Li W, Xu C, et al. Efficient precise integration of large DNA sequences with 3’-overhang dsDNA donors using CRISPR/Cas9. Proceedings of the National Academy of Sciences of the United States of America. 2023; 120: e2221127120. https://doi.org/10.1073/pnas.2221127120.

[64] Tei C, Hata S, Mabuchi A, Okuda S, Ito KK, Genova M, et al. Comparative analysis of multiple DNA double-strand break repair pathways in CRISPR-mediated endogenous tagging. Communications Biology. 2025; 8: 749. https://doi.org/10.1038/s42003-025-08187-5.

[65] Tan J, Sun X, Zhao H, Guan H, Gao S, Zhou PK. Double-strand DNA break repair: molecular mechanisms and therapeutic targets. MedComm. 2023; 4: e388. https://doi.org/10.1002/mco2.388.

[66] Stewart GS, Stankovic T, Byrd PJ, Wechsler T, Miller ES, Huissoon A, et al. RIDDLE immunodeficiency syndrome is linked to defects in 53BP1-mediated DNA damage signaling. Proceedings of the National Academy of Sciences of the United States of America. 2007; 104: 16910–16915. https://doi.org/10.1073/pnas.0708408104.

[67] Zong D, Adam S, Wang Y, Sasanuma H, Callén E, Murga M, et al. BRCA1 Haploinsufficiency Is Masked by RNF168-Mediated Chromatin Ubiquitylation. Molecular Cell. 2019; 73: 1267–1281.e7. https://doi.org/10.1016/j.molcel.2018.12.010.

[68] Chauhan AS, Mackintosh MJW, Cassar J, Lanz AJ, Jamshad M, Mackay HL, et al. PIN1-SUMO2/3 motif suppresses excessive RNF168 chromatin accumulation and ubiquitin signaling to promote IR resistance. Nature Communications. 2025; 16: 3399. https://doi.org/10.1038/s41467-025-56974-9.

[69] Yang Y, Jayaprakash D, Jhujh SS, Reynolds JJ, Chen S, Gao Y, et al. PCNA-binding activity separates RNF168 functions in DNA replication and DNA double-stranded break signaling. Nucleic Acids Research. 2024; 52: 13019–13035. https://doi.org/10.1093/nar/gkae918.

[70] Huang TH, Fowler F, Chen CC, Shen ZJ, Sleckman B, Tyler JK. The Histone Chaperones ASF1 and CAF-1 Promote MMS22L-TONSL-Mediated Rad51 Loading onto ssDNA during Homologous Recombination in Human Cells. Molecular Cell. 2018; 69: 879–892.e5. https://doi.org/10.1016/j.molcel.2018.01.031.

[71] Lee KY, Dutta A. Chk1 promotes non-homologous end joining in G1 through direct phosphorylation of ASF1A. Cell Reports. 2021; 34: 108680. https://doi.org/10.1016/j.celrep.2020.108680.

[72] Feng S, Ma S, Li K, Gao S, Ning S, Shang J, et al. RIF1-ASF1-mediated high-order chromatin structure safeguards genome integrity. Nature Communications. 2022; 13: 957. https://doi.org/10.1038/s41467-022-28588-y.

[73] Min S, Ji JH, Heo Y, Cho H. Transcriptional regulation and chromatin dynamics at DNA double-strand breaks. Experimental & Molecular Medicine. 2022; 54: 1705–1712. https://doi.org/10.1038/s12276-022-00862-5.

[74] Bayley R, Borel V, Moss RJ, Sweatman E, Ruis P, Ormrod A, et al. H3K4 methylation by SETD1A/BOD1L facilitates RIF1-dependent NHEJ. Molecular Cell. 2022; 82: 1924–1939.e10. https://doi.org/10.1016/j.molcel.2022.03.030.

