Research Progress on DNA Damage Repair Efficiency Assessment Strategies

Xiao Liu; Yiwei Zhu; Ling Li; Siyu Hang; Sipei Wu; Song-Bai Liu

doi:10.3791/71031

Review Article

Research Progress on DNA Damage Repair Efficiency Assessment Strategies

DOI:

10.3791/71031

⸱

June 23rd, 2026

Xiao Liu^*¹^,² , Yiwei Zhu^*³ , Ling Li¹^,² , Siyu Hang¹ , Sipei Wu⁴ , Song-Bai Liu¹^,²

¹Jiangsu Province Engineering Research Center of Development and Translation of Key Technologies for Chronic Disease Prevention and Control, Suzhou Vocational Health College, ²School of Chemistry and Life Sciences, Suzhou University of Science and Technology, ³Department of Orthopedics, The First Affiliated Hospital of Soochow University, ⁴Department of Hematology, The First Affiliated Hospital of Soochow University

^* These authors contributed equally

Summary

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DNA damage repair efficiency assays can be broadly categorized into three groups: cell-based functional assays, in vitro biochemical assays, and molecular marker assays. These approaches enable qualitative or quantitative evaluation of repair efficiency across multiple DNA damage repair pathways.

Abstract

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Cellular genomic DNA is continuously exposed to endogenous and exogenous factors that induce various forms of DNA damage. In mammalian cells, the daily burden of DNA damage may reach 10⁴–10⁵ lesions per cell. DNA damage that persists into the S phase can lead to mutations. When such mutations occur in oncogenes, tumor suppressor genes, or genes involved in cell proliferation, they may promote cellular transformation and carcinogenesis. Therefore, DNA damage repair plays a critical role in maintaining genomic stability and preventing cancer development. Assessment of DNA repair capacity is best achieved by measuring the functional activity of DNA repair systems. DNA damage repair pathways and their repair efficiencies are highly relevant to chemotherapeutic drug development, gene function validation, and cancer therapy. Accordingly, investigation of the repair efficiency of individual DNA damage repair pathways has important theoretical and practical significance. This review summarizes the major DNA damage repair pathways, methods used to assess repair efficiency, and the advantages and limitations of these approaches.

Introduction

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The DNA damage repair system is a critical cellular defense mechanism that maintains genomic stability by reducing mutation frequency and suppressing carcinogenesis¹. Different types of DNA damage are repaired through distinct but interconnected repair pathways². For example, base mismatches generated during DNA replication are corrected through mismatch repair (MMR), whereas chemically modified bases, such as alkylated bases, are primarily repaired through base excision repair (BER)³. Ultraviolet (UV)-induced 6-4 photoproducts and bulky nucleotide adducts generated by certain chemical agents are removed through nucleotide excision repair (NER). Interstrand crosslink (ICL) damage is repaired through coordinated activities of proteins associated with the Fanconi anemia pathway⁴. Single-strand breaks (SSBs) are repaired through the single-strand break repair (SSBR) pathway, whereas double-strand breaks (DSBs) are primarily repaired through nonhomologous end joining (NHEJ) or homologous recombination (HR). In most cases, complex DNA lesions are resolved through coordinated interactions among multiple DNA repair pathways⁵.

Assessment of DNA damage and repair efficiency in cells typically relies on a combination of experimental approaches, including the Comet assay, immunofluorescence (IF), Western blotting (WB), host cell reactivation (HCR), and plasmid-based repair assays. These methods evaluate DNA damage and repair by detecting DNA strand breaks, DNA damage markers, and the expression or localization of repair-associated proteins⁶. However, accurate quantification of cellular repair efficiency generally requires dynamic challenge-based assays. In these approaches, specific DNA lesions are induced experimentally, and cells are sampled at sequential time points to monitor the formation and clearance of damage signals. Comparison of residual damage levels over time relative to the initial damage burden enables quantitative assessment of repair kinetics and pathway functionality.

To systematically compare the characteristics and applications of currently available DNA damage repair efficiency assays, this review summarizes major detection methods according to five parameters: lesion/substrate specificity, analytical resolution, sensitivity, quantitative capability, and instrumentation requirements (Table 1). This review aims to provide a practical methodological guide for selecting appropriate approaches to investigate DNA damage repair pathways in basic, translational, and clinical research.

