Now, a comprehensive critical review published in Molecular Biology Reports by Tianshun Xu, Yaoyang Shen, and colleagues at Ningbo University’s Health Science Center has taken stock of the entire toolbox that scientists use to measure how well human cells perform this vital repair job. The review, led by corresponding authors Geng Wang and Na Li, systematically compares the biochemical, cellular, and genome-wide methods currently available for functional NER analysis, examining what each technique actually measures, how it works, and where its blind spots lie. In doing so, it offers researchers a kind of field guide to an area of biology that has grown fragmented across decades of methodological innovation, and it arrives at a deceptively simple conclusion: no single assay tells the whole story.

To appreciate why measuring NER is so tricky, it helps to understand what the pathway actually does. NER operates through two intertwined branches. Global-genome NER, abbreviated GG-NER, patrols the entire genome, scanning for distortions caused by lesions such as the cyclobutane pyrimidine dimers and pyrimidine (6-4) pyrimidinone photoproducts generated by ultraviolet light. In this surveillance mode, the XPC-RAD23B complex, often assisted by the UV-damaged DNA-binding protein complex, serves as the primary damage sensor. The second branch, transcription-coupled NER, is triggered when RNA polymerase II stalls at a lesion blocking its progress along a gene. Stalled transcription is itself the alarm bell, recruiting factors such as CSB, CSA, and UVSSA that hand the problem over to the shared repair machinery. Both branches converge on a common core: the transcription factor IIH, with its XPB and XPD helicase subunits, verifies the damage and opens the DNA, XPA and RPA help position the incision nucleases XPF-ERCC1 and XPG, and the damaged stretch is cut out as a short oligonucleotide roughly 24 to 32 nucleotides long. The resulting gap is filled by repair synthesis and sealed by ligation.

Because the pathway unfolds in discrete stages, from damage recognition through dual incision, gap filling, and ligation, different assays capture different snapshots of the process. The Ningbo review organizes the field around exactly this principle, grouping methods by the endpoint they interrogate: excision-product formation, repair-associated DNA synthesis, transcriptional or reporter gene-expression recovery, lesion removal inferred from restored DNA amplifiability, and genome-wide mapping of damage and excision events. Each category carries its own technical logic, its own strengths, and its own interpretive caveats, and the review’s central message is that assay selection must be guided by the specific research question at hand, with complementary readouts providing the most complete picture of NER function.

At the biochemical end of the spectrum sit the in vitro excision assays, which trace their lineage to the pioneering work of Aziz Sancar’s laboratory and others who reconstituted human NER in cell-free extracts in the late 1980s and early 1990s. In these systems, damaged plasmid DNA is incubated with cell extracts or purified protein fractions, and the dual-incision products, the small excised oligonucleotides carrying the lesion, are detected by gel electrophoresis or immunological methods. The approach offers exquisite mechanistic resolution: researchers can pinpoint exactly which protein factors are required for each step, as demonstrated in landmark studies that defined the minimal set of human factors needed to reconstitute dual incision. Recent structural biology has pushed this mechanistic understanding even further, with cryo-electron microscopy revealing the pre-incision complexes and the scanning-to-incision switch in TFIIH-XPG that licenses repair. Yet the review is careful to note the limitations of cell-free systems. Extract-based assays strip away the chromatin context in which repair actually occurs in living cells, and studies going back decades have shown that NER is suppressed in reconstituted nucleosomes, meaning that what works beautifully in a test tube may not reflect the chromatin-wrapped reality of the nucleus.