[75] Kolobynina KG, Rapp A, Cardoso MC. Chromatin Ubiquitination Guides DNA Double Strand Break Signaling and Repair. Frontiers in Cell and Developmental Biology. 2022; 10: 928113. https://doi.org/10.3389/fcell.2022.928113.

[76] Bradshaw PC. Acetyl-CoA Metabolism and Histone Acetylation in the Regulation of Aging and Lifespan. Antioxidants (Basel, Switzerland). 2021; 10: 572. https://doi.org/10.3390/antiox10040572.

[77] Willnow P, Teleman AA. Nuclear position and local acetyl-CoA production regulate chromatin state. Nature. 2024; 630: 466–474. https://doi.org/10.1038/s41586-024-07471-4.

[78] Ruszkiewicz JA, Bürkle A, Mangerich A. Fueling genome maintenance: On the versatile roles of NAD⁺ in preserving DNA integrity. The Journal of Biological Chemistry. 2022; 298: 102037. https://doi.org/10.1016/j.jbc.2022.102037.

[79] Mekonnen N, Yang H, Shin YK. Homologous Recombination Deficiency in Ovarian, Breast, Colorectal, Pancreatic, Non-Small Cell Lung and Prostate Cancers, and the Mechanisms of Resistance to PARP Inhibitors. Frontiers in Oncology. 2022; 12: 880643. https://doi.org/10.3389/fonc.2022.880643.

[80] Tsuchimochi S, Yamamoto Y, Taguchi A, Kawazu M, Sone K, Ikemura M, et al. BRCA1 Promoter Methylation in Ovarian Cancer: Clinical Relevance and a Novel Diagnostic Approach Using Fragment Analysis. Cancer Science. 2025; 116: 1996–2007. https://doi.org/10.1111/cas.70078.

[81] Wang Z, Jia R, Wang L, Yang Q, Hu X, Fu Q, et al. The Emerging Roles of Rad51 in Cancer and Its Potential as a Therapeutic Target. Frontiers in Oncology. 2022; 12: 935593. https://doi.org/10.3389/fonc.2022.935593.

[82] Li Y, Yu H, Luo RZ, Zhang Y, Zhang MF, Wang X, et al. Elevated expression of Rad51 is correlated with decreased survival in resectable esophageal squamous cell carcinoma. Journal of Surgical Oncology. 2011; 104: 617–622. https://doi.org/10.1002/jso.22018.

[83] Song JE, Alves TC, Stutz B, Šestan-Peša M, Kilian N, Jin S, et al. Mitochondrial Fission Governed by Drp1 Regulates Exogenous Fatty Acid Usage and Storage in Hela Cells. Metabolites. 2021; 11: 322. https://doi.org/10.3390/metabo11050322.

[84] Hu J, Zhang Z, Zhao L, Li L, Zuo W, Han L. High expression of RAD51 promotes DNA damage repair and survival in KRAS-mutant lung cancer cells. BMB Reports. 2019; 52: 151–156. https://doi.org/10.5483/BMBRep.2019.52.2.213.

[85] Compadre AJ, van Biljon LN, Valentine MC, Llop-Guevara A, Graham E, Fashemi B, et al. RAD51 Foci as a Biomarker Predictive of Platinum Chemotherapy Response in Ovarian Cancer. Clinical Cancer Research: an Official Journal of the American Association for Cancer Research. 2023; 29: 2466–2479. https://doi.org/10.1158/1078-0432.CCR-22-3335.

[86] Shkundina IS, Gall AA, Dick A, Cocklin S, Mazin AV. New RAD51 Inhibitors to Target Homologous Recombination in Human Cells. Genes. 2021; 12: 920. https://doi.org/10.3390/genes12060920.

[87] Saito S, Kato S, Arai U, En A, Tsunezumi J, Mizushima T, et al. HR eye & MMR eye: one-day assessment of DNA repair-defective tumors eligible for targeted therapy. Nature Communications. 2025; 16: 4239. https://doi.org/10.1038/s41467-025-59462-2.