Detection method	Lesion/substrate specificity	Analytical resolution	Sensitivity/detection threshold	Quantitative vs. qualitative output	Instrumentation requirements
Western blot	CPDs, 6-4PPs, repair proteins (e.g., MGMT, RAD51)	Population	Moderate	Semi-quantitative	Protein electrophoresis, membrane transfer, chemiluminescence imaging system
Immunofluorescence	CPDs, DSB foci (e.g., γ-H2AX, RAD51)	Single-cell	Moderate	Semi-quantitative	Fluorescence microscope
HPLC-MS/MS	CPDs, 6-4PPs, oxidative and alkylative base damage	Population/In vitro	Extremely high	Quantitative	HPLC-MS/MS
MGMT enzyme activity assay	O⁶-meG alkylation damage	Population/In vitro	High	Quantitative	Microplate reader
Comet assay	SSBs, DSBs, oxidized bases (with FPG/OGG pretreatment)	Single-cell	High	Quantitative	Fluorescence microscope/electrophoresis system
Incision assay	AP sites, NER incisions, BER incisions	Population/In vitro	High	Quantitative	Gel imaging, radiolabeling/fluorescence detection
Reconstituted repair system	Site-specific base damage, AP sites, strand breaks	In vitro	High	Quantitative	Gel electrophoresis, fluorescence/MS detection
Plasmid-based repair assay	Site-specific damage	Population	Moderate	Quantitative	Microplate reader, flow cytometer, colony counting
Reporter assay	Site-specific damage; BER, NER, HR, NHEJ, MMR	Population	High	Quantitative	Microplate reader, flow cytometer, fluorescence microscope
HCR/FM-HCR	UV damage, platinum crosslinks, site-specific damage	Population	High	Quantitative	Flow cytometer, fluorescence/chemiluminescence detector
RADD	Broad-spectrum DNA damage	Single-cell	Extremely high	Quantitative	Single-molecule fluorescence microscope
qPCR	UV damage, chain-blocking lesions	Population	Moderate	Quantitative	Real-time qPCR instrument
MSI detection	MMR deficiency-associated microsatellite instability	Population	Moderate	Qualitative	PCR instrument/capillary electrophoresis
DR-GFP/EJ5-GFP	HR- and NHEJ-mediated DSB repair	Population	High	Quantitative	Flow cytometer/fluorescence microscope
BLESS/BLISS	Genome-wide DSB mapping	Single-cell	Extremely high	Quantitative	High-throughput sequencer
iPOND	DDR proteins at replication forks/TLS-associated repair	Replication fork/proteome	High	Semi-quantitative	Mass spectrometry/sequencing platform

Table 1: Systematic comparison of methods for detecting DNA damage repair efficiency. Comparison of commonly used methods for assessing DNA damage repair efficiency across multiple repair pathways. The table summarizes lesion or substrate specificity, analytical resolution, sensitivity, quantitative capability, and instrumentation requirements for each assay, including biochemical, molecular, and cell-based approaches.

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Review and Perspective

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1. Direct repair (DR)

Direct repair (DR) is the simplest form of DNA repair and refers to the direct restoration of damaged DNA without removal of bases or nucleotides⁷. DR primarily repairs two major types of DNA lesions. The first is cyclobutane pyrimidine dimer (CPD) damage induced by ultraviolet (UV) radiation. In many organisms, photolyase recognizes and binds pyrimidine dimers. Upon exposure to visible light, photolyase becomes activated, directly reverses DNA damage, and then dissociates from the DNA molecule. This photoreactivation mechanism is particularly important in plants and lower organisms. However, placental mammals, including humans, lack functional photolyase and therefore cannot perform photoreactivation. Consequently, UV-induced CPD damage in human cells is repaired primarily through nucleotide excision repair (NER)⁸.

The second major type of damage repaired through DR is alkylation-induced base damage. Repair is mediated by O⁶-alkylguanine-DNA alkyltransferase (AGT), also known as O⁶-methylguanine-DNA methyltransferase (MGMT). This enzyme transfers alkyl groups from the O⁶ position of guanine residues onto itself, thereby directly restoring the damaged DNA base.

