Aaron M. Fleming

Ribosomal RNA has a high density of epitranscriptomic modifications essential for faithful translation, and their levels vary across cell types and in disease. Typically, rRNA modifications are quantified in bulk, and therefore, coordination on individual RNA molecules has remained poorly understood. Herein, modification-aware, single-molecule nanopore sequencing enabled detection of the co-occurrence of rRNA modifications on individual transcripts, resolving coordination invisible to ensemble methods. An analytical framework was established that separates co-occurrence from read-quality, false-positive, calling-artifact, near-saturation, and global modification-level confounds, using human rRNAs from HEK293T cells as the test dataset. The method was applied to four human cell lines to find that co-occurrence is not generally explained by a shared small nucleolar RNA (snoRNA) guide; instead, coordinated modifications cluster locally, within ∼100 nucleotides and 35-40 angstroms in the folded ribosome. For one shared-guide pair, coordination increased as guide levels fell across cell lines, possibly indicating an all-or-nothing mode of modification per molecule under limiting guide availability. Further, the results revealed rRNA heterogeneity between the cells in overall modification levels. Finally, levofloxacin remodeled specific modification sites and their local co-occurrence networks, showing that the approach can reveal small-molecule perturbation of rRNA modification networks.

The iron-Fenton reaction in biology is influenced by diverse cellular molecules that coordinate with redox-active ferrous ion. Contemporary research proposed that hydroxyl radical or a ferryl species (HO^•/Fe=O²⁺) are the primary oxidants; however, this was challenged by the observation that physiological bicarbonate redirects the reaction to form carbonate radical anion (CO₃^•-). Questions remained regarding the roles of O₂ concentration, ascorbate, and iron speciation in CO₃^•- formation. Accordingly, in cellulo studies were conducted under physiological O₂ (∼25 μM) with ascorbate replenishment to monitor bicarbonate-dependent telomeric DNA damage. Under these conditions,. physiological bicarbonate (25 mM) yielded 2'-deoxyguanosine-specific oxidation consistent with CO₃^•- formation at a ratio exceeding 80:1 relative to HO^•/Fe=O²⁺. In parallel in vitro experiments using a plasmid nicking assay, the cellular low molecular weight (LMW) ultrafiltrate was used as the source of iron and its endogenous coordination partners; under physiological O₂, bicarbonate, and 500 nM H₂O₂ (the concentration required to produce detectable signal), the DNA damage profile was consistent with exclusive CO₃^•- formation, mirroring the cell culture result. A panel of iron complexes approximating the intracellular labile iron pool (5 μM) was examined: hexaaquo-ferrous ion, ferrous citrate, ferrous α-ketoglutarate, ferrous pyrophosphate, ferrous glutathione, and hemin. Of these, only hemin reproduced the bicarbonate-dependent CO₃^•- damage profile observed in cells with 100 nM H₂O₂ and 25 mM bicarbonate present. This finding was corroborated using a defined biomimetic metabolome in which hemin, ferrous ion, or their combination was tested; hemin consistently supported CO₃^•- as the dominant oxidant. Roles for the Udenfriend reaction (Fe(II), O₂, and reductant) and superoxide dismutase were also studied. Collectively, these results identify heme iron as a likely candidate to drive CO₃^•- formation via the bicarbonate iron-Fenton reaction to damage dG in DNA during endogenous oxidative stress.

Ribosomal RNA (rRNA) modifications are tuned to regulate protein synthesis; however, their temporal dynamics during oxidative or inflammatory stress remain poorly understood. Nanopore direct RNA sequencing using Dorado v5.2.0 modification-aware models for the data analysis was employed to map human rRNA epitranscriptomic marks in a cell line undergoing oxidative stress, inflammatory stress, or ferroptosis. Oxidative stress triggered a global trend of decreased modification occupancy in which six modifications shifted significantly over 48 h, particularly, 18S Ψ573 and 18S m⁶A1832. Conversely, inflammatory stress induced a complex response involving an acute pulse of hypermodifications at 28S Um1773 and 28S Ψ1779, for example, and chronic hypomodification at specific target sites (e.g., 18S Gm1328 and 28S Gm4228). In this work, the pseudouridine modifications 28S Ψ4296 and 28S Ψ4353 were identified as "universal stress markers" that decreased under all stressors studied, including ferroptosis. Mapping these changes onto the ribosome structure revealed that they reside in functional regions such as the decoding center and A-site finger, supporting a role in functional ribosome reprogramming during stress. Analysis of mitochondrial rRNA (mt-rRNA) revealed modification shifts within the peptidyl transferase center, suggesting a mechanism to attenuate mitochondrial translation during chronic stress. This work demonstrates that oxidative and inflammatory stress drive distinct, time-resolved remodeling of the human rRNA epitranscriptome and provides a framework for using rRNA modifications as biomarkers of cellular health during oxidative or inflammatory stress exposure.

