Epitranscriptomic RNA modifications can regulate mRNA function; however, there is a major gap in our understanding of the biochemical mechanisms mediating their effects. Here, we develop a chemical proteomics approach relying upon photo-cross-linking with synthetic diazirine-containing RNA probes and quantitative proteomics to profile RNA-protein interactions regulated by N-methyladenosine (mA), the most abundant internal modification in eukaryotic RNA. In addition to identifying YTH domain-containing proteins and ALKBH5, known interactors of this modification, we find that FMR1 and LRPPRC, two proteins associated with human disease, "read" this modification. Surprisingly, we also find that mA disrupts RNA binding by the stress granule proteins G3BP1/2, USP10, CAPRIN1, and RBM42. Our work provides a general strategy for interrogating the interactome of RNA modifications and reveals the biochemical mechanisms underlying mA function in the cell.

Epitranscriptomic RNA modifications can serve as recognition elements for the recruitment of effector proteins (i.e., “readers”) to modified transcripts. While these interactions play an important role in mRNA regulation, there is a major gap in our understanding of the sequence determinants critical for the binding of readers to modified sequence motifs. Here, we develop a high-throughput platform, relying upon in vitro selection with a site-specifically modified random sequence RNA library and next-generation sequencing, to profile the binding specificity of RNA modification reader proteins. We apply our approach to interrogate the effect of sequence context on the interactions of YTH-domain proteins with N6-methyladenosine (m6A)-modified RNA. We find that while the in vitro binding preferences of YTHDC1 strongly overlap with the well-characterized DR­(m6A)­CH motif, the related YTH-domain proteins YTHDF1 and YTHDF2 can bind tightly to noncanonical m6A-containing sequences. Our results reveal the principles underlying substrate selection by m6A reader proteins and provide a powerful approach for investigating protein-modified RNA interactions in an unbiased manner.

Epitranscriptomic RNA modifications can regulate fundamental biological processes, but we lack approaches to map modification sites and probe writer enzymes. Here we present a chemoproteomic strategy to characterize RNA 5-methylcytidine (m5C) dioxygenase enzymes in their native context based upon metabolic labeling and activity-based crosslinking with 5-ethynylcytidine (5-EC). We profile m5C dioxygenases in human cells including ALKBH1 and TET2 and show that ALKBH1 is the major hm5C- and f5C-forming enzyme in RNA. Further, we map ALKBH1 modification sites transcriptome-wide using 5-EC-iCLIP and ARP-based sequencing to identify ALKBH1-dependent m5C oxidation in a variety of tRNAs and mRNAs and analyze ALKBH1 substrate specificity in vitro. We also apply targeted pyridine borane-mediated sequencing to measure f5C sites on select tRNA. Finally, we show that f5C at the wobble position of tRNA-Leu-CAA plays a role in decoding Leu codons under stress. Our work provides powerful chemical approaches for studying RNA m5C dioxygenases and mapping oxidative m5C modifications and reveals the existence of novel epitranscriptomic pathways for regulating RNA function.

Fluorescence imaging is a powerful method for probing macromolecular dynamics in biological systems; however, approaches for cellular RNA imaging are limited to the investigation of individual RNA constructs or bulk RNA labeling methods compatible primarily with fixed samples. Here, we develop a platform for fluorescence imaging of bulk RNA dynamics in living cells. We show that fluorescent bicyclic and tricyclic cytidine analogues can be metabolically incorporated into cellular RNA by overexpression of uridine−cytidine kinase 2. In particular, metabolic feeding with the tricyclic cytidine-derived nucleoside tC combined with confocal imaging enables the investigation of RNA synthesis, degradation, and trafficking at single-cell resolution. We apply our imaging modality to study RNA metabolism and localization during the oxidative stress response and find that bulk RNA turnover is greatly accelerated upon NaAsO 2 treatment. Furthermore, we identify cytoplasmic RNA granules containing RNA transcripts generated during oxidative stress that are distinct from canonical stress granules and P-bodies and co-localize with the RNA helicase DDX6. Taken together, our work provides a powerful approach for live-cell RNA imaging and reveals how cells reshape RNA transcriptome dynamics in response to oxidative stress.

A DNA-imidazole conjugate, designed to mimic enzyme function, site-selectively hydrolyzes a target ester, but not other esters, with >100-fold rate enhancement compared to free imidazole.