Methods for detecting DR efficiency

Detection of repair efficiency for CPD damage induced by ultraviolet (UV) radiation
1. Western blot (WB) assay for detecting changes in CPD content
  Using antibodies specific for CPDs, investigators can detect changes in CPD abundance at different time points following UV irradiation and infer repair efficiency from the level of residual damage. Higher repair efficiency corresponds to more rapid CPD removal. WB-based CPD detection is a classical and effective approach for assessing DNA repair activity; however, several limitations exist. First, the assay measures average damage levels across cell populations and therefore lacks single-cell resolution. Second, and more importantly, the assay cannot distinguish among repair pathways. Because CPDs in human cells are primarily repaired through NER, WB analysis alone cannot differentiate whether observed repair activity reflects photolyase-mediated repair or NER activity without additional experimental context.
2. Immunofluorescence (IF) positioning CPD focus dynamic changes
  Cells subjected to UV irradiation and photorepair treatment can be stained using CPD-specific antibodies, and changes in nuclear CPD foci can be visualized by fluorescence microscopy. Quantitative analysis of CPD foci formation and clearance enables assessment of DNA repair capacity at both population and single-cell levels. The rate of CPD foci disappearance directly reflects repair efficiency. IF provides spatial and temporal visualization of repair dynamics within individual cells and allows detection of cell-to-cell heterogeneity. However, fluorescence signals are susceptible to quenching, and quantitative analysis often relies on manual counting, which introduces subjectivity. In mammalian cells lacking photoreactivation activity, this method primarily reflects NER-mediated repair efficiency.
3. High-performance liquid chromatography-mass spectrometry (HPLC-MS/MS) for quantitative CPD analysis
  Extracted DNA is enzymatically digested into nucleosides, after which CPDs are separated from normal nucleosides by high-performance liquid chromatography (HPLC). Quantitative analysis is subsequently performed using tandem mass spectrometry (MS/MS). Repair efficiency is determined by comparing CPD abundance before and after repair⁹. HPLC-MS/MS provides extremely high specificity and picomolar-level sensitivity; however, the method requires expensive instrumentation, complex sample preparation, and does not reflect the in situ repair state of intact cells. In human cell samples, the measured repair outcome predominantly reflects NER activity.
Efficacy of DNA base damage repair induced by alkylating agents
1. MGMT Enzyme Activity Assay
  Repair efficiency of DNA damage induced by alkylating agents is commonly evaluated by measuring MGMT activity¹⁰. Radioactively labeled methylated DNA substrates are used to monitor transfer of methyl groups from damaged DNA to the MGMT protein, thereby enabling quantitative assessment of repair efficiency.
2. Reporting gene assay for detecting O⁶-meG repair efficiency
  Reporter plasmids containing O⁶-methylguanine (O⁶-meG) lesions can be constructed to assess repair efficiency. The introduced lesion suppresses reporter gene expression until MGMT-mediated repair restores reporter activity. Repair efficiency is subsequently quantified by measuring reporter gene expression levels. Reporter gene assays provide intuitive and quantitative evaluation of repair activity in living cells¹¹. However, these assays require construction of lesion-specific vectors and involve relatively long experimental cycles¹².
Damage repair efficiency upon detection of chain breakage
1. Comet Assay
  The Comet assay, also known as single-cell gel electrophoresis (SCGE), is a sensitive method for detecting DNA strand breaks at the single-cell level. Damaged DNA fragments migrate from the nucleus during electrophoresis, forming comet-like tails, whereas intact DNA remains concentrated within the nuclear region. Parameters such as comet tail length, tail DNA content, and tail moment can be quantified using fluorescence microscopy and image analysis software. These metrics enable quantitative evaluation of DNA strand break severity. By comparing comet profiles immediately after DNA damage induction with those obtained after repair, repair efficiency can be estimated¹³.
  The Comet assay offers several advantages, including simple operation, relatively low cost, high sensitivity, and single-cell resolution. Further refinement of this method has led to development of enzyme-modified variants and comet-based in vitro DNA repair assays, which can be used to evaluate BER and NER activities.

2. Base excision repair (BER)

Base excision repair (BER) is a highly conserved DNA repair pathway that primarily removes small base lesions generated by oxidation, alkylation, and deamination¹⁴. BER shares substantial mechanistic overlap with single-strand break repair (SSBR), and certain SSBR events are considered extensions of BER. According to the repair patch length, BER is categorized into short-patch BER (SP-BER) and long-patch BER (LP-BER).

BER is initiated by DNA glycosylases that recognize and excise damaged bases by cleaving the N-glycosidic bond between the base and deoxyribose, thereby generating apurinic/apyrimidinic (AP) sites¹⁵. AP endonuclease 1 (APE1) subsequently cleaves the DNA backbone at the 5′ side of the AP site, producing 3′-OH and 5′-deoxyribose phosphate (dRP) termini. During SP-BER, the dRP group is removed, DNA polymerase β (Polβ) fills the single-nucleotide gap, and DNA ligase III (LigIII) completes ligation.

Repair of alkylation-induced single-strand breaks containing 3′-blocked termini follows a similar mechanism but requires additional end processing. Polynucleotide kinase 3′-phosphatase (PNKP) first removes abnormal 3′ termini to generate a proper 3′-OH group. Polβ subsequently fills the missing nucleotide, and LigIII seals the nick to complete repair.