[88] Cheong A, Nagel ZD. Human Variation in DNA Repair, Immune Function, and Cancer Risk. Frontiers in Immunology. 2022; 13: 899574. https://doi.org/10.3389/fimmu.2022.899574.

[89] Ray U, Gopinatha VK, Sharma S, Goyary L, Choudhary B, Mantelingu K, et al. Identification and characterization of mercaptopyrimidine-based small molecules as inhibitors of nonhomologous DNA end joining. The FEBS Journal. 2023; 290: 796–820. https://doi.org/10.1111/febs.16615.

[90] Hildebrand LS, Jost T, Schindler M, Derer A, Fuhrmann G, Fietkau R, et al. Inhibiting NHEJ in HNSCC cell lines by the ligase IV inhibitor SCR130 has limited radiosensitizing effects. Scientific Reports. 2025; 15: 17871. https://doi.org/10.1038/s41598-025-03159-5.

[91] Chen BR, Pham T, Reynolds LD, Dang N, Zhang Y, Manalang K, et al. Senataxin and DNA-PKcs redundantly promote non-homologous end joining repair of DNA double strand breaks during V(D)J recombination. Science Advances. 2025; 11: eads5272. https://doi.org/10.1126/sciadv.ads5272.

[92] Kumari N, Kaur E, Raghavan SC, Sengupta S. Regulation of pathway choice in DNA repair after double-strand breaks. Current Opinion in Pharmacology. 2025; 80: 102496. https://doi.org/10.1016/j.coph.2024.102496.

[93] Uziel T, Lerenthal Y, Moyal L, Andegeko Y, Mittelman L, Shiloh Y. Requirement of the MRN complex for ATM activation by DNA damage. The EMBO Journal. 2003; 22: 5612–5621. https://doi.org/10.1093/emboj/cdg541.

[94] Stucki M, Jackson SP. gammaH2AX and MDC1: anchoring the DNA-damage-response machinery to broken chromosomes. DNA Repair. 2006; 5: 534–543. https://doi.org/10.1016/j.dnarep.2006.01.012.

[95] Thompson LH. Recognition, signaling, and repair of DNA double-strand breaks produced by ionizing radiation in mammalian cells: the molecular choreography. Mutation Research. 2012; 751: 158–246. https://doi.org/10.1016/j.mrrev.2012.06.002.

[96] Thorslund T, Ripplinger A, Hoffmann S, Wild T, Uckelmann M, Villumsen B, et al. Histone H1 couples initiation and amplification of ubiquitin signalling after DNA damage. Nature. 2015; 527: 389–393. https://doi.org/10.1038/nature15401.

[97] Mattiroli F, Vissers JHA, van Dijk WJ, Ikpa P, Citterio E, Vermeulen W, et al. RNF168 ubiquitinates K13-15 on H2A/H2AX to drive DNA damage signaling. Cell. 2012; 150: 1182–1195. https://doi.org/10.1016/j.cell.2012.08.005.

[98] Chang HHY, Pannunzio NR, Adachi N, Lieber MR. Non-homologous DNA end joining and alternative pathways to double-strand break repair. Nature Reviews. Molecular Cell Biology. 2017; 18: 495–506. https://doi.org/10.1038/nrm.2017.48.

[99] Noordermeer SM, Adam S, Setiaputra D, Barazas M, Pettitt SJ, Ling AK, et al. The shieldin complex mediates 53BP1-dependent DNA repair. Nature. 2018; 560: 117–121. https://doi.org/10.1038/s41586-018-0340-7.

[100] Setiaputra D, Durocher D. Shieldin - the protector of DNA ends. EMBO Reports. 2019; 20: e47560. https://doi.org/10.15252/embr.201847560.

[101] Zhu Q, Pao GM, Huynh AM, Suh H, Tonnu N, Nederlof PM, et al. BRCA1 tumour suppression occurs via heterochromatin-mediated silencing. Nature. 2011; 477: 179–184. https://doi.org/10.1038/nature10371.

