Biochemical Methods To Image and Analyze RNA Localization: From

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Biochemical Methods to Image and Analyze RNA Localization: from One to Many Ying Li, Ke Ke, and Robert C. Spitale Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b01087 • Publication Date (Web): 16 Nov 2018 Downloaded from http://pubs.acs.org on November 18, 2018

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Biochemical Methods to Image and Analyze RNA Localization: from One to Many Ying Li $,1, Ke Ke$,1, Robert C. Spitale*,1,2 $These authors contributed equally to this manuscript. (1) Department of Pharmaceutical Sciences and (2) Department of Chemistry. University of California, Irvine. Irvine, California. 92697 Supporting Information Placeholder

ABSTRACT: Recent analysis of transcriptomes has revealed that RNAs perform a myriad of functions beyond coding for proteins. Critical to RNA function is its transport to unique subcellular locations, where it can be spatially regulated to perform its biological function. Within this perspective we highlight the techniques developed to image and discover RNAs within their subcellular compartments, and discuss the utility of these techniques in understanding RNA localization.

Understanding how a cell organizes its molecular components is one of the great remaining challenges in basic and medical research. Proteins were once thought to be the only molecules with specific localization properties. Nevertheless, since the mid-1980s, an overwhelming amount of evidence has been gathered which suggests that cells localize proteins by directing the corresponding messenger RNAs (mRNAs) .1 The localization of RNAs to subcellular compartments provides a mechanism for regulating gene expression with exquisite temporal and spatial control. Localization of RNAs is widespread and evolutionarily conserved. 2-5 In Saccharomyces cerevisiae a subset of mRNAs is selectively localized during cell budding, ensuring that certain proteins are only expressed in the daughter cell.6 Fluorescent in situ hybridization (FISH) in the developing Drosophila embryo revealed that 71% of 3370 genes analyzed encode subcellularly localized mRNAs.7 RNA localization is a hallmark of neurobiology, where mRNAs can be targeted to specific neuronal subcellular domains enabling rapid changes in the spatial proteome through local translation.8 In many cases RNA localization pre-determines protein synthesis and localization, suggesting that RNA localization can be a predictor of protein placement and organization.9 Misregulated RNA localization can lead to many disease phenotypes.2-5 In metastatic breast cancer cells specific mRNAs have been localized in cellular protrusions, where their protein products are locally translated to control kinase signaling pathways.4 Gprotein receptor translation and membrane insertion is controlled by mRNA localization to the plasma membrane.10 Altered mRNA axonal transport has been

found in spinal muscular atrophy and has been hypothesized to be causative for dysfunction and degeneration of motorneurons.11 These examples are just a few of the potentially vast roles in which the localization of RNAs can contribute to disease. The majority of work in the field has focused on the localization of mRNAs; nevertheless, long non-coding RNAs (lncRNAs) are often subject to specific subcellular localization as well. lncRNAs are a relatively new class of described RNAs, but they play important roles in disease and development.12 Elucidating where and why RNAs are localized to specific cellular compartments in a more comprehensive manner would be extremely empowering to research focused on RNA biology and potentially answer longstanding biological questions.

Figure 1: The lifetime of an RNA and changes in localization as it moves through the cell, from birth to decay. From its birth to decay, the localization of an RNA can change dramatically (Figure 1). Initially, mRNAs and some non-coding RNAs are transcribed and processed in the nucleus.13 Second, protein complexes can form to shepherd RNAs through the nuclear pore and out into the cytoplasm.14 There, recognition of an RNA motif by an RNA-binding protein (RBP) connects the RNA with the cytoskeletal network.15 These RNAprotein interactions present a molecular “code” for where an RNA is spatially organized. RNAs can then be trafficked great distances within cells to unique organelles (mitochondria, ER, membrane, even back into the nucleus). Upon correct localization RNAs can