Methods for detecting BER efficiency

Enzyme-modified comet assay
The enzyme-modified Comet assay extends the standard Comet assay by incorporating lesion-specific repair enzymes. These enzymes recognize and cleave damaged bases, converting otherwise undetectable lesions into detectable strand breaks. The resulting comet tail intensity reflects the level of DNA damage¹⁶˒¹⁷.
To evaluate BER efficiency, formamidopyrimidine DNA glycosylase (FPG) or its mammalian homolog 8-oxoguanine DNA glycosylase (OGG1) is used to convert oxidized base lesions into single-strand breaks, which are subsequently analyzed using the Comet assay¹⁸.
Incision assay
The incision assay is a core in vitro approach for evaluating BER activity. DNA substrates containing AP sites are incubated with purified enzymes or cellular extracts, and the proportion of cleaved DNA substrates is quantified. This assay directly reflects APE1-mediated incision activity and provides high specificity and sensitivity while minimizing interference from the intracellular environment¹⁹.
However, the incision assay does not fully recapitulate BER activity in living cells because it cannot assess dynamic regulation of the complete repair process, including lesion recognition, base excision, AP site cleavage, DNA synthesis, and ligation²⁰.
Reconstituted repair system
The reconstituted repair system is an in vitro experimental platform that reconstructs DNA repair pathways using purified repair proteins. This system enables detailed analysis of the molecular functions and interactions of individual repair proteins and is widely used for mechanistic studies and drug screening.
To evaluate BER efficiency, purified glycosylases, AP endonucleases, DNA polymerases, and ligases are sequentially incubated with damaged DNA substrates to reconstitute lesion recognition, excision, synthesis, and ligation. Repair products are analyzed by gel electrophoresis, fluorescence labeling, or mass spectrometry²¹. Although this approach enables highly controlled mechanistic studies, it does not account for chromatin structure or global cellular regulation of repair activity²².
Plasmid-based repair assay
Plasmid-based repair assays involve introduction of plasmids containing defined DNA lesions into host cells or incubation with cellular extracts containing repair enzymes, followed by assessment of repair efficiency using reporter assays, flow cytometry, or colony formation analysis²³.
To evaluate BER efficiency, plasmids containing oxidative DNA damage are incubated with cellular extracts and subsequently transformed into repair-deficient bacterial strains, such as Escherichia coli. Repair efficiency is then estimated from colony formation rates.
Reporter gene assay
Reporter gene assays indirectly evaluate DNA repair efficiency or gene regulatory activity by measuring restoration of reporter gene expression²⁴. In these assays, DNA lesions or regulatory elements are incorporated into reporter constructs, and repair efficiency is quantified through changes in reporter activity following repair.
For BER analysis, an 8-oxoG lesion can be inserted into the coding region of a luciferase reporter gene. If the lesion remains unrepaired, translation is disrupted; successful repair restores full-length reporter protein expression. Undamaged plasmids, such as pRL-TK Renilla luciferase vectors, are commonly used as internal controls.
Host cell reactivation (HCR)
Host cell reactivation (HCR) is a classical assay that indirectly evaluates DNA repair efficiency by measuring the ability of host cells to restore expression from damaged reporter plasmids²⁵. Plasmids carrying defined lesions, such as UV-induced pyrimidine dimers or chemically induced crosslinks, are transfected into host cells. Repair efficiency is subsequently quantified by measuring reporter gene expression.
HCR assays provide high sensitivity and allow assessment of repair activity toward specific lesion types. However, efficient transfection and preparation of high-quality damaged plasmids are required. A major limitation of conventional HCR is that treatment of plasmids with DNA-damaging agents often generates heterogeneous lesion populations throughout the plasmid, thereby complicating mechanistic interpretation²⁶.
To address these limitations, flow cytometry-based host cell reactivation (FM-HCR) assays using site-specific single-lesion plasmids have been developed. FM-HCR enables rapid quantitative analysis of repair activity in living cells, although the assay requires specialized instrumentation and has limitations for simultaneous mechanistic analysis of multiple BER pathways²⁷.
Plaque host cell reactivation (PL-HCR) assays based on lesion-specific BER substrates have also been developed to investigate BER mechanisms in living cells (Figure 1)²⁸. In this approach, plasmids containing a single 1,N⁶-ethenoadenine (εA) lesion are introduced into mammalian cells. Following repair, plasmid DNA is recovered, digested with restriction enzymes, transformed into E. coli, and quantified by plaque formation analysis. Completely repaired plasmids become linearized after restriction digestion and fail to generate plaques, whereas unrepaired or incompletely repaired plasmids remain circular and form plaques. BER efficiency is therefore determined by plaque quantification.
Repair-assisted damage detection
Repair-assisted damage detection (RADD) is a single-molecule method for detecting and quantifying DNA damage²⁹. In this approach, lesion-specific repair enzymes recognize and excise damaged bases, after which DNA polymerases incorporate fluorescently labeled nucleotides into the resulting gaps. DNA molecules are subsequently visualized by fluorescence microscopy, enabling direct detection and quantification of DNA lesions (Figure 2)³⁰.
RADD provides highly sensitive qualitative and quantitative analysis of DNA damage at the single-molecule level. However, the method primarily measures DNA damage burden rather than directly evaluating repair kinetics or overall repair efficiency in cell populations.

Plasmid DNA transfection, restriction enzyme digestion, and plaque assay process diagram.
Figure 1: Plaque host cell reactivation (PL-HCR). M13mp18 circular single-stranded DNA containing a defined lesion is transfected into competent cells. Following transfection, cells are cultured on agar plates to allow plaque formation. Replicative-form M13 DNA is subsequently isolated from plaques, digested with restriction enzymes, and analyzed by gel electrophoresis to distinguish mutant and wild-type products and determine whether lesion repair has occurred. Please click here to view a larger version of this figure.

DNA damage assessment diagram; DNA repair monitoring, fluorescence intensity analysis.
Figure 2: Repair-assisted damage detection (RADD). Single-stranded gaps generated during DNA repair are filled with biotin-labeled nucleotides. Fluorescently labeled avidin or streptavidin is then used to detect incorporated nucleotides, enabling quantitative assessment of DNA damage levels by fluorescence analysis. Please click here to view a larger version of this figure.