[102] Tripathi V, Agarwal H, Priya S, Batra H, Modi P, Pandey M, et al. MRN complex-dependent recruitment of ubiquitylated BLM helicase to DSBs negatively regulates DNA repair pathways. Nature Communications. 2018; 9: 1016. https://doi.org/10.1038/s41467-018-03393-8.

[103] Truong LN, Li Y, Shi LZ, Hwang PYH, He J, Wang H, et al. Microhomology-mediated End Joining and Homologous Recombination share the initial end resection step to repair DNA double-strand breaks in mammalian cells. Proceedings of the National Academy of Sciences of the United States of America. 2013; 110: 7720–7725. https://doi.org/10.1073/pnas.1213431110.

[104] Keeney S, Chang GJ, Linn S. Characterization of a human DNA damage binding protein implicated in xeroderma pigmentosum E. The Journal of Biological Chemistry. 1993; 268: 21293–21300.

[105] Legerski R, Peterson C. Correction: Expression cloning of a human DNA repair gene involved in xeroderma pigmentosum group C. Nature. 1992; 360: 610. https://doi.org/10.1038/360610b0.

[106] Nishi R, Okuda Y, Watanabe E, Mori T, Iwai S, Masutani C, et al. Centrin 2 stimulates nucleotide excision repair by interacting with xeroderma pigmentosum group C protein. Molecular and Cellular Biology. 2005; 25: 5664–5674. https://doi.org/10.1128/MCB.25.13.5664-5674.2005.

[107] An S, Kusakabe M, Kim HS, Kozono H, Cheon NY, Kim J, et al. XPC-RAD23B enhances UV-DDB binding to DNA to facilitate lesion search in nucleotide excision repair. Nucleic Acids Research. 2025; 53: gkaf463. https://doi.org/10.1093/nar/gkaf463.

[108] Nakazawa Y, Oka Y, Matsunaga T, Ogi T. Transcription-coupled repair - mechanisms of action, regulation, and associated human disorders. FEBS Letters. 2025; 599: 166–167. https://doi.org/10.1002/1873-3468.15073.

[109] Khobta A, Sarmini L. Transcription-Coupled Nucleotide Excision Repair: A Faster Solution or the Only Option? Biomolecules. 2025; 15: 1026. https://doi.org/10.3390/biom15071026.

[110] Chaurasia P, Sen R, Pandita TK, Bhaumik SR. Preferential repair of DNA double-strand break at the active gene in vivo. The Journal of Biological Chemistry. 2012; 287: 36414–36422. https://doi.org/10.1074/jbc.M112.364661.

[111] Bohr VA, Smith CA, Okumoto DS, Hanawalt PC. DNA repair in an active gene: removal of pyrimidine dimers from the DHFR gene of CHO cells is much more efficient than in the genome overall. Cell. 1985; 40: 359–369. https://doi.org/10.1016/0092-8674(85)90150-3.

[112] Wright WD, Shah SS, Heyer WD. Homologous recombination and the repair of DNA double-strand breaks. The Journal of Biological Chemistry. 2018; 293: 10524–10535. https://doi.org/10.1074/jbc.TM118.000372.

[113] Aymard F, Bugler B, Schmidt CK, Guillou E, Caron P, Briois S, et al. Transcriptionally active chromatin recruits homologous recombination at DNA double-strand breaks. Nature Structural & Molecular Biology. 2014; 21: 366–374. https://doi.org/10.1038/nsmb.2796.

[114] Scully R, Panday A, Elango R, Willis NA. DNA double-strand break repair-pathway choice in somatic mammalian cells. Nature Reviews. Molecular Cell Biology. 2019; 20: 698–714. https://doi.org/10.1038/s41580-019-0152-0.