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perform their functions. For example, mRNAs can be translated and their protein products are imported into the mitochondria.16 ncRNAs can bind to chromatin modifying enzymes to control gene expression.17 mRNAs can be trafficked to the ER to be translated and protein products are imported into the ER lumen.18 mRNAs can be asymmetrically divided in cells to ensure proper localization of protein translation to regulate signaling networks. 19 These examples highlight just some of the prevailing paradigms we know of in RNA localization. Despite decades studying RNA localization, many fundamental questions need to still be further explored to better understand how large classes of RNAs are localized and what their contribution to biological regulation is. What locations within the cell are “hot spots” for organized RNA localization? How do RNA localization, organization, and dynamics contribute to cellular fate decisions? What novel RNA sequence and structural motifs program RNA transport and localization? What factors are important for RNA spatial organization? An incomplete understanding of RNA localization is a key obstacle that impedes progress toward obtaining a holistic understanding of RNA biology and being able to design RNA-based medicines. Paramount to addressing these questions is understanding some of the techniques that are used to assay RNA localization within cells. Herein we detail more mature approaches, which are primarily focused on studying single RNA at a time. Further, we give perspective on the maturation of biochemical approaches to engineer cells to report on the localization of their RNAs. Lastly, we introduce some of the new methods being developed to discover localized RNAs near pointsof-interest in cells. Hybridization-Based Methods of RNA Detection The majority of focus on developing methods to analyze RNA localization, stems from a desire to know where single transcript is in cells. This is more often than not accomplished now through RNA imaging. Hybridization methods are ideal for this as they rely on the primary sequence of an RNA of interest to design antisense oligos for visualization.20 Once an RNA sequence of interest is chosen, antisense RNA, DNA, or other modified sequences can be designed to ensure robust base-pairing through Watson-Crick interactions. Fluorescence in situ hybridization (FISH; Figure 2, a)21 is the most commonly used method of choice since it is now sophisticated enough to distinguish between two RNA molecules that differ by a single nucleotide.22 In a typical FISH experiment, cells are grown to a desired confluency, fixed (formaldehyde or methanol), permeabilized, and then antisense oligos are incubated for extended periods of time to enable base pairing. Traditional FISH experiments have been widely utilized to understand transcriptional output, RNA localization,

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and changes in localization due to cellular stress or development.20 Although widely adopted FISH does struggle in some contexts due to background issues attributed to nonspecific binding of oligos or fluorophores to endogenous cellular components or crosslinkers23, extension of FISH has relied on sophisticated chemical approaches which only work when antisense oligos are hybridized to their RNA target of interest. One such strategy involves taking advantage of the ability of fluorophores to be quenched by electron-rich quencher moieties as molecular beacons (Figure 2, b). A molecular beacon24 consists of a DNA probe that covalently links a fluorophore with a quencher and the fluorescence is quenched. Once bound to the target the fluorophore and the quencher are separated by long enough distance to enable fluorescence. Many examples of molecular beacons exist in the literature.24 Using quencher-fluorophore pairs to suppress fluorescence results in significantly lowered fluorescence, but can still have significant background due to the breathing of distance between quencher and fluorophore. Pairing Förster resonance energy transfer (FRET) pairs with quenching helps to minimize this effect, because the reaction is controlled through pairing and the FRET donor-acceptor distance needs to be optimal for energy transfer (Figure 2, c). Hybridization bridging between two oligos can bring a nucleophile (Nu:) in close proximity to a tethered quencher to release the quencher upon binding. One of the early adoptions of this approach is nicely executed by utilizing thioateconjugated oligos (Nu:) and sulfo-dansyl quencher pairs that once hybridized release a dansyl quencher and enable Förster resonance energy transfer (FRET) for imaging of RNAs inside cells.25 Bioorthogonal templated reactions can also serve the role for controlling the removal of protecting groups on fluorophores by adjacent reactive groups to enable visualization (Figure 2, d). For example, localized Staudinger reduction of aryl azide moieties enables 1,6elimination reactions to liberate quenchers and enable fluorescence. Additionally, aryl azides can be reduced by hybridization-based proximity to phosphines that enable the formation of aryl amine fluorophores (such as rhodamine dyes).26 Another example is the utility of biorthogonal tetrazene reactivity to unmask vinyl ether caged fluorophores, including a near-infrared (NIR)emitting cyanine dye.27 Overall, these examples, and others nicely demonstrate the utility of proximity-driven reactions to remove protecting groups (or chemical moieties that limit fluorescence) to liberate fluorophores that are controlled by RNA target engagement through hybridization. Many different interactions of this approach are reported in the literature and are nicely summarized in the reference.28 Hybridization-based imaging of RNA localization has been utilized to understand the distribution of RNA molecules and how they change