3. Nucleotide excision repair (NER)

Nucleotide excision repair (NER) is a major DNA repair pathway responsible for removing bulky DNA lesions that distort the DNA double helix³¹. NER substrates include UV-induced cyclobutane pyrimidine dimers (CPDs), 6-4 pyrimidine-pyrimidone photoproducts (6-4PPs), oxidative purine lesions, bulky chemical adducts, and cisplatin-induced intrastrand crosslinks³². Defects in NER are associated with disorders such as xeroderma pigmentosum, which is characterized by extreme UV sensitivity and elevated cancer susceptibility.

NER consists of two major subpathways: global genome NER (GG-NER)³³ and transcription-coupled NER (TC-NER)³⁴. GG-NER surveys the entire genome for helix-distorting lesions, thereby suppressing mutagenesis and carcinogenesis. In contrast, TC-NER selectively removes transcription-blocking lesions from actively transcribed genes and is essential for maintaining transcriptional integrity. Defects in TC-NER contribute to disorders such as Cockayne syndrome and UV-sensitive syndrome³⁵.

Methods for detecting NER efficiency

Incision assay
The incision assay evaluates NER efficiency by detecting strand incisions introduced at DNA damage sites by repair proteins. In this assay, damaged DNA substrates, such as UV-irradiated DNA containing CPDs, are incubated with repair proteins or cellular extracts, and incision products are subsequently detected³⁶.
Radiolabeling or fluorescence labeling can be used to improve sensitivity and permit precise monitoring of incision activity³⁷. The assay directly measures the activity of key NER proteins responsible for damage recognition and excision. However, successful implementation requires preparation of high-quality damaged substrates and active protein extracts.
Enzyme repair comet assay
NER efficiency can also be assessed using enzyme-modified Comet assays. In this approach, T4 endonuclease V is incubated with damaged cells to convert pyrimidine dimers into detectable single-strand breaks, which are subsequently analyzed using the Comet assay³⁸.
Real-time fluorescence quantitative PCR (qPCR)
Real-time quantitative PCR (qPCR) indirectly evaluates DNA repair efficiency by measuring the inhibitory effects of DNA lesions on PCR amplification³⁹. During NER analysis, UV irradiation is first used to induce pyrimidine dimers. These lesions impede DNA polymerase progression and reduce amplification efficiency, resulting in increased cycle threshold (Ct) values. Following repair by the NER pathway, DNA integrity is restored, and amplification efficiency improves, leading to lower Ct values. Repair efficiency can therefore be quantified by comparing amplification before and after repair relative to an undamaged internal control region⁴⁰.
In addition, qPCR can be used to assess expression levels of genes involved in DNA damage signaling and repair pathways, including ATM, ATR, BRCA1, CDKN1A, and XPC, thereby indirectly reflecting DNA repair capacity⁴¹.
Flow cytometry-based host cell reactivation (FM-HCR)
FM-HCR assays utilize site-specific lesion-containing plasmids transfected into host cells and quantify repair efficiency by flow cytometric analysis of fluorescence intensity and the proportion of fluorescent cells (Figure 3)⁴²˒⁴³. Increased fluorescence intensity generally reflects greater DNA repair capacity⁴⁴˒⁴⁵.
Excision repair sequencing (XR-seq)
Excision repair sequencing (XR-seq) is a genome-wide technique for mapping NER activity at single-nucleotide resolution. This method selectively enriches and sequences damage-containing oligonucleotides excised during NER, thereby generating high-resolution maps of repair activity across the genome. XR-seq enables quantitative analysis of repair efficiency and distribution patterns associated with both GG-NER and TC-NER pathways and represents a major technological advance in NER research.
Because NER consists of two mechanistically distinct subpathways and involves numerous repair proteins, multiple complementary methods are required for comprehensive assessment of repair activity⁴⁶˒⁴⁷. In addition to the methods described above, NER efficiency may also be evaluated using Western blotting, immunofluorescence assays, plasmid-based repair assays, and reporter gene assays.

DNA damage process diagram; flow cytometry for DRC analysis, plasmid co-transfection method.
Figure 3: Detection principle of flow cytometry-based host cell reactivation (FM-HCR). Plasmids containing four distinct types of DNA damage are co-transfected into cultured mammalian cells. DNA repair capacity (DRC) is subsequently quantified by flow cytometry based on reporter fluorescence intensity. DRC, DNA repair capacity. Please click here to view a larger version of this figure.

4. Mismatch repair (MMR)

Mismatch repair (MMR) is a postreplicative error-correction pathway responsible for repairing base mismatches, such as G-T and A-C mispairs, as well as small insertion-deletion loops generated during DNA replication⁴⁸. MMR is initiated when MutS recognizes mismatched bases, after which MutL recruits downstream repair factors and exonucleases to activate the repair process⁴⁹. A critical step in MMR is discrimination between the parental and newly synthesized DNA strands⁵⁰.

