Mitochondria perform diverse metabolic and signaling functions, yet how these activities are spatially organized within the mitochondrial network of cells remains poorly understood. Organelle contact sites are spatially restricted hubs that regulate mitochondrial metabolism, signaling, and dynamics, and are therefore well positioned to influence mitochondrial specialization. Investigation of contact site-associated mitochondrial populations has been hindered by a lack of methods to isolate these subpopulations. Here, we develop Organelle Contact-dependent Affinity Purification (ORCA), a workflow for the isolation and analysis of subpopulations of intact mitochondria and associated proteins defined by their organelle contacts. ORCA revealed distinct proteomes for mitochondria associated with the endoplasmic reticulum, lysosomes, peroxisomes, and the Golgi apparatus, demonstrating that organelle contacts define biochemically specialized mitochondrial populations. Focused analysis of Golgi-associated mitochondria showed enrichment of mitochondrial ribosomes and increased mitochondrial translation, revealing an unexpected role for Golgi-mitochondria contacts in regulating mitochondrial protein homeostasis. ORCA also identified the previously uncharacterized Golgi protein KIAA0930/GMO1 as an evolutionarily conserved regulator of oxidative phosphorylation at Golgi-mitochondria contacts. Together, our findings establish ORCA as a broadly applicable approach for investigating the spatial organization of intracellular organelles and reveal organelle contacts as key determinants of mitochondrial specialization.

The accuracy of crucial nuclear processes such as transcription, replication, and repair depends on the local composition of chromatin and the regulatory proteins that reside there. Understanding these DNA-protein interactions at the level of specific genomic loci has remained challenging due to technical limitations. Here, we introduce a method termed 'DNA O-MAP', which uses programmable peroxidase-conjugated oligonucleotide probes to biotinylate nearby proteins. We show that DNA O-MAP can be coupled with label-free or sample multiplexed quantitative proteomics, targeted chemical perturbations, and next-generation sequencing to quantify DNA-proximal proteins and DNA-DNA interactions at specific genomic loci in human and murine cells. Furthermore, we establish that DNA O-MAP is applicable to both repetitive and unique genomic loci of varying sizes, from kilobase HOX gene clusters to megabase alpha-satellite repeats, and that DNA O-MAP can measure proximal molecular effectors in a homolog-specific manner.

Understanding how cells compartmentalize their biomolecules into discrete structures remains one of the fundamental goals of cell biology. The proliferation of proximity labeling (PL) technologies has been invaluable toward this goal, enabling biochemical “dissection” of compartments that would be intractable to classical biochemical methods. While robust PL approaches have long been established for targeting proteins of interest, targeting nucleic acids—RNAs and genomic loci—has remained significantly more challenging. Here, we review recent advancements in the field that overcome this longstanding roadblock by using programmable DNA oligonucleotides to direct PL enzyme localization. These tools are already revealing new insights into the molecular architecture of cellular compartments that lie at the heart of gene expression. They also provide a foundation for developing a new generation of PL tools that exploit the modularity and programmability of oligonucleotide-based devices to enable precise spatiotemporal control at previously inaccessible targets and in challenging specimen types.

Deciphering RNA molecules' function and regulation requires an in-depth understanding of the myriad interactions these RNAs form within their cellular environment. In this review, we present a comprehensive overview of recent technological advances that collectively form a molecular toolkit for mapping the molecular environment of RNA. We discuss innovative RNA-centric methods designed to overcome long-standing challenges in capturing direct RNA-protein interactions in living cells. Additionally, we explore transformative proximity-labeling techniques that leverage engineered enzymes and chemical catalysts to spatially resolve the composition of RNA-associated microenvironments. By critically evaluating the strengths and limitations of these emerging methodologies, we highlight how they are reshaping our understanding of RNA function, from local binding events to the dynamic organization of RNA-scaffolded compartments. These advancements not only promise to elucidate the molecular grammar underlying RNA regulatory networks in unprecedented details but also pave the way for an integrative, system-level understanding of complex RNA-mediated cellular processes.