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under stress, development, or disease. One such compelling utilization of FISH has been used to understand lncRNA localization within the nucleus. lncRNAs are a class of emergent RNAs, many of which function to control chromatin modifying enzymes or genome structure to regulate transcription. FISH has recently been used to drive hypotheses about RNA function by categorizing the spatial distribution of lncRNAs within cells, by characterizing their cytoplasmic : nucleus ratio. By systematically curating the localization of 61 lncRNAs, Rinn and Raj, were able to understand the distribution of lncRNAs on the chromatin, how their localization changed during the cell cycle, and also made comparisons to mRNAs.29 Further, two-colored FISH has been used to characterize lncRNA localization on the genome – marking sites of transcription through intronFISH and also the final destination of mature lncRNAs (exon-exon FISH; Figure 2, e).30 Utilizing FISH in this way has given a localization-based understanding of RNA function and trafficking around the nucleus. Single-cell FISH can reveal not only the spatial distribution of RNAs in cells, but also the variability in gene expression among cells of the same genetic background. A recent highlight of this approach comes from understanding transcriptional variability in human melanoma cells, which were found to display profound transcriptional variability at the single cell level that predicts which cells will ultimately resist drug treatment.31 This variability involves infrequent, semicoordinated transcription of a number of resistance markers at high levels in a very small percentage of cells. Identification of such “jackpot cells” (Figure 2, f) was made possible by single-cell FISH to characterize the expression of the RNAs and also their coordinated cytoplasmic localization leading to enhanced translation of drug resistance proteins. Lastly, the extensive pallet of fluorophores and conjugation chemistries have also enabled emerging methods for serial hybridization to obtain the localization of many RNAs in parallel. For example, in SeqFISH32 and MERFISH33, oligos with unique primary sequence identifiers and corresponding fluorophores are hybridized, then stripped away. Following stripping, a next set of oligos are hybridized with a new set of sequence and fluorophore identifiers. This process is repeated multiples times. Finally, the individual localizations of hybridization events are overlaid and the primary sequences of the oligos can be collated to give the identity of thousands of RNA molecules from a single experiment (Figure 2, g). Efforts combining hybridization-based WatsonCrick pairing and chemical approaches to reduce background and enhance signal have resulted in sophisticated methods to image RNAs within cells. As shown above, exploiting these efforts to gain additional insight into localization and expression variation has revealed how specialized placement and transcriptional variability can play in RNA function within cells.

Figure 2: Examples of RNA detection and localization by hybridization-based approaches. (a) Example of simple hybridization-based detection of RNA for FISH. (b) Schematic of a molecular beacon, whereby in the absence of the RNA binding partner fluorescence is quenched. (c) Schematic of hybridization-based nucleophilic attack to promote liberation of a quencher and enable FRET. (d) Hybridization-based proximity can promote removal of a protecting group (PG) by reactive group (RG) to enable fluorescence. (e) Cartoon depicting the use of two-colored FISH to image the site of transcription and the site of mature RNA localization, within the nucleus. (f) FISH can be used to image single cells and identify cells that have different RNA