In prokaryotes, strand discrimination is achieved through DNA methylation, which marks the parental strand. In eukaryotes, strand identity is determined through transient nicks in newly synthesized DNA and interactions with proliferating cell nuclear antigen (PCNA). MMR substantially improves replication fidelity and plays an essential role in maintaining genomic stability⁵¹. Defects in MMR are strongly associated with multiple cancers, including Lynch syndrome.

Methods for assessing the efficiency of MMR

Microsatellite instability testing
Defective MMR results in failure to correct replication slippage errors at microsatellite loci, leading to microsatellite instability (MSI). MSI testing involves PCR amplification of representative microsatellite markers followed by capillary electrophoresis-based fragment analysis. Instability at multiple loci indicates impaired MMR activity, whereas stable microsatellite profiles indicate intact MMR function. MSI testing provides high sensitivity and specificity for identifying MMR deficiency; however, it remains an indirect measure of MMR function and does not quantitatively assess repair efficiency.
Reporter plasmids containing defined mismatches or insertion-deletion loops can also be used to evaluate MMR efficiency through reporter gene assays. In addition, Western blotting and immunofluorescence assays targeting MMR-associated proteins may provide indirect assessment of pathway activity.

5. Double-strand break (DSB) repair

Double-strand breaks (DSBs) are among the most cytotoxic forms of DNA damage. Failure to repair DSBs accurately can result in mutations, chromosomal deletions, translocations, genomic instability, and tumorigenesis. DSB repair primarily occurs through homologous recombination (HR) and nonhomologous end joining (NHEJ), which operate preferentially during different phases of the cell cycle⁵².

Methods for assessing the efficiency of DSB repair

gamma-H2AX foci assay
The γ-H2AX foci assay is one of the most widely used methods for indirectly detecting DSBs. Following DSB formation, histone H2AX is rapidly phosphorylated at Ser139 to generate γ-H2AX foci. The number of γ-H2AX foci correlates with the extent of DSB formation, and repair efficiency can be inferred by monitoring the rate of foci disappearance over time using immunofluorescence microscopy.
This assay offers several advantages, including high sensitivity, single-cell resolution, and established experimental protocols, making it a central method in DNA damage response (DDR) research.
BLESS and BLISS
BLESS (Breaks Labeling, Enrichment on Streptavidin, and Sequencing) and BLISS (Breaks Labeling In Situ and Sequencing) are genome-wide DSB mapping methods based on in situ labeling and high-throughput sequencing. Both approaches identify DSB sites through direct ligation of sequencing adapters to broken DNA ends, enabling precise genome-wide localization and quantification of DSBs. These methods can also be used to evaluate overall DSB repair kinetics.
In addition, DSB-specific reporter gene assays can quantitatively evaluate DSB repair efficiency and distinguish HR-mediated repair from NHEJ-mediated repair (Figure 4). However, although γ-H2AX assays, BLESS, and BLISS effectively measure global DSB burden and repair kinetics, they cannot determine whether repair occurred specifically through HR or NHEJ pathways.

DSB reporter transfection process diagram; microscopy analysis; DNA repair study.
Figure 4: Reporter gene assay for double-strand break (DSB) repair. DSB reporter plasmids are introduced into target cells. Following an appropriate repair period after transfection, reporter gene fluorescence is monitored using fluorescence microscopy to evaluate DSB repair efficiency. DSB, double-strand break. Please click here to view a larger version of this figure.

6. Homologous recombination repair (HR)

Homologous recombination (HR) is a high-fidelity DNA repair pathway that primarily functions during the S and G2 phases of the cell cycle, when an undamaged sister chromatid is available as a repair template⁵³. Following DSB formation, the MRE11-RAD50-NBS1 (MRN) complex recognizes DNA ends and initiates DNA end resection in cooperation with CtIP⁵⁴. This process generates 3′ single-stranded DNA (ssDNA) overhangs.

The resulting ssDNA is initially coated by replication protein A (RPA), which protects the DNA from nuclease degradation and prevents formation of secondary structures⁵⁵. Subsequently, BRCA2 mediates replacement of RPA with RAD51 recombinase, resulting in formation of a RAD51 nucleoprotein filament⁵⁶. This filament searches for homologous DNA sequences on the sister chromatid and promotes strand invasion into the homologous duplex DNA template, generating a displacement loop (D-loop) structure⁵⁷˒⁵⁸. DNA synthesis and strand exchange then proceed to complete repair.

HR efficiency can be quantitatively evaluated using DR-GFP reporter systems in reporter gene assays. Alternatively, indirect assessment may be performed using Western blotting or immunofluorescence assays targeting HR-associated proteins.

7. Non-homologous DNA end joining (NHEJ)

Nonhomologous end joining (NHEJ) repairs DSBs without requiring extensive sequence homology between DNA ends⁵⁹. Instead, broken DNA ends are directly processed and ligated, making NHEJ an inherently error-prone repair pathway. Because DNA ends may undergo degradation before ligation and ends from unrelated DSBs may be joined together, NHEJ can lead to mutations, deletions, and chromosomal rearrangements⁶⁰.