Moving into living cells, the review highlights the classic unscheduled DNA synthesis assay, a technique with roots stretching back to 1964, when Rasmussen and Painter first reported evidence of DNA repair in cultured mammalian cells. Because repair synthesis inserts new nucleotides outside the normal S phase of the cell cycle, it was dubbed unscheduled, and measuring this incorporation, originally with radioactive thymidine and later with the click-chemistry-compatible analog EdU, became the gold-standard functional readout of global-genome NER for decades. The assay remains clinically relevant: it has been used for prenatal diagnosis of xeroderma pigmentosum, the devastating hereditary disorder in which NER defects cause extreme sun sensitivity and a thousands-fold elevated skin cancer risk, and for complementation analysis of patient cells. Modern refinements, including tyramide signal amplification, have made the assay non-radioactive and sensitive enough for single-cell quantification, and parallel measurements of recovery of RNA synthesis provide the complementary readout for transcription-coupled repair, which is the branch specifically defective in Cockayne syndrome and UV-sensitive syndrome.

Perhaps the most versatile family of cellular assays is host cell reactivation, or HCR, in which a reporter plasmid is deliberately damaged in vitro, for example by UV irradiation or treatment with the cigarette-smoke carcinogen benzo[a]pyrene diol epoxide, and then transfected into the cells under study. If the cells’ NER machinery repairs the plasmid, the reporter gene, historically chloramphenicol acetyltransferase and now typically luciferase or fluorescent proteins, switches back on, and the level of expression becomes a proxy for repair capacity. The elegance of the method lies in its scalability and its epidemiological reach: modified HCR assays have been used to measure DNA repair capacity in peripheral blood lymphocytes from large cohorts, revealing associations between reduced repair capacity and elevated risk of skin, lung, head and neck, and bladder cancers. The latest evolution, fluorescence multiplexed HCR, or FM-HCR, deploys barcoded reporter plasmids carrying different types of damage, each tagged with a distinct fluorescent protein, allowing simultaneous quantification of repair across multiple lesion types and multiple pathways in a single sample by flow cytometry. Recent large-scale applications have measured inter-individual variation in DNA repair capacity across healthy populations, opening the door to personalized assessments of genome maintenance.

The most technologically ambitious methods in the review are the genome-wide approaches, chief among them excision repair sequencing, or XR-seq, developed in Sancar’s laboratory. Rather than measuring repair indirectly, XR-seq captures the actual excised oligonucleotides that NER releases, maps them back to the reference genome by next-generation sequencing, and thereby produces single-nucleotide-resolution maps of where repair happened across the entire genome. The technique has generated striking biological insights: repair is faster in transcribed genes than in silent regions, it is modulated by chromatin state, it follows circadian rhythms in mouse tissues, and it varies systematically across the three-dimensional organization of the genome, as shown in recent work mapping the repair of aflatoxin-induced and cisplatin-induced damage. Complementary damage-mapping methods such as Damage-seq and high-sensitivity variants locate the lesions themselves, so that comparing damage maps with excision maps reveals repair kinetics directly. These approaches have even been extended beyond humans, illuminating repair patterns in plants, yeast, and bacteria, and connecting transcription-coupled repair to the Mfd translocase in E. coli.

Why does all this methodological scrutiny matter beyond the laboratory? The review emphasizes that NER sits at a fascinating double-edged intersection with cancer. On one hand, robust NER protects normal cells from mutagenesis, and epidemiological studies have linked weaker repair phenotypes to higher cancer risk. On the other hand, cancer cells can co-opt elevated NER activity to survive the DNA-damaging chemotherapy drugs that oncologists deploy against them. Cisplatin adducts and other bulky lesions are themselves NER substrates, so tumors with high repair capacity can shrug off treatment. This has fueled a therapeutic strategy of NER inhibition: the review’s authors themselves have previously shown that triptolide, a natural product, enhances carboplatin-induced apoptosis in melanoma by suppressing NER activity. Accurate, well-matched assays are therefore essential not only for understanding basic biology but also for identifying which tumors might respond to such combination therapies and for monitoring how interventions actually change repair function in cells.