[115] Yang H, Lan L. Transcription-coupled DNA repair protects genome stability upon oxidative stress-derived DNA strand breaks. FEBS Letters. 2025; 599: 168–176. https://doi.org/10.1002/1873-3468.14938.

[116] Saur F, Lesage E, Pradel L, Collins S, Finoux AL, Alghoul E, et al. Transcriptional repression facilitates RNA:DNA hybrid accumulation at DNA double-strand breaks. Nature Cell Biology. 2025; 27: 992–1005. https://doi.org/10.1038/s41556-025-01669-y.

[117] Jalan M, Brambati A, Shah H, McDermott N, Patel J, Zhu Y, et al. RNA transcripts serve as a template for double-strand break repair in human cells. Nature Communications. 2025; 16: 4349. https://doi.org/10.1038/s41467-025-59510-x.

[118] Mojumdar A, Granger C, Lunke M, Evans E, Cobb JA. Dna2 nuclease resolves RNA:DNA hybrids at double-strand breaks. iScience. 2025; 28: 113235. https://doi.org/10.1016/j.isci.2025.113235.

[119] Ouyang J, Yadav T, Zhang JM, Yang H, Rheinbay E, Guo H, et al. RNA transcripts stimulate homologous recombination by forming DR-loops. Nature. 2021; 594: 283–288. https://doi.org/10.1038/s41586-021-03538-8.

[120] Pessina F, Giavazzi F, Yin Y, Gioia U, Vitelli V, Galbiati A, et al. Functional transcription promoters at DNA double-strand breaks mediate RNA-driven phase separation of damage-response factors. Nature Cell Biology. 2019; 21: 1286–1299. https://doi.org/10.1038/s41556-019-0392-4.

[121] Liu S, Hua Y, Wang J, Li L, Yuan J, Zhang B, et al. RNA polymerase III is required for the repair of DNA double-strand breaks by homologous recombination. Cell. 2021; 184: 1314–1329.e10. https://doi.org/10.1016/j.cell.2021.01.048.

[122] Bader AS, Bushell M. DNA:RNA hybrids form at DNA double-strand breaks in transcriptionally active loci. Cell Death & Disease. 2020; 11: 280. https://doi.org/10.1038/s41419-020-2464-6.

[123] Modafferi S, Esposito F, Tavella S, Gioia U, Francia S. Traffic light at DSB-transit regulation between gene transcription and DNA repair. FEBS Letters. 2025; 599: 177–189. https://doi.org/10.1002/1873-3468.15024.

[124] Keskin H, Shen Y, Huang F, Patel M, Yang T, Ashley K, et al. Transcript-RNA-templated DNA recombination and repair. Nature. 2014; 515: 436–439. https://doi.org/10.1038/nature13682.

[125] Wei L, Nakajima S, Böhm S, Bernstein KA, Shen Z, Tsang M, et al. DNA damage during the G0/G1 phase triggers RNA-templated, Cockayne syndrome B-dependent homologous recombination. Proceedings of the National Academy of Sciences of the United States of America. 2015; 112: E3495–E3504. https://doi.org/10.1073/pnas.1507105112.

[126] Chen L, Chen JY, Zhang X, Gu Y, Xiao R, Shao C, et al. R-ChIP Using Inactive RNase H Reveals Dynamic Coupling of R-loops with Transcriptional Pausing at Gene Promoters. Molecular Cell. 2017; 68: 745–757.e5. https://doi.org/10.1016/j.molcel.2017.10.008.

[127] Dumelie JG, Jaffrey SR. Defining the location of promoter-associated R-loops at near-nucleotide resolution using bisDRIP-seq. eLife. 2017; 6: e28306. https://doi.org/10.7554/eLife.28306.

[128] Skourti-Stathaki K, Kamieniarz-Gdula K, Proudfoot NJ. R-loops induce repressive chromatin marks over mammalian gene terminators. Nature. 2014; 516: 436–439. https://doi.org/10.1038/nature13787.