Three major forms of nonhomologous repair are recognized: classical NHEJ, single-strand annealing (SSA), and microhomology-mediated end joining (MMEJ).

Classical NHEJ
Classical NHEJ operates throughout the cell cycle. The Ku70/Ku80 heterodimer first recognizes and binds DSB ends⁶¹. This complex recruits DNA-dependent protein kinase catalytic subunit (DNA-PKcs), leading to activation of kinase signaling and initiation of NHEJ. End-processing enzymes, including nucleases and phosphatases, subsequently process damaged DNA termini before the XRCC4-DNA ligase IV complex mediates ligation of DNA ends⁶². Because terminal processing frequently results in nucleotide loss or inaccurate rejoining, NHEJ is considered a mutagenic repair pathway⁶³˒⁶⁴.
Single-strand annealing (SSA)
Single-strand annealing (SSA) repairs DSBs occurring between repeated homologous sequences. Following DSB formation, nucleases resect the 5′ DNA ends, generating extended 3′ ssDNA overhangs. RAD52 mediates annealing between complementary homologous repeats⁶⁵. Subsequently, the XPF-ERCC1 endonuclease complex removes unpaired DNA flaps, and the SLX1-SLX4 complex may also participate in end processing. DNA ligase then seals the remaining gaps. Because sequences located between homologous repeats are deleted during repair, SSA is considered a rapid but inherently mutagenic repair mechanism⁶⁶.
Microhomology-mediated end joining (MMEJ)
Microhomology-mediated end joining (MMEJ), also referred to as alternative NHEJ (Alt-NHEJ), is another error-prone DSB repair pathway⁶⁷. In MMEJ, short microhomologous sequences located near DNA break ends are used to align broken DNA termini prior to repair. Following recognition of these microhomologous regions, DNA ends are joined through a recombination-like process⁶⁸. Because MMEJ relies on very short homologous sequences, deletions and genomic rearrangements commonly occur during repair.
NHEJ efficiency can be quantitatively evaluated using EJ5-GFP reporter systems in reporter gene assays. Alternatively, repair activity may be indirectly assessed through Western blotting and immunofluorescence analysis of NHEJ-associated proteins.

8. Interstrand crosslink (ICL) repair

Interstrand crosslinks (ICLs) are highly toxic DNA lesions that covalently link the two strands of the DNA double helix, thereby blocking DNA replication and transcription. ICL repair is mechanistically complex and primarily involves the Fanconi anemia (FA) pathway and the NEIL3 pathway⁶⁹.

In the FA pathway, FANCM recognizes ICL lesions and promotes monoubiquitination of the FANCI-FANCD2 complex, thereby activating downstream repair proteins⁷⁰. Structure-specific nucleases subsequently incise DNA near the crosslink to separate the two DNA strands, and the resulting DSBs are repaired through HR or NHEJ pathways.

In the NEIL3 pathway, the DNA glycosylase NEIL3 cleaves glycosidic bonds associated with the ICL lesion, generating AP sites that are subsequently bypassed by translesion synthesis (TLS) polymerases⁷¹. Because ICLs prevent strand separation required for replication and transcription, these lesions are highly cytotoxic and contribute to aging, neurodegeneration, and cancer⁷²˒⁷³.

ICL repair efficiency can be assessed using plasmid-based repair assays, reporter gene assays, host cell reactivation assays, and in vitro reconstituted repair systems.

9. Translesion synthesis (TLS)

Translesion synthesis (TLS) is a DNA damage tolerance mechanism that enables replication to continue across unrepaired DNA lesions⁷⁴. TLS primarily involves specialized DNA polymerases, including polymerases κ, η, ι, and ζ. When DNA replication stalls at damaged sites, TLS polymerases bypass the lesion and allow replication to proceed, thereby maintaining genome stability⁷⁵.

Unlike BER, NER, MMR, HR, and NHEJ, which directly remove or repair DNA lesions, TLS does not eliminate DNA damage. Instead, it bypasses unrepaired lesions during DNA replication and is therefore classified as a damage tolerance mechanism rather than a canonical DNA repair pathway⁷⁶.

TLS can be divided into error-free and error-prone bypass pathways. Error-free TLS inserts the correct nucleotide opposite the lesion, whereas error-prone TLS frequently introduces incorrect nucleotides and contributes to mutagenesis. Error-prone TLS is considered a major mechanism by which environmental carcinogens induce genomic mutations. Consequently, modulation of error-prone TLS pathways may represent a potential strategy for cancer prevention and therapy⁷⁷.

