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

The story begins in 1969, when Robert Roeder and William Rutter first separated multiple forms of DNA-dependent RNA polymerase from eukaryotic cells. Over the following decades, biochemists painstakingly identified the general transcription factors, TFIID, TFIIB, TFIIE, TFIIF, and TFIIH, that guide polymerase II to promoters and position it precisely at the transcription start site. The TATA box, discovered in histone genes in the late 1970s, became the archetypal promoter element, and the crystal structure of the TATA-box-binding protein bound to DNA in 1993 revealed how this saddle-shaped factor bends the double helix nearly ninety degrees. But these early pictures were static snapshots of isolated parts. What the machinery does as a whole, and how it moves, remained opaque.

Cryo-electron microscopy changed everything. Where X-ray crystallography demanded rigid, homogenous crystals that giant, flexible assemblies rarely form, cryo-EM can capture heterogeneous, dynamic complexes frozen in vitreous ice, and computational classification can sort millions of particle images into distinct conformational states. Structures of the human pre-initiation complex, or PIC, the mega-assembly of polymerase II and general transcription factors on promoter DNA, progressed from low-resolution models to structures resolved at near-atomic detail. In 2021, researchers resolved the polymerase II pre-initiation complex at 2.9 angstroms, revealing the initial stages of DNA opening, and subsequent work captured the PIC in complex with the Mediator co-activator, with TFIIH, and even with the first nucleosome downstream of the promoter.

At the heart of the new understanding is TFIID, a massive 1.2-megadalton complex of the TATA-box-binding protein and thirteen TBP-associated factors, or TAFs. TFIID serves as the primary scaffold for promoter recognition. In its free state, it adopts a compact, autoinhibited conformation in which the lobes that will grip promoter DNA are folded inward. Binding of TBP to the TATA box triggers a dramatic rearrangement, and recent structures show that TFIID engages not only the TATA box but also downstream promoter elements, including the downstream core promoter region, or DPR, discovered through machine-learning analysis of human promoter sequences. Because the majority of human promoters lack a TATA box altogether, these TAF-mediated contacts with downstream DNA are essential for accurate initiation across most of the genome, and different TAF subunits are now known to dictate promoter selectivity and to integrate signals from sequence-specific activator proteins.

Superimposed on this scaffold is Mediator, the thirty-subunit co-activator that physically bridges enhancer-bound transcription factors to the promoter-bound machinery. Structural studies of the complete Mediator-PIC complex, including landmark human structures published in 2021, revealed how Mediator’s head, middle, and tail modules wrap around the polymerase and how its dissociable kinase module can regulate the interaction. The Pol II C-terminal domain, an intrinsically disordered tail of heptad repeats extending from the largest polymerase subunit, threads into the PIC-Mediator assembly, and its phosphorylation by the TFIIH-associated kinase module, through cyclin-dependent kinase 7, marks a critical checkpoint. Serine 5 phosphorylation of the CTD not only helps trigger promoter escape but also recruits the mRNA capping enzyme, coupling the earliest steps of transcription to RNA processing.

Perhaps the most striking theme to emerge from recent structural work is the role of chromatin itself. The +1 nucleosome, the first nucleosome downstream of the transcription start site, is no longer viewed as a mere obstacle. Structures of the +1 nucleosome-bound PIC-Mediator complex show that this histone package acts as a position-dependent regulator, physically orienting the pre-initiation complex, influencing transcription start-site selection, and even enforcing directionality. Epigenetic marks matter here: trimethylation of histone H3 lysine 4 anchors TFIID through the bromodomain of TAF3, while acetylation marks recruit BET-family factors that bridge TFIID and nucleosomes. The nucleosome, in other words, transmits epigenetic information directly into the transcription machinery, helping the PIC integrate genetic sequence and chromatin state into a single decision about where and in which direction to initiate.

The transition from initiation to elongation emerges as a choreographed, programmed process rather than a simple handoff. Before productive synthesis begins, polymerase II must melt the promoter DNA, a task performed primarily by the XPB translocase subunit of TFIIH, which pulls double-stranded DNA into the polymerase cleft to generate torsional stress. Early attempts at synthesis are often abortive: the enzyme repeatedly synthesizes and releases short RNAs of fewer than five nucleotides because the nascent transcript is thermodynamically unstable in the complex. During this phase, the enzyme scrunches downstream DNA into its active site, storing mechanical energy. Structural and single-molecule studies now delineate a three-step mechanism of promoter escape, in which steric pressure from accumulated template DNA and forward translocation of the RNA-DNA hybrid rupture the narrow channel formed by Pol II and TFIIB, causing general transcription factors to dissociate, the transcription bubble to collapse from roughly twenty-one to a stable eleven nucleotides, and the polymerase to commit to elongation, typically after transcribing about eight to twenty-three nucleotides.

Crucially, the review frames transcription initiation as a process governed by surface competition. Each stage of initiation involves factors that bind overlapping or competing surfaces on polymerase II. As CTD phosphorylation proceeds, inhibitory factors are released and regulatory factors are progressively exchanged: Mediator dissociates, DSIF and NELF bind to impose promoter-proximal pausing, and elongation factors await release signals. This model explains how the same binding surfaces can recruit different partners at different times, and how re-initiation, in which scaffold factors remain promoter-bound to launch successive rounds of polymerase loading, can generate the transcriptional bursting observed in single cells. It also rationalizes why paused polymerase can inhibit new initiation, creating feedback loops that shape gene expression dynamics.

The medical implications are substantial. TAF subunits such as TAF1 and TAF10 are required during development but dispensable in certain adult tissues, and PROTAC-mediated degradation of TAF1 has been shown to induce apoptosis in acute myeloid leukemia cells, making components of the initiation machinery emerging drug targets. As cryo-EM continues to push toward imaging these assemblies inside cells, and as single-molecule and nascent-RNA sequencing methods add temporal resolution, the field is converging on a unified view: transcription initiation is not a static lock-and-key event but a dynamic, chromatin-informed, programmed pipeline, and seeing it in atomic detail is finally telling us how life reads its own blueprint.

Subject of Research: Structural and mechanistic basis of transcription initiation by RNA polymerase II in eukaryotes

Article Title: The molecular basis of transcription initiation by RNA polymerase II

Article References: Chen, X., & Xu, Y. (2026). The molecular basis of transcription initiation by RNA polymerase II. Nature Reviews Molecular Cell Biology. https://doi.org/10.1038/s41580-026-01019-2

Image Credits: AI Generated

DOI: 10.1038/s41580-026-01019-2

Keywords: RNA polymerase II, transcription initiation, pre-initiation complex, TFIID, Mediator, cryo-electron microscopy, TATA box, promoter escape, plus-one nucleosome, CTD phosphorylation, TFIIH, gene regulation