The lncRNA Xist silences gene expression by recruiting the protein SPEN through its 5'-proximal Repeat A domain. How Repeat A recruits SPEN, how SPEN enacts silencing, and why Repeat A is required for Xist biogenesis in addition to silencing remain unclear. We find that sequences in Repeat A critical for SPEN recruitment, silencing, and Xist biogenesis directly bind SR-rich splicing factors and not SPEN. SRSF1, one such factor, is necessary and sufficient for SPEN recruitment to Repeat A. SPEN and SR protein-binding motifs in Repeat A enable Xist 's association with many proteins, including those involved in m 6 A methylation, RNA turnover, and transcriptional elongation. Our results reveal an unexpectedly essential role for splicing factors in coordinating silencing by SPEN and suggest a unifying model for the origin of Repeat A, the function of SPEN, and their roles in Xist biogenesis and silencing.

RNA molecules form complex networks of molecular interactions that are central to their function and to cellular architecture. But these interaction networks are difficult to probe in situ. Here, we introduce Oligonucleotide-mediated proximity-interactome MAPping (O-MAP), a method for elucidating the biomolecules near an RNA of interest, within its native context. O-MAP uses RNA-fluorescence in situ hybridization-like oligonucleotide probes to deliver proximity-biotinylating enzymes to a target RNA in situ, enabling nearby molecules to be enriched by streptavidin pulldown. This induces exceptionally precise biotinylation that can be easily optimized and ported to new targets or sample types. Using the noncoding RNAs 47S, 7SK and Xist as models, we develop O-MAP workflows for discovering RNA-proximal proteins, transcripts and genomic loci, yielding a multiomic characterization of these RNAs’ subcellular compartments and new regulatory interactions. O-MAP requires no genetic manipulation, uses exclusively off-the-shelf parts and requires orders of magnitude fewer cells than established methods, making it accessible to most laboratories. Oligonucleotide-mediated proximity-interactome MAPping (O-MAP) enables precise multiomic characterization of biomolecular interaction networks at target RNAs. Distinct O-MAP workflows reveal RNA-adjacent proteins, transcripts and genomic loci.

Calcium ions play important roles in nearly every biological process, yet whole-proteome analysis of calcium effectors has been hindered by a lack of high-throughput, unbiased, and quantitative methods to identify protein-calcium engagement. To address this, we adapted protein thermostability assays in budding yeast, human cells, and mouse mitochondria. Based on calcium-dependent thermostability, we identified 2,884 putative calcium-regulated proteins across human, mouse, and yeast proteomes. These data revealed calcium engagement of signaling hubs and cellular processes, including metabolic enzymes and the spliceosome. Cross-species comparison of calcium-protein engagement and mutagenesis experiments identified residue-specific cation engagement, even within well-known EF-hand domains. Additionally, we found that the dienoyl-coenzyme A (CoA) reductase DECR1 binds calcium at physiologically relevant concentrations with substrate-specific affinity, suggesting direct calcium regulation of mitochondrial fatty acid oxidation. These discovery-based proteomic analyses of calcium effectors establish a key resource to dissect cation engagement and its mechanistic effects across multiple species and diverse biological processes.

'Ribo-organisms' of the primordial RNA World would have needed ribozymes that catalyze RNA replication. McRae, Wan, Kristoffersen et al. recently revealed how these RNA replicases might have functioned by solving the structure of an artificial polymerase ribozyme. This work illustrates how complex RNA structures evolve, with implications for the origins of life.

In situ hybridization (ISH) is a powerful tool for investigating the spatial arrangement of nucleic acid targets in fixed samples. ISH is typically visualized using fluorophores to allow high sensitivity and multiplexing or with colorimetric labels to facilitate covisualization with histopathological stains. Both approaches benefit from signal amplification, which makes target detection effective, rapid and compatible with a broad range of optical systems. Here, we introduce a unified technical platform, termed ‘pSABER’, for the amplification of ISH signals in cell and tissue systems. pSABER decorates the in situ target with concatemeric binding sites for a horseradish peroxidase-conjugated oligonucleotide, enabling the localized deposition of fluorescent or colorimetric substrates. We demonstrate that pSABER effectively labels DNA and RNA targets in cultured cells and FFPE specimens. Furthermore, pSABER can achieve fivefold signal amplification over conventional signal amplification by exchange reaction (SABER) and can be serially multiplexed using solution exchange. Therefore, by linking nucleic acid detection to robust signal amplification capable of diverse readouts, pSABER will have broad utility in research and clinical settings. pSABER combines the power of signal amplification by exchange reaction (SABER) with the deposition of fluorescent or colorimetric substrates by horseradish peroxidase to enable enhanced signals for in situ hybridization in cells and tissues.