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abundance or “bursting” of RNA expression. (g) Schematic of SeqFISH and MERFISH Biochemical methods to image RNA within living cells. Hybridization-based approaches are very useful for analyzing a static picture of RNA localization or expression. However, RNA localization is a dynamic process that requires analysis within living cells to create the complete picture. Toward this goal, a combination of sophisticated biological approaches, with live-cell compatible chemical methods is starting to expand our ability to image RNAs without the use of antisense oligos. The most widely used approach combines green fluorescent protein (GFP) imaging and the construction of an RNA aptamer in the 3’-UTR of the RNA of interest (Figure 3, a).34 MS2 RNA aptamer and a GFP-MS2 binding protein fusion has been applied in a variety of biological systems. This system has characterized RNA localization dynamics during yeast cell budding,35 analyzed dynamics of RNA trafficking and translation in neurons,36 and also revealed the principals of RNA decay and translation in many different species. While extremely useful, the GFP system does have some drawbacks. In particular, the expression of GFP is usually high and the 3’-UTR fusions often exist as an array of MS2 aptamers. This combination can very easily result in high background, which can be supplanted by forcing free GFP to be localized to the nucleus. 34 One way to get around the significant background has been to instead employ split GFP proteins that are fused to two different RNA binding proteins with unique RNA aptamers separately (Figure 3, b).37 In this way, reconstruction of GFP is dependent on recognition of the two RNA binding proteins in proximity on a single RNA, while unbound split GFP proteins do not fluoresce. Extension of the split GFP system has been applied to more sophisticated RNA-binding protein scaffolds. The RNA-binding protein PumHD (Pumilio homology domain), which has been commonly applied in native or modified form for targeting RNA, can be engineered to recognize pre-determined RNA sequences through RNA-protein interfacial engineering.38 The PumHD protein can be programmed with varying compositions and length, to bind desired target RNA sequences (Figure 3, c). The utilization of PumHD interactions results in high specificity, with undetectable binding of a Pumilio domain to RNA sequences that bear three or more mismatches from the target sequence.38 This specific binding further contributes to high signalto-noise ratio along with split proteins. While very effective in decreasing the background fluorescence, split protein systems are still restricted by lacking tight control of protein expression and protein folding for GFP chromophore maturation. A way to mitigate some of the problems with exogenous fluorescent proteins and background signals

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would be combined the split GFP system with the binding-dependent fluorescence turn-on, which has been achieved with hybridization-based methods in fixed samples. An improving approach is to develop turn-on RNA-small molecule interactions in live cells. The first widely used approach to turn on fluorescence was the application of the GFP-like RNA aptamer Spinach.39 Whereas, GFP fluorescence is due to protein folding and chromophore formation, Spinach relies on the binding of a small molecule (DFHBI), which does not fluorescence in solution (Figure 3, d). Spinach was initially discovered through in vitro selection methods along with several other small molecule ligands which when bound emit at several different wavelengths.39 Crystal structures of DFHBI bound to its RNA aptamer revealed that Spinach activates the small molecule by immobilizing it in a specific sequence context (Figure 3, e & f).40 The G-quadruplex is well suited for an RNA to induce fluorescence of a chromophore like DFHBI by simultaneously restricting it to a planar conformation and also forming hydrogen bonds with the functional groups on its edges. Structural insights have been critical to designing novel fluorescent RNA-small molecule interactions. Another RNA aptamer-small molecule interaction that serves a similar role as Spinach was also very recently discovered. Mango, is a much smaller RNA aptamer that binds to a non-fluorescent ligand thiazole orange (TO; ~ 3nM affinity) and when bound permits fluorescence from the ligand increasing by as much as 1,100-fold (Figure 3, g).41 Depending on the specific TO ligand, TO-Mango could be one of the most redshifted fluorescent macromolecular tags known. Mango was also demonstrated to work in living animals as injection of TO into C. elegans gonads enabled the visualization of a Mango aptamer-fused RNA.41, 42 Interestingly, crystallographic analysis of the TO-mango interaction also revealed that, like DFHBI, TO is positioned directly against a G-quadruplex, forcing the two heterocycles of TO to form a small angle of 45°with respect to each other (Figure 3, h & i).43 This orientation is thought to contribute to increased fluorescence. Overall, the identification and structural analysis of Mango and Spinach aptamers has uncovered a new way of imaging RNAs inside living cells. Turn-on fluorescent aptamers do provide a novel approach to imaging RNAs inside cells; however, the application of this method is limited by the chemical diversity of the analogs that can be used to interrogate RNA localization and also not permitting stringent washing steps on cells to remove unbound analogs, which can result in spurious fluorescent signal. To overcome these obstacles, it would be beneficial to have methods for covalent tagging of RNAs to create permeant fluorophore-RNA interactions. Recent reports have moved in this direction by developing enzymatic reactions to covalently link fluorophores and other