Methods for assessing the efficiency of TLS

Isolation of Proteins On Nascent DNA (iPOND)
Isolation of proteins on nascent DNA (iPOND) is a technique used to analyze proteins associated with newly synthesized DNA at stalled replication forks. Because TLS activity frequently occurs at stalled forks, iPOND enables dynamic analysis of recruitment and assembly of TLS polymerases and DNA damage response (DDR) proteins during replication stress. This method can therefore be used to evaluate TLS activity and replication-coupled repair processes.
TLS efficiency may also be evaluated using reporter plasmids containing TLS-specific DNA lesions in reporter gene assays. In addition, Western blotting and immunofluorescence assays targeting TLS-associated proteins may provide indirect assessment of pathway activity.

10. Quality control and reproducibility considerations in experimental design

Biological replication and technical replication
Clear distinction between biological replicates and technical replicates is essential for ensuring reliability and reproducibility in DNA repair studies. In general, at least three independent biological replicates should be included for each experiment, whereas the number of technical replicates should be determined according to the variability of the detection platform.
For single-cell assays, such as the Comet assay and immunofluorescence-based foci analysis, at least 50–100 cells should be analyzed per biological replicate to minimize sampling bias. In flow cytometry-based reporter assays, including DR-GFP and FM-HCR assays, a minimum of 10,000 live-cell events should be collected per sample to ensure statistical robustness.
Standardization strategy
Accurate quantitative comparison of repair efficiency across different samples or experimental batches requires rigorous normalization procedures to minimize systematic variation arising from differences in transfection efficiency, cell viability, sample loading, and instrument performance. Common normalization strategies include the following:
1. Co-transfection of normalization plasmids
  In HCR assays, reporter-based repair assays, and plasmid-based repair assays, co-transfection with a normalization plasmid unaffected by DNA damage is recommended. Commonly used controls include pRL-TK Renilla luciferase plasmids or constitutively expressed GFP vectors. Repair efficiency is normalized by calculating the ratio between the experimental reporter signal and the control reporter signal.
2. Normalization using housekeeping proteins or total protein staining
  In Western blot analyses assessing repair protein expression or residual CPD levels, normalization should be performed using housekeeping proteins, such as GAPDH, β-actin, or histone H3, or through total protein staining methods, including Ponceau S or REVERT staining.
3. Undamaged controls
  In qPCR-based repair assays, undamaged genomic regions should be co-amplified as internal controls, and relative damage levels should be calculated using the ΔCt method. Similarly, HCR assays should include undamaged plasmid controls representing maximal reporter expression, allowing repair efficiency to be expressed relative to undamaged reference plasmids.
4. Cell number normalization
  In flow cytometry-based assays, including FM-HCR and DR-GFP assays, equal numbers of viable cells should be analyzed across samples. Repair efficiency should subsequently be reported as the percentage of positive cells or relative changes in fluorescence intensity.
Kinetic dimension of repair efficiency measurement
DNA repair is a dynamic process, and measurements obtained from a single time point may fail to distinguish between delayed repair and complete pathway deficiency. Therefore, time-course experimental designs are strongly recommended.
Following induction of DNA damage, samples should be collected at multiple sequential time points, such as 0, 0.5, 1, 2, 4, 8, 12, and 24 h, to monitor the accumulation and clearance of damage markers or repair products. The kinetics of signal disappearance provide important information regarding repair rate, pathway integrity, and overall repair efficiency.
Multiple hypothesis testing correction:
High-throughput and multiplexed approaches frequently involve simultaneous analysis of multiple repair pathways or experimental conditions, thereby increasing the risk of false-positive findings due to multiple statistical comparisons. Examples include simultaneous assessment of BER, NER, HR, and NHEJ activities using FM-HCR assays; genome-wide DSB analysis using BLESS or BLISS; and qPCR array-based analysis of DNA repair gene expression.
Accordingly, statistical correction methods for multiple hypothesis testing should be predefined during experimental design. The selected correction methods and adjusted significance thresholds should be clearly reported in the Materials and Methods and statistical analysis sections.

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Conclusions

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Accurate detection and quantitative analysis of DNA damage repair processes are essential for understanding the molecular mechanisms underlying DNA repair, evaluating the biological effects of environmental exposures, and assessing therapeutic responses⁷⁸. Therefore, in-depth investigation of DNA damage repair pathways and repair efficiency has substantial significance in both basic research and clinical applications⁷⁹.

Numerous studies have demo...

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Disclosures

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The authors declare no competing financial interests or personal relationships that could have influenced the work reported in this study.

Acknowledgements

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This work was supported by the Jiangsu Province Engineering Research Center of Development and Translation of Key Technologies for Chronic Disease Prevention and Control; the Jiangsu Higher Education Institution Innovative Research Team for Science and Technology (2021); the Program of Jiangsu Vocational College Engineering Technology Research Center (2023); the Natural Science Key Foundation of the Jiangsu Higher Education Institutions of China (Grant No. 24KJA310008); the Key Program of Suzhou Vocational Health College (Grant No. szwzy202406); and the State Key Laboratory of Radiation Medicine and Protection, Soochow University (Grant No. GZK1202506).

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Research Progress on DNA Damage Repair Efficiency Assessment Strategies

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