The review’s final verdict is a call for methodological humility and deliberate design. Because each assay interrogates a distinct stage or consequence of NER, a positive result in one assay cannot be automatically generalized to the whole pathway, and apparent discrepancies between studies may simply reflect different endpoints rather than conflicting biology. Biochemical assays offer mechanistic precision but lack cellular context; cellular reporter assays capture functional outcomes but can be influenced by transcription, replication, and toxicity; genome-wide maps deliver unprecedented resolution but demand substantial sequencing resources and computational expertise. The authors argue that researchers should choose their tools to match their questions and, wherever possible, layer complementary readouts, for example pairing an excision assay with a repair-synthesis measurement or a genome-wide map with a functional reporter, to build a comprehensive assessment of NER capability. As DNA repair biology moves from correlation toward causation in cancer prevention, diagnosis, and treatment, that disciplined, multi-angle approach may prove to be the review’s most enduring contribution.

Subject of Research: Functional methods for assessing nucleotide excision repair capability in human cells

Article Title: Assessment and detection of nucleotide excision repair capability in human cells: a critical review of current functional methods

Article References: Xu, T., Shen, Y., Dai, X., Shao, Y., Zhang, Y., Zhang, Y., Wang, G., & Li, N. (2026). Assessment and detection of nucleotide excision repair capability in human cells: a critical review of current functional methods. Molecular Biology Reports, 53(1), Article 1635. https://doi.org/10.1007/s11033-026-12816-5

Image Credits: AI Generated

DOI: 10.1007/s11033-026-12816-5

Keywords: nucleotide excision repair, DNA damage, DNA repair assays, host cell reactivation, unscheduled DNA synthesis, XR-seq, xeroderma pigmentosum, transcription-coupled repair, genome stability, cancer therapy resistance, TFIIH, functional genomics

Interstrand crosslinks, or ICLs, are among the most cytotoxic forms of DNA damage because they covalently tether the two strands of the double helix to one another, blocking both DNA replication and transcription. Cisplatin, a platinum-based drug, induces these crosslinks as part of its antitumor arsenal, and the resulting suppression of transcription contributes significantly to the drug’s ability to kill rapidly dividing cancer cells. However, cells possess a dedicated pathway known as transcription-coupled repair that can remove such lesions from actively transcribed genes, and this repair capacity can mitigate cisplatin’s effects, contributing to treatment resistance in some tumors. Understanding exactly how the crosslink stalls the polymerase, and how that stall is recognized by the repair machinery, is therefore of direct clinical relevance.

The Zhejiang team assembled defined RNA polymerase II elongation complexes on DNA templates containing a site-specific cisplatin-induced interstrand crosslink, positioning the lesion at various distances from the enzyme’s active site. By vitrifying these complexes and imaging them with cryo-electron microscopy, the researchers were able to reconstruct a molecular movie of sorts, following the polymerase as it advanced toward the crosslink and ultimately ground to a halt. The eight captured states span the journey from the initial approach of the lesion through the final stalled configuration, offering a step-by-step structural narrative that has never before been available for this class of DNA damage.

The most striking discovery emerged when the polymerase reached the crosslink itself. Rather than simply blocking progress like a physical roadblock, the crosslinked guanine bases underwent an unexpected conformational transformation. The enzyme actively remodeled the crosslink, prying the damaged guanines into a novel configuration positioned above the bridge helix, a structural element of the polymerase that plays a central role in translocating DNA and RNA through the enzyme. This rearrangement had not been observed for any other type of transcription-blocking lesion and reveals RNA polymerase II in an unanticipated role: that of a structural remodeler of the very damage it encounters.

The consequence of this remodeling is functionally decisive. By relocating the crosslinked base above the bridge helix, the polymerase prevents the damaged guanine from entering the active site, the catalytic chamber where templated nucleotide addition normally occurs. With the lesion physically excluded from the position where it would need to reside to serve as a template, transcription cannot proceed in the conventional manner. The structural data show that the remodeled crosslink creates an empty nucleotide-binding site poised in an abnormal configuration, setting the stage for a secondary failure mode that the researchers characterized in detail.

That secondary failure is ATP misincorporation followed by backtracking. Because the active site is left in an aberrant state, the polymerase can misinsert adenosine triphosphate opposite the position where the crosslinked guanine should have been read. Following this misincorporation event, the enzyme undergoes backtracking, a process in which the polymerase slides backward along the DNA and RNA, extruding the freshly synthesized 3′ end of the RNA transcript from the active site. Backtracking is a well-known mechanism of transcriptional arrest, and the new structures demonstrate how the remodeled crosslink actively drives the enzyme into this arrested configuration rather than merely obstructing forward motion.