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chemical handles to RNA aptamer sequences. One such example is the utility of RNA-guanine transglycosylase (TGT). TGT is a key enzyme involved in the posttranscriptional modification of tRNA across the three kingdoms of life. In eukaryotes and eubacteria, TGT is involved in the introduction of queuine into the anticodon of the cognate tRNAs. Because TGT binds to a specific tRNA sequence, the tRNA-like RNA sequence can be appended to an RNA of interest. Queuine-conjugated fluorophores can then be transferred to the RNA for covalent linkage (Figure 3, j).44 Structural inspection of the RNA-TGT co-crystal structure (Figure 3, k) with the inert analog N-9-deazaguanine45 reveals how the enzyme would allow for functionality at the N-7 potion (Figure 3, l), where a solvent-exposed pocket would simply enable long tethers for attachment. TGT has been used to image RNAs in cells and further develop queuinefluorogenic thiazole orange probes that significantly lower nonspecific binding and background fluorescence and, as a result, provide up to a 100-fold fluorescence intensity increase after labeling.46 This addition to the enzymatic method enabled RNA imaging in living cells in a wash-free manner.

aptamer bound to TO (PDB 5V3F). (i) Zoom in onto the TO binding site and the G-quadruplex. (j) Schematic of protein-based covalent labeling of RNA (TGTase). (k) Overall crystal structure of the TGT-RNA interaction (PDB 1Q2R). (l) Zoom in of the TGT-RNA active site. Beyond TGT, another enzyme-RNA motif pair was developed for enzymatic imaging of RNA in cells: labeling of RNAs in mammalian cells by using tRNAIle2‐agmatidine synthetase (Tias)47 and click chemistry. The unique RNA sequence specificity and plastic Tias/substrate recognition enable the site‐specific transfer of azide/alkyne groups to an RNA molecule of interest in vitro and in mammalian cells. Subsequent click chemistry reactions facilitate the versatile labeling, functionalization, and visualization of target RNAs. The examples illustrated within this section highlight efforts to combine the attributes of biological engineering with chemical biology. The methods mentioned are historically employed to understand the localization and/or expression of specific RNAs, often relying on knowing the sequence of an RNA of interest, as well as appending artificial RNA sequences that enable small molecule or protein recognition for imaging. Further development should be employed to expand the pool of specific aptamers, fluorescent proteins, or fluorescent ligands that can enable higher dimensional imaging of many RNAs in parallel. An important consideration for these experiments is to evaluate if changes to the primary sequence of the target RNAs are altered in such a way that it can alter its normal function. For example, it has been demonstrated that fusing repeated aptamer sequences can alter the half-life or localization of RNA molecules.48 As such, special care should be taken in the design and implementation of RNA fusions for imaging or interrogating RNA function. Emerging methods to interrogate the localization of large groups of RNAs.

Figure 3. Biochemical methods for imaging RNA without the use of antisense oligos. (a) Schematic of the MS2-GFP system for protein imaging of RNA. (b) Split GFP system using two different RNA binding proteins MBP and PCP with known aptamer sequences MBS and PBS respectively. (c) Split GFP using two different pumilio domain proteins, which can be engineered to bind to specific sequences of interest in RNAs. (d) Schematic of spinach turn-on fluorescent aptamer. (e) Overall structure of the spinach aptamer bound to DFHBI (PDB 4TS0). (f) Zoom in onto the DFHBI binding site and the G-quadruplex. (g) Schematic of mango turn-on fluorescent aptamer. (h) Overall structure of the MANGO