Eukaryotic nuclei adopt a highly compartmentalized architecture that influences nearly all genomic processes. Understanding how this architecture impacts gene expression has been hindered by a lack of tools for elucidating the molecular interactions at individual genomic loci. Here, we adapt oligonucleotide-mediated proximity-interactome mapping (O-MAP) to biochemically characterize discrete, micron-scale nuclear neighborhoods. By targeting O-MAP to introns within the TTN pre-mRNA, we systematically map the chromatin loci, RNAs, and proteins within a muscle-specific RNA factory organized around the TTN locus. This reveals an unanticipated compartmental architecture that organizescis-andtrans-interacting chromosomal domains, including a hub of transcriptionally silenced chromatin. The factory also recruits dozens of unique RNA-binding and chromatin-scaffolding factors, including QKI and SAFB, along with their target transcripts. Loss of the cardiac-specific splicing factor RBM20—a master regulator of TTN splicing that is mutated in dilated cardiomyopathy—remodels nearly every facet of this architecture. This establishes O-MAP as a pioneering method for probing single-locus, microcompartment-level interactions that are opaque to conventional tools. Our findings suggest new mechanisms by which coding genes can "moonlight" in nuclear-architectural roles.

In situ hybridization (ISH) is a powerful tool for investigating the spatial arrangement of nucleic acid targets in fixed samples. ISH is typically visualized using fluorophores to allow high sensitivity and multiplexing or with colorimetric labels to facilitate co-visualization with histopathological stains. Both approaches benefit from signal amplification, which makes target detection effective, rapid, and compatible with a broad range of optical systems. Here, we introduce a unified technical platform, termed ‘pSABER’, for the amplification of ISH signals in cell and tissue systems. pSABER decorates the in situ target with concatemeric binding sites for a horseradish peroxidase-conjugated oligonucleotide which can then catalyze the massive localized deposition of fluorescent or colorimetric substrates. We demonstrate that pSABER effectively labels DNA and RNA targets, works robustly in cultured cells and challenging formalin fixed paraffin embedded (FFPE) specimens. Furthermore, pSABER can achieve 25-fold signal amplification over conventional signal amplification by exchange reaction (SABER) and can be serially multiplexed using solution exchange. Therefore, by linking nucleic acid detection to robust signal amplification capable of diverse readouts, pSABER will have broad utility in research and clinical settings.

Throughout biology, RNA molecules form complex networks of molecular interactions that are central to their function, but remain challenging to investigate. Here, we introduce Oligonucleotide-mediated proximity-interactome MAPping (O-MAP), a straightforward method for elucidating the biomolecules near an RNA of interest, within its native cellular context. O-MAP uses programmable oligonucleotide probes to deliver proximity-biotinylating enzymes to a target RNA, enabling nearby molecules to be enriched by streptavidin pulldown. O-MAP induces exceptionally precise RNA-localized in situ biotinylation, and unlike alternative methods it enables straightforward optimization of its targeting accuracy. Using the 47S pre-ribosomal RNA and long noncoding RNA Xist as models, we develop O-MAP workflows for unbiased discovery of RNA-proximal proteins, transcripts, and genomic loci. This revealed unexpected co-compartmentalization of Xist and other chromatin-regulatory RNAs and enabled systematic characterization of nucleolar-chromatin interactions across multiple cell lines. O-MAP is portable to cultured cells, organoids, and tissues, and to RNAs of various lengths, abundances, and sequence composition. And, O-MAP requires no genetic manipulation and uses exclusively off-the-shelf parts. We therefore anticipate its application to a broad array of RNA phenomena.

CRISPR-Display uses the S. pyogenes Cas9 protein to posttranscriptionally localize noncoding RNA (ncRNA) domains to any genomic site, by directly fusing these domains to the Cas9 sgRNA cofactor. This versatile technology enables numerous applications for interrogating natural chromatin-regulatory ncRNAs, or for utilizing artificial ncRNA and ribonucleoprotein (RNP) devices at individual chromatin loci. To achieve these, a successful CRISPR-Display experiment requires that chimeric sgRNA-ncRNA fusions are stably expressed and incorporated into Cas9 complexes, and that they retain their ncRNA "cargo" domains at the targeted genomic sites. Here, I describe a workflow for designing, building, and testing such chimeric sgRNA-ncRNA expression constructs. I detail strategies for choosing expression systems and sgRNA insertion topologies, for assaying the incorporation of sgRNA-ncRNA fusions into functional Cas9 complexes, and for surveying the activities of ncRNA domains at targeted genomic loci. This establishes an initial set of "best practices" for the design and implementation of CRISPR-Display experiments.