The study also examined oxaliplatin, another clinically important platinum chemotherapy drug, and found that its interstrand crosslinks stall polymerase II through a conserved mechanism. Notably, the structural analyses revealed that when the platinum crosslink sits several nucleotides downstream of the active site, the polymerase drives the platinum adduct from the minor groove of the DNA helix into the major groove, and the crosslinked guanine on the non-template strand migrates toward the upstream side of the complex as the lesion approaches the catalytic center. These observations, together with experiments using alternative DNA sequences, demonstrated that the remodeling behavior is not an artifact of a particular template but a general property of how polymerase II handles platinum crosslinks.

A central question in transcription-coupled repair concerns what happens after the polymerase stalls. Cockayne syndrome protein B, or CSB, is an ATP-dependent translocase that is recruited to stalled polymerase II and is considered the initiating factor of transcription-coupled repair. One might expect CSB to either help the polymerase bypass the lesion or actively displace the stalled enzyme to clear the way for repair enzymes. The new data show that CSB does neither. In the reconstituted system, CSB neither promoted bypass of the cisplatin crosslink nor displaced the stalled polymerase, and the transcription factor TFIIS, which can stimulate RNA cleavage and restart arrested polymerases, likewise failed to rescue transcription through the lesion. This failure to resolve the blockage effectively commits the crosslinked site to the transcription-coupled repair pathway, ensuring that the damage is flagged for processing rather than erroneously bypassed.

The structural comparisons revealed additional mechanistic elegance. The fork loop-2 region of the polymerase was observed to separate the template and non-template DNA strands at the downstream fork of the transcription bubble, and this separation appears to facilitate the migration of the crosslinked non-template guanine as the lesion is remodeled. Furthermore, alignment of the cisplatin-stalled complexes with previously determined structures of polymerase II bound to CSB homologs, including yeast Rad26 and the human CSB-CSA-UVSSA complex, showed that the crosslink-stalled enzyme adopts a conformation competent for recruiting these repair factors, mirroring the way polymerase II arrested by bulky lesions on the template strand presents itself to the repair machinery.

The implications of this work extend from basic molecular biology to the clinic. By identifying RNA polymerase II as an active remodeler of cisplatin-induced interstrand crosslinks and defining the structural basis of the resulting stall, the study fills a long-standing gap in our understanding of how platinum drugs exert their transcription-blocking effects at atomic resolution. Because the efficiency of transcription-coupled repair influences cellular sensitivity to cisplatin, with repair-deficient cells such as certain testis tumor cells showing heightened drug sensitivity, the molecular details uncovered here could inform strategies to modulate repair pathways and improve therapeutic outcomes. The cryo-electron microscopy maps and atomic models have been deposited in public databases, ensuring that the structural framework established by the Zhejiang team will serve as a foundation for future studies of crosslink repair, polymerase stalling, and the cellular response to platinum-based chemotherapy.

Subject of Research: Structural mechanism of transcription stalling by cisplatin-induced DNA interstrand crosslinks

Article Title: Transcription remodeling and stalling by cisplatin-induced interstrand crosslinks

Article References: Zhu, L., Zhao, W., Ye, L., Liu, Y., Deng, W., Qian, B., Yang, X., & Xu, J. (2026). Transcription remodeling and stalling by cisplatin-induced interstrand crosslinks. Nature Chemical Biology. https://doi.org/10.1038/s41589-026-02338-3

Image Credits: AI Generated

DOI: 10.1038/s41589-026-02338-3

Keywords: cisplatin, interstrand crosslink, RNA polymerase II, transcription-coupled repair, cryo-electron microscopy, DNA damage, Cockayne syndrome protein B, oxaliplatin, backtracking, platinum chemotherapy, bridge helix, DNA repair