[129] Cooke MB, Herman C, Sivaramakrishnan P. Clues to transcription/replication collision-induced DNA damage: it was RNAP, in the chromosome, with the fork. FEBS Letters. 2025; 599: 209–243. https://doi.org/10.1002/1873-3468.15063.

[130] Zhu MY, Zhang YZ, Guo T, Ren J. Nascent RNA at the crossroad of transcription and replication. Trends in Genetics: TIG. 2025. https://doi.org/10.1016/j.tig.2025.05.010. (online ahead of print)

[131] Bayona-Feliu A, Aguilera A. Transcription-Replication Conflicts: Unlocking New Frontiers in Cancer. BioEssays: News and Reviews in Molecular, Cellular and Developmental Biology. 2025; 47: e70025. https://doi.org/10.1002/bies.70025.

[132] Hu Y, Stillman B. Origins of DNA replication in eukaryotes. Molecular Cell. 2023; 83: 352–372. https://doi.org/10.1016/j.molcel.2022.12.024.

[133] Hamperl S, Bocek MJ, Saldivar JC, Swigut T, Cimprich KA. Transcription-Replication Conflict Orientation Modulates R-Loop Levels and Activates Distinct DNA Damage Responses. Cell. 2017; 170: 774–786.e19. https://doi.org/10.1016/j.cell.2017.07.043.

[134] Goehring L, Huang TT, Smith DJ. Transcription-Replication Conflicts as a Source of Genome Instability. Annual Review of Genetics. 2023; 57: 157–179. https://doi.org/10.1146/annurev-genet-080320-031523.

[135] Werner M, Trauner M, Schauer T, Ummethum H, Márquez-Gómez E, Lalonde M, et al. Transcription-replication conflicts drive R-loop-dependent nucleosome eviction and require DOT1L activity for transcription recovery. Nucleic Acids Research. 2025; 53: gkaf109. https://doi.org/10.1093/nar/gkaf109.

[136] Hartono SR, Malapert A, Legros P, Bernard P, Chédin F, Vanoosthuyse V. The Affinity of the S9.6 Antibody for Double-Stranded RNAs Impacts the Accurate Mapping of R-Loops in Fission Yeast. Journal of Molecular Biology. 2018; 430: 272–284. https://doi.org/10.1016/j.jmb.2017.12.016.

[137] Gorthi A, Romero JC, Loranc E, Cao L, Lawrence LA, Goodale E, et al. EWS-FLI1 increases transcription to cause R-loops and block BRCA1 repair in Ewing sarcoma. Nature. 2018; 555: 387–391. https://doi.org/10.1038/nature25748.

[138] Kotsantis P, Silva LM, Irmscher S, Jones RM, Folkes L, Gromak N, et al. Increased global transcription activity as a mechanism of replication stress in cancer. Nature Communications. 2016; 7: 13087. https://doi.org/10.1038/ncomms13087.

[139] Hatchi E, Skourti-Stathaki K, Ventz S, Pinello L, Yen A, Kamieniarz-Gdula K, et al. BRCA1 recruitment to transcriptional pause sites is required for R-loop-driven DNA damage repair. Molecular Cell. 2015; 57: 636–647. https://doi.org/10.1016/j.molcel.2015.01.011.

[140] Shivji MKK, Renaudin X, Williams ÇH, Venkitaraman AR. BRCA2 Regulates Transcription Elongation by RNA Polymerase II to Prevent R-Loop Accumulation. Cell Reports. 2018; 22: 1031–1039. https://doi.org/10.1016/j.celrep.2017.12.086.

[141] Petermann E, Lan L, Zou L. Sources, resolution and physiological relevance of R-loops and RNA-DNA hybrids. Nature Reviews. Molecular Cell Biology. 2022; 23: 521–540. https://doi.org/10.1038/s41580-022-00474-x.