In a short 11 nt dye-labelled oligonucleotide hybrid, we investigated the oxidation of a single guanine, G, to 8-oxo-7,8-dihydroguanosine, 8oG, a reaction proceeding via two reaction pathways. In single-molecule fluorescence traces, we were able to detect the conversion of G to 8oG and assign different reaction pathways. Reactions are initiated by a photoinduced electron transfer to the dye, creating a G radical cation, G^•+, which is in equilibrium with the neutral G radical, G(−H)^•. Results demonstrate that 8oG formation via water addition to G^•+ proceeds within less than the time resolution of 150 ms. The reaction of G(−H)^• with superoxide is more than 1000 times slower due to the formation of an intermediate with a lifetime of minutes. The overall yield of 8oG formation is low but increases considerably upon addition of dithiothreitol, thus preferring water addition over reactions depending on the presence of oxygen. Our results highlight the advantages of single-molecule real-time technology for the investigation of chemical reactions and reaction pathways. The method enables the direct observation of product formation comparable to fast kinetic methods, including flash photolysis and laser spectroscopy, but on much longer timescales. Time-resolved measurements over minutes to hours not only allow monitoring of reaction sequences that show the formation and decay of products and intermediates, but also the order of their appearance, information that is lost in classical analysis. Product yields can be obtained by counting events, in contrast to ensemble measurements that require considerable manipulation, during which intermediates may get lost.

The bacterial pore-forming toxin α-hemolysin has dimensions appropriate for capture and translocation of DNA strands in a single-molecule electrophoresis experiment. We used the nanopore’s properties to study G-quadruplex unfolding in the human telomere repeat sequence with and without the presence of DNA lesions introduced into either the GGG tracks of a potential G-quadruplex (via oxidation) or the TTA loops (via photodimerization). Different topological folds of the G-quadruplex could be distinguished by either their current-time signatures or by their unfolding rates when a dA25 tail was added. Another more compact four-stranded structure, the i-motif, was also studied and found to be exceptionally well folded and stable inside the nanopore cavity. Comparisons are made between early studies with a home-built nanopore device vs. the currently available instrument from Oxford Nanopore Technologies (ONT), showing that we approach, but do not yet achieve, single-molecule sequencing of DNA damage sites in human telomere repeats. These studies aid in our understanding of the structure and dynamics of non-canonically folded DNA, its behaviour in crowded environments that mimic intracellular conditions, and the ability to use nanopore sequencing to identify DNA damage sites in this oxidation-prone segment of the genome.

Nanopore direct RNA sequencing (DRS) is revolutionizing our ability to analyze the epitranscriptome to evaluate nucleoside modifications in both cellular and synthetic RNA. The process involves minimal handling of fragile RNA strands, one round of reverse transcription to provide a DNA:RNA duplex, and library preparation to directly read nucleotides with their modifications as they pass through a protein nanopore embedded in a membrane. Simultaneous sequencing of hundreds of strands on a chip provides unprecedented access to whole transcriptome information. A key advantage is the long-read length that permits, for example, operon-specific epitranscriptomics of ribosomal RNA modifications as a function of cellular stress. By analyzing the entire transcriptome, the interplay of different modifications on the same RNA, or the correlation of changes in different RNAs in the same cell type, can be monitored. This review presents several recent examples of the types of experiments that are suitable for nanopore DRS as well as some of the current challenges and future expectations.