Examining RNA localization of single molecules has given us insight into how transcripts can have programmable localization in cells. However, it is still unclear how different groups of RNAs are moved around and what the general principals are controlling the localization of classes of RNAs. To overcome this barrier, some new approaches are currently underway to discover RNAs localized to distinct subcellular organelles in an unbiased manner. Our lab has been investigating spatiallyrestricted oxidation methods to identify and tag RNAs localized to subcellular regions. We initially developed a novel method to assay RNA localization within intact living cells through spatially-restricted nucleobase oxidation.49 Using Halo-tag fusion proteins, we localized the oxygen photosensitizer dibromofluorescein (DBF) within subcellular compartments. Blue light exposure


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generated short-lived singlet oxygen, resulting in guanosine oxidation in RNAs nearby in space. Later we expanded the labeling scope to proteins. Oxidized guanosines in RNA, as well as electron-rich side chains in proteins, can react with nucleophiles in solution. We utilized propargyl amine (PA) to append alkyne handles onto these adjacent RNAs and proteins for downstream study. RT-qPCR data demonstrated that this oxidative approach provides subcellular, and even sub-organellar resolution for RNA localization studies (Figure 4, a & b). We note that this short-lived singlet oxygen method was of high enough resolution to distinguish between chromatin-associated, nucleoplasm, and nucleolus localized RNAs, all within the nucleus.50 Future work in our lab is focused on systematically localizing Halo-tag fusion proteins around the cells and identifying localized RNAs.

Figure 4. Biochemical methods for identifying large groups of RNAs with subcellular resolution. (a) Schematic for RNA tagging using spatially-restricted singlet oxygen generation. (b) Schematic of oxidized RNA tagging with propargyl amine. The alkyne handle enables attachment of biotin to tagged RNAs using “click” chemistry. (c) Schematic for protein-RNA tagging using spatially-restricted biotin tyramide generation. (d) Schematic of tyramide-conjugated protein-RNA complexes, which can be enriched using streptavidin-conjugated resins. (e) Schematic of localized ribosome profiling. (f) Close-up diagram detailing the use of BirA to conjugate biotin to a ribosome tag, which can then be used to enrich ribosomes from a subcellular location. Another exciting development has been the utilization of peroxidase enzymes to spatially tag proteins, and their associated RNAs with high spatial resolution. Proximity biotinylation using activated tyramide radicals generated with exogenously incorporated hydrogen peroxide and APEX peroxidase enzymes has been extremely fruitful in the discovery of protein localization with subcellular resolution.51 Extension of this methodology was recently performed to

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identify adjacent RNAs through the pulldown of tyramide-labeled protein-RNA complexes (Figure 4, c & d).52 This method was used to identify nuclear, cytoplasmic, mitochondrial and ER-associated RNAs. These exciting results with APEX open up the possibility of systematically studying subcellular RNA localization. As APEX relies on short-lived phenoxyl radicals (< 1 ms53), and our approach with short-lived singlet oxygen (< 3.5 μs54, 55) both should give high-resolution complimentary approaches. Identifying RNA localization in the cell does not necessarily reveal associated function. As such, more sophisticated methods are needed to close the gap and address how RNA localization is contributing to its function. Ribosome profiling is a method to determine where on mRNAs ribosomes are located.56 recent extension of ribosome profiling was performed to identify subcellular-localized ribosomes and the RNAs being translated at those positions in cells. To accomplish this, the BirA biotin ligase was localized to either the outer membrane of the ER or mitochondria. An accessory ribosomal protein was also engineered to have the BirA substrate sequence. When the localized BirA came in contact with the ribosome bearing the substrate peptide, a biotin was transferred to the ribosome (Figure 4, e & f). After cell lysis biotinylated ribosomes can be enriched and profiled for the mRNAs being translated. As expected ER ribosome profiling revealed RNAs enriched for translated proteins that would be secreted57 and mitochondrial ribosome profiling enriched for RNAs that encode for proteins to be imported into the mitochondria.58 Extension of this method to different organelles or parts of the cells that have not had translation examined would reveal a comprehensive assay of localized mRNAs and their corresponding translation. Conclusions and Future Directions Methods to analyze RNA localization with high specificity have matured tremendously over the past couple of decades. Expansion of hybridization-based methods and their merger with higher-resolution microscopy has begun to yield a more complete picture of RNA localization and differential RNA expression in unique cell types. In addition, more sophisticated biochemical methods that rely on cellular engineering and the synthesis of novel enzymatic cofactors and substrates are rapidly expanding our ability to image RNA localization in not only fixed and permeabilized cells, but living ones as well. Lastly, emergent methods for the discovery of localized RNAs are sure to open the door to additional analysis and identification of novel RNA localization “hot spots” inside cells. One example is the recent development of using CRISPR-related technology to track RNA in living cells. This is accomplished by utilizing nuclease-inactive S. pyogenes CRISPR/Cas9