[142] Santos-Pereira JM, Aguilera A. R loops: new modulators of genome dynamics and function. Nature Reviews. Genetics. 2015; 16: 583–597. https://doi.org/10.1038/nrg3961.

[143] Damasceno JD, Briggs EM, Krasilnikova M, Marques CA, Lapsley C, McCulloch R. R-loops acted on by RNase H1 influence DNA replication timing and genome stability in Leishmania. Nature Communications. 2025; 16: 1470. https://doi.org/10.1038/s41467-025-56785-y.

[144] Sollier J, Stork CT, García-Rubio ML, Paulsen RD, Aguilera A, Cimprich KA. Transcription-coupled nucleotide excision repair factors promote R-loop-induced genome instability. Molecular Cell. 2014; 56: 777–785. https://doi.org/10.1016/j.molcel.2014.10.020.

[145] Makharashvili N, Arora S, Yin Y, Fu Q, Wen X, Lee JH, et al. Sae2/CtIP prevents R-loop accumulation in eukaryotic cells. eLife. 2018; 7: e42733. https://doi.org/10.7554/eLife.42733.

[146] Matos DA, Zhang JM, Ouyang J, Nguyen HD, Genois MM, Zou L. ATR Protects the Genome against R Loops through a MUS81-Triggered Feedback Loop. Molecular Cell. 2020; 77: 514–527.e4. https://doi.org/10.1016/j.molcel.2019.10.010.

[147] Chang EYC, Tsai S, Aristizabal MJ, Wells JP, Coulombe Y, Busatto FF, et al. MRE11-RAD50-NBS1 promotes Fanconi Anemia R-loop suppression at transcription-replication conflicts. Nature Communications. 2019; 10: 4265. https://doi.org/10.1038/s41467-019-12271-w.

[148] Bhatia V, Barroso SI, García-Rubio ML, Tumini E, Herrera-Moyano E, Aguilera A. BRCA2 prevents R-loop accumulation and associates with TREX-2 mRNA export factor PCID2. Nature. 2014; 511: 362–365. https://doi.org/10.1038/nature13374.

[149] García-Rubio ML, Pérez-Calero C, Barroso SI, Tumini E, Herrera-Moyano E, Rosado IV, et al. The Fanconi Anemia Pathway Protects Genome Integrity from R-loops. PLoS Genetics. 2015; 11: e1005674. https://doi.org/10.1371/journal.pgen.1005674.

[150] Chung C, Verheijen BM, Navapanich Z, McGann EG, Shemtov S, Lai GJ, et al. Evolutionary conservation of the fidelity of transcription. Nature Communications. 2023; 14: 1547. https://doi.org/10.1038/s41467-023-36525-w.

[151] Chung C, Verheijen BM, Zhang X, Huang B, Coakley A, McGann E, et al. The fidelity of transcription in human cells. Proceedings of the National Academy of Sciences of the United States of America. 2023; 120: e2210038120. https://doi.org/10.1073/pnas.2210038120.

[152] Barton O, Naumann SC, Diemer-Biehs R, Künzel J, Steinlage M, Conrad S, et al. Polo-like kinase 3 regulates CtIP during DNA double-strand break repair in G1. The Journal of Cell Biology. 2014; 206: 877–894. https://doi.org/10.1083/jcb.201401146.

[153] Biehs R, Steinlage M, Barton O, Juhász S, Künzel J, Spies J, et al. DNA Double-Strand Break Resection Occurs during Non-homologous End Joining in G1 but Is Distinct from Resection during Homologous Recombination. Molecular Cell. 2017; 65: 671–684.e5. https://doi.org/10.1016/j.molcel.2016.12.016.

[154] Mateos-Gomez PA, Kent T, Deng SK, McDevitt S, Kashkina E, Hoang TM, et al. The helicase domain of Polθ counteracts RPA to promote alt-NHEJ. Nature Structural & Molecular Biology. 2017; 24: 1116–1123. https://doi.org/10.1038/nsmb.3494.