When cellular inflammation yields destructive radicals, the tentacle hairpins of rRNA expansion sequences bear the brunt of the damage. We leverage nanopore sequencing to capture real-time oxidative and nitrosative base lesions in cellular RNA.

Apurinic/apyrimidinic endonuclease‐1 (APE1) is a repair enzyme that efficiently cleaves abasic (AP) site damage in duplex DNA. Reports of in vitro activity assays between APE1 and single‐stranded G‐quadruplex (ssG4) reveal significant decreases in the endonuclease activity. Here, we identify that the low yields observed represent cleavage of the noncanonical folds that did not adopt a complete G4 fold. This conclusion is supported through circular dichroism analysis and activity assays analyzing the cleavage rate, folding impact on cleavage, and product inhibition. Studies were performed on AP‐containing ssG4 and duplex‐embedded G4 scaffolds. The CD spectra of a non‐G4 containing potential quadruplex sequence reveal a noncanonical structure. APE1 can cleave an AP in these non‐G4 conformation(s) in high yields comparable to the preferred duplex substrate. There is direct evidence of decreasing APE1 activity with increasing G4 folding in ssG4 and duplex‐G‐quadruplex‐duplex (DGD) systems. Also observed is a positional dependency on yield in the non‐G4 DGD scaffolds, but not in the non‐G4 ssG4 scaffolds. In conclusion, our studies provide evidence that APE1 efficiently cleaves noncanonical conformations in G4‐like structures, highlighting the control of secondary structure on APE1 endonuclease activity.

Antimicrobial resistance (AMR) threatens global health; however, the molecular adaptations underlying resistance to emerging antibiotic classes remain poorly defined. Here, we applied long-read DNA and direct RNA nanopore sequencing and developed methods to deconvolute operon-specific epitranscriptomic changes. Together, this platform uncovered a previously unrecognized, operon-specific pathway of resistance in Staphylococcus aureus to the naphthyridone antibiotic A-692345. Genomic nanopore sequencing identified a single 23S rRNA mutation (T1732C) confined to one of the six rRNA operons (operon 2), which uniquely contains nine tRNA genes. RNA direct nanopore sequencing generated a comprehensive and updated rRNA modification map for S. aureus and revealed extensive remodeling of rRNA modifications in the resistant strain. Differentially incorporated modifications included pseudouridine, dihydrouridine, and 5-hydroxycytidine at functionally relevant positions within the ribosome. Upon mapping these epitranscriptomic changes, we noted they were operon specific. This likely gives rise to ribosome heterogeneity with potential for selective translation of stress-response genes that favor resistance. Collectively, these findings establish nanopore sequencing as a powerful platform for resolving coupled genomic and epitranscriptomic adaptations, providing molecular insight into how bacteria evolve resistance to antibiotics.

Telomere dysfunction and attrition are observed in cells subjected to oxidative or inflammatory stress, largely due to DNA damage. In this study, we applied a DNA glycosylase-assisted quantitative PCR (qPCR) assay to assess telomeric DNA damage in human cells exposed to defined sources of oxidative or inflammatory stress. As a consequence of the endogenous chemistry of these stressors being different, we were able to identify distinct types of telomere DNA damage that differentiate oxidative from inflammatory stress. By selecting lesion-specific DNA glycosylases before qPCR analysis, we determined that reactive oxygen species generated under physiological bicarbonate buffering during both oxidative and inflammatory stress primarily damaged 2′-deoxyguanosine (dG) residues. In contrast, inflammatory stress increased dG oxidation sites and nitrosative DNA damage, evidenced by deamination of 2′-deoxycytosine (dC) to 2′-deoxyuridine (dU) and 2′-deoxyadenosine (dA) to 2′-deoxyinosine (dI; hypoxanthine). During inflammation, telomeric DNA contained 1.5-fold more dU than dI. These nitrosative lesions require different DNA repair enzymes compared with dG oxidation products, and they may impact telomere structure differently. Our findings suggest that beyond dG oxidation, nitrosative DNA damage can be a major source of lesions in telomeres. The profile of telomere DNA lesions can serve as a biomarker to distinguish between oxidative and inflammatory stress.