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that can bind RNA in a nucleic-acid-programmed manner.59 Additionally, new Cas proteins, such as class 2 type VI RNA-guided RNA-targeting CRISPR-Cas effector Cas13a (previously known as C2c2), can also be programmed to bind and image selected RNAs in cells. These programmable approaches holds unlimited opportunity once its sensitivity and specificity are improved in the future. An urgent need in this area is to understand the dynamics of RNA localization and how that relates to processing and function. Live cell imaging with some of the methods described in Figure 3 is sure to be fruitful. As an example, the recent development of translating RNA imaging by coat protein knock-off (TRICK)60, which is designed to image the localization of mRNAs and distinguish untranslated mRNAs from those that have undergone at least one round of translation, has the potential to dramatically increase our understanding of the factors controlling nuclear export and regulating translation. In a related development, 3′RNA end accumulation during turnover (TREAT)61 also revealed the localization of RNAs to subcellular compartments and also enabled kinetic characterization of TREAT mRNA degradation within living cells. Additionally, more methods directed toward understanding an RNA translocates through the cell would be transformative. To be able to “mark” RNA at different positions within the cell and map back the order of events of localization would reveal where an RNA has been during its lifetime and potentially identify RNAs that have multiple functions throughout their time in the cell. Perhaps co-opting additional enzymatic reactions, like Tias and TGT (Figure 3, j) would enable programmed marking of RNAs as they process through different subcellular distributions in their lifetimes sequentially. To accomplish this, it’s crucial to develop additional chemical methods and merge them with cellular engineering to achieve RNA tagging with high temporal and spatial resolution. Lastly, the merge of more sophisticated bioinformatic analyses with some methods as outlined in Figure 4 should reveal several outstanding questions in RNA localization, for example, do different isoforms localize to different parts of cells? A hint of this being true has come from fractionation of neurons where different isoforms can be localized within the cell body and the axons respectively.62 Differential isoform localization can give rise to unique protein synthesis that may have different function depending on their place in the cell and the local environment. Further, being able to utilize RNA localization datasets to build testable models to understand the mechanisms of RNA localization would be extremely useful. The prevailing model in the field is that RNA-binding proteins interact with RNAs and other scaffolding proteins to bring them to their cellular destinations. As the motifs for many RNA-binding proteins are starting to be experimentally identified,63, 64 it presents an opportunity to merge RBP datasets with

localization to predict RNA-protein interactions contributing to the localization of sets of RNAs in cells. Together, multimodal, systems-level approaches, coupled with rapidly emerging technologies for quantitative RNA localization, will usher a new era in our understanding of the functional roles as well as the mechanisms of RNA localization. AUTHOR INFORMATION Corresponding Author [email protected] Author Contributions All authors have given approval to the final version of the manuscript. Funding Sources No competing financial interests have been declared. ACKNOWLEDGMENT We apologize for any work left out in this perspective. We thank members of the Spitale lab for their careful reading and critique of the manuscript. Work on RNA localization in the Spitale lab is supported by the NIH (1DP2GM119164 to RCS). RCS is a Pew Biomedical Scholar.

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