[155] Hopkins JL, Lan L, Zou L. DNA repair defects in cancer and therapeutic opportunities. Genes & Development. 2022; 36: 278–293. https://doi.org/10.1101/gad.349431.122.

[156] Baxter JS, Zatreanu D, Pettitt SJ, Lord CJ. Resistance to DNA repair inhibitors in cancer. Molecular Oncology. 2022; 16: 3811–3827. https://doi.org/10.1002/1878-0261.13224.

[157] Groelly FJ, Fawkes M, Dagg RA, Blackford AN, Tarsounas M. Targeting DNA damage response pathways in cancer. Nature Reviews. Cancer. 2023; 23: 78–94. https://doi.org/10.1038/s41568-022-00535-5.

[158] Awwad SW, Serrano-Benitez A, Thomas JC, Gupta V, Jackson SP. Revolutionizing DNA repair research and cancer therapy with CRISPR-Cas screens. Nature Reviews. Molecular Cell Biology. 2023; 24: 477–494. https://doi.org/10.1038/s41580-022-00571-x.

[159] Chen BR, Sleckman BP. A Whole Genome CRISPR/Cas9 Screening Approach for Identifying Genes Encoding DNA End-Processing Proteins. Methods in Molecular Biology (Clifton, N.J.). 2022; 2444: 15–27. https://doi.org/10.1007/978-1-0716-2063-2_2.

[160] Fielden J, Siegner SM, Gallagher DN, Schröder MS, Dello Stritto MR, Lam S, et al. Comprehensive interrogation of synthetic lethality in the DNA damage response. Nature. 2025; 640: 1093–1102. https://doi.org/10.1038/s41586-025-08815-4.

[161] Wu Y, Wang Y, Wang Y, Qiu H, Yuan X, Xiong H, et al. Exploring synthetic lethality in cancer therapy: CRISPR-Cas9 technology offers new hope. Biochimica et Biophysica Acta. Reviews on Cancer. 2025; 1880: 189370. https://doi.org/10.1016/j.bbcan.2025.189370.

[162] Kulkarni S, Brownlie J, Jeyapalan JN, Mongan NP, Rakha EA, Madhusudan S. Evolving DNA repair synthetic lethality targets in cancer. Bioscience Reports. 2022; 42: BSR20221713. https://doi.org/10.1042/BSR20221713.

[163] Peng S, Long M, Chen Q, Yin Z, Zeng C, Zhang W, et al. Perspectives on cancer therapy-synthetic lethal precision medicine strategies, molecular mechanisms, therapeutic targets and current technical challenges. Cell Death Discovery. 2025; 11: 179. https://doi.org/10.1038/s41420-025-02418-8.

[164] Smith G, Alholm Z, Coleman RL, Monk BJ. DNA Damage Repair Inhibitors-Combination Therapies. Cancer Journal (Sudbury, Mass.). 2021; 27: 501–505. https://doi.org/10.1097/PPO.0000000000000561.

[165] Liu Y. How is synthetic lethality and CRISPR able to revolutionize the field of cancer treatment? Journal of Student Research. 2025; 14. https://doi.org/10.47611/jsrhs.v14i1.8539.

[166] Xiong Q, Jin J, Lin J, Zhang B, Jiang Y, Sun Z, et al. The role of synthetic lethality in overcoming cancer therapy resistance: Emerging paradigm and recent advances. Drug Resistance Updates: Reviews and Commentaries in Antimicrobial and Anticancer Chemotherapy. 2025; 83: 101290. https://doi.org/10.1016/j.drup.2025.101290.

Funding

Vedecká grantová agentúra (VEGA) - Grant Agency of the Slovak Republic(2/0140/23)

Recovery and Resilience Plan of the Slovak Republic under the project [CU-ESG-2024-05-PD](09I03-03-V05-00019)