Modern proximity labeling techniques have enabled significant advances in understanding biomolecular interactions. However, current tools primarily utilize activation modes that are incompatible with complex biological environments, limiting our ability to interrogate cell- and tissue-level microenvironments in animal models. Here, we report μMap-Red, a proximity labeling platform that uses a red-light-excited SnIV chlorin e6 catalyst to activate a phenyl azide biotin probe. We validate μMap-Red by demonstrating photonically controlled protein labeling in vitro through several layers of tissue, and we then apply our platform in cellulo to label EGFR microenvironments and validate performance with STED microscopy and quantitative proteomics. Finally, to demonstrate labeling in a complex biological sample, we deploy μMap-Red in whole mouse blood to profile erythrocyte cell-surface proteins. This work represents a significant methodological advance toward light-based proximity labeling in complex tissue environments and animal models.

Adenosine-to-inosine (A-to-I) RNA editing is a widespread and conserved post-transcriptional modification, producing significant changes in cellular function and behavior. Accurately identifying, detecting, and quantifying these sites in the transcriptome is necessary to improve our understanding of editing dynamics, its broader biological roles, and connections with diseases. Chemical labeling of edited bases coupled with affinity enrichment has enabled improved characterization of several forms of RNA editing. However, there are no approaches currently available for pull-down of inosines. To address this need, we explore acrylamide as a labeling motif and report here an acrylamidofluorescein reagent that reacts with inosine and enables enrichment of inosine-containing RNA transcripts. This method provides improved sensitivity in the detection and identification of inosines toward a more comprehensive transcriptome-wide analysis of A-to-I editing. Acrylamide derivatization is also highly generalizable, providing potential for the labeling of inosine with a wide variety of probes and affinity handles.

Creating accurate maps of A-to-I RNA editing activity is vital to improving our understanding of the biological role of this process and harnessing it as a signal for disease diagnosis. Current RNA sequencing techniques are susceptible to random sampling limitations due to the complexity of the transcriptome and require large amounts of RNA material, specialized instrumentation, and high read counts to accurately interrogate A-to-I editing sites. To address these challenges, we show that Escherichia coli Endonuclease V (eEndoV), an inosine-cleaving enzyme, can be repurposed to bind and isolate A-to-I edited transcripts from cellular RNA. While Mg2+ enables eEndoV to catalyze RNA cleavage, we show that similar levels of Ca2+ instead promote binding of inosine without cleavage and thus enable high affinity capture of inosine in RNA. We leverage this capability to demonstrate EndoVIPER-seq (Endonuclease V inosine precipitation enrichment sequencing) as a facile and effective method to enrich A-to-I edited transcripts prior to RNA-seq, producing significant increases in the coverage and detection of identified editing sites. We envision the use of this approach as a straightforward and cost-effective strategy to improve the epitranscriptomic informational density of RNA samples, facilitating a deeper understanding of the functional roles of A-to-I editing.

Controlling the structure and activity of nucleic acids dramatically expands their potential for application in therapeutics, biosensing, nanotechnology, and biocomputing. Several methods have been developed to impart responsiveness of DNA and RNA to small-molecule and light-based stimuli. However, heat-triggered control of nucleic acids has remained largely unexplored, leaving a significant gap in responsive nucleic acid technology. Moreover, current technologies have been limited to natural nucleic acids and are often incompatible with polymerase-generated sequences. Here we show that glyoxal, a well-characterized compound that covalently attaches to the Watson–Crick–Franklin face of several nucleobases, addresses these limitations by thermoreversibly modulating the structure and activity of virtually any nucleic acid scaffold. Using a variety of DNA and RNA constructs, we demonstrate that glyoxal modification is easily installed and potently disrupts nucleic acid structure and function. We also characterize the kinetics of decaging and show that activity can be restored via tunable thermal removal of glyoxal adducts under a variety of conditions. We further illustrate the versatility of this approach by reversibly caging a 2′-O-methylated RNA aptamer as well as synthetic threose nucleic acid (TNA) and peptide nucleic acid (PNA) scaffolds. Glyoxal caging can also be used to reversibly disrupt enzyme–nucleic acid interactions, and we show that caging of guide RNA allows for tunable and reversible control over CRISPR-Cas9 activity. We also demonstrate glyoxal caging as an effective method for enhancing PCR specificity, and we cage a biostable antisense oligonucleotide for time-release activation and titration of gene expression in living cells. Together, glyoxalation is a straightforward and scarless method for imparting reversible thermal responsiveness to theoretically any nucleic acid architecture, addressing a significant need in synthetic biology and offering a versatile new tool for constructing programmable nucleic acid components in medicine, nanotechnology, and biocomputing.

Straightforwardm ethods for detecting adenosine-to-inosine (A-to-I) RNA editinga re key to ab etter understanding of its regulation, function, and connection with disease. Wea ddress this need by developing an ovel reagent, N-(4-ethynylphenyl)acrylamide (EPhAA), and illustrating its ability to selectively label inosine in RNA. EPhAA is synthesized in as ingle step, reacts rapidlyw ith inosine, and is "click"-compatible, enabling flexible attachment of fluorescent probesa te diting sites. We first validate EPhAA reactivity and selectivityf or inosine in both ribonucleosides and RNA substrates, and then apply our approach to directly monitor in vitro A-to-I RNA editing activity using recombinantA DAR enzymes. This method improves upon existing inosine chemical-labeling techniques and provides ac ost-effective, rapid, and non-radioactive approach for detecting inosine formation in RNA. We envision this method will improve the study of A-to-I editing and enable better characterization of RNA modification patterns in different settings. RNA is chemically modified by anumber of enzymes after transcription, in turn influencing RNA stability, localizationa nd activity within the cell. Adenosine-to-inosine (A-to-I) RNA editing is one of the most widespread modifications, and is performed by adenosine deaminases acting on RNA (ADARs) (Scheme 1a). [1] Adenosine deaminationc hanges the molecular structurea nd hydrogen-bonding pattern of the nucleobase, and resulting inosines insteadb ase pairw ith cytidinet oe ffectively recode these sites as guanosine. Editings ites within protein-coding mRNAsd irectly alter amino acid sequences and produce different protein isoforms. Non-coding RNAs also undergo extensive editing, including microRNAs and small-interfering RNAs, significantly alteringt heir biosynthesis, localization, and gene regulation properties. [2-3] A-to-I editing is essential for an umber of biological processes including tissue development, [4-5] neurologicalf unction, [6] and immune system activation. [7] Dysfunctional editing is also directlyl inked with autoimmune diseases, [8-9] neurological disorders, [10] and several types of cancer. [11-12] Despite this importance,o ur overall understanding of A-to-I editingr egulation is limited. In particular,w hile many sites have been identified (> 5million), [13-14] it is unclear why certain sites are edited at higherf requency than others and what precise function they each serve. [15] Efforts to map A-to-I locations and ADAR binding sites have revealed that editingp atterns are highly complex and variable in humans, [7, 16-18] and the precise mechanismsb yw hich ADAR enzymes bind to and edit specific RNA sequences remain unclear.T his gap is also significant for therapeutic site-directed RNA editing strategies, [19] as both the design and precise implementation of this machinery is reliant on at horough understanding of ADAR regulation. Detecting inosine formation in RNA is of central importance for characterizing editingm echanisms. While high-throughput RNA sequencin...