APEX is an engineered peroxidase that catalyzes the oxidation of a wide range of substrates, facilitating its use in a variety of applications from subcellular staining for electron microscopy to proximity biotinylation for spatial proteomics and transcriptomics. To further advance the capabilities of APEX, we used directed evolution to engineer a split APEX tool (sAPEX). A total of 20 rounds of fluorescence activated cell sorting (FACS)-based selections from yeast-displayed fragment libraries, using 3 different surface display configurations, produced a 200-amino-acid N-terminal fragment (with 9 mutations relative to APEX2) called "AP" and a 50-amino-acid C-terminal fragment called "EX". AP and EX fragments were each inactive on their own but were reconstituted to give peroxidase activity when driven together by a molecular interaction. We demonstrate sAPEX reconstitution in the mammalian cytosol, on engineered RNA motifs within a non-coding RNA scaffold, and at mitochondria-endoplasmic reticulum contact sites.

Imaging and chromatin capture techniques have provided important insights into our understanding of nuclear organization. A limitation of these techniques is the inability to resolve allele-specific spatiotemporal properties of genomic loci in living cells. Here, we describe an allele-specific CRISPR live-cell DNA imaging technique (SNP-CLING) to provide the first comprehensive insights into allelic positioning across space and time in mouse embryonic stem cells and fibroblasts. With 3D imaging, we studied alleles on different chromosomes in relation to one another and relative to nuclear substructures such as the nucleolus. We find that alleles maintain similar positions relative to each other and the nucleolus; however, loci occupy unique positions. To monitor spatiotemporal dynamics by SNP-CLING, we performed 4D imaging and determined that alleles are either stably positioned or fluctuating during cell state transitions, such as apoptosis. SNP-CLING is a universally applicable technique that enables the dissection of allele-specific spatiotemporal genome organization in live cells.

SummaryOne of the biggest surprises since the sequencing of the human genome has been the discovery of thousands of long noncoding RNAs (lncRNAs)1–6. Although lncRNAs and mRNAs are similar in many ways, they differ with lncRNAs being more nuclear-enriched and in several cases exclusively nuclear7,8. Yet, the RNA-based sequences that determine nuclear localization remain poorly understood9–11. Towards the goal of systematically dissecting the lncRNA sequences that impart nuclear localization, we developed a massively parallel reporter assay (MPRA). Unlike previous MPRAs12–15that determine motifs important for transcriptional regulation, we have modified this approach to identify sequences sufficient for RNA nuclear enrichment for 38 human lncRNAs. Using this approach, we identified 109 unique, conserved nuclear enrichment regions, originating from 29 distinct lncRNAs. We also discovered two shorter motifs within our nuclear enrichment regions. We further validated the sufficiency of several regions to impart nuclear localization by single molecule RNA fluorescencein situhybridization (smRNA-FISH). Taken together, these results provide a first systematic insight into the sequence elements responsible for the nuclear enrichment of lncRNA molecules.

The spatial organization of RNA within cells is a crucial factor influencing a wide range of biological functions throughout all kingdoms of life. However, a general understanding of RNA localization has been hindered by a lack of simple, high-throughput methods for mapping the transcriptomes of subcellular compartments. Here, we develop such a method, termed APEX-RIP, which combines peroxidase-catalyzed, spatially restricted in situ protein biotinylation with RNA-protein chemical crosslinking. We demonstrate that, using a single protocol, APEX-RIP can isolate RNAs from a variety of subcellular compartments, including the mitochondrial matrix, nucleus, cytosol, and endoplasmic reticulum (ER), with specificity and sensitivity that rival or exceed those of conventional approaches. We further identify candidate RNAs localized to mitochondria-ER junctions and nuclear lamina, two compartments that are recalcitrant to classical biochemical purification. Since APEX-RIP is simple, versatile, and does not require special instrumentation, we envision its broad application in a variety of biological contexts.