RNA Tagging Research Articles

Terbium ion as RNA tag for slide-free pathology with deep-ultraviolet excitation fluorescence

Deep-ultraviolet excitation fluorescence microscopy has enabled molecular imaging having an optical sectioning capability with a wide-field configuration and its usefulness for slide-free pathology has been shown in recent years. Here, we report usefulness of terbium ions as RNA-specific labeling probes for slide-free pathology with deep-ultraviolet excitation fluorescence. On excitation in the wavelength range of 250–300 nm, terbium ions emitted fluorescence after entering cells. Bright fluorescence was observed at nucleoli and cytoplasm while fluorescence became weak after RNA decomposition by ribonuclease prior to staining. It was also found that the fluorescence intensity at nucleoplasm increased with temperature during staining and that this temperature-dependent behavior resembled temperature-dependent hypochromicity of DNA due to melting. These findings indicated that terbium ions stained single-stranded nucleic acid more efficiently than double-stranded nucleic acid. We further combined terbium ions and DNA-specific dyes for dual-color imaging. In the obtained image, nucleolus, nucleoplasm, and cytoplasm were distinguished. We demonstrated the usefulness of dual-color imaging for rapid diagnosis of surgical specimen by showing optical sectioning of unsliced tissues. The present findings can enhance deep-ultraviolet excitation fluorescence microscopy and consequently expand the potential of fluorescence microscopy in life sciences.

Influence of 16S rRNA Hypervariable Region on Estimates of Bacterial Diversity and Community Composition in Seawater and Marine Sediment.

To assess the influence of 16S ribosomal RNA (rRNA) tag choice on estimates of microbial diversity and/or community composition in seawater and marine sediment, we examined bacterial diversity and community composition from a site in the Central North Atlantic and a site in the Equatorial Pacific. For each site, we analyzed samples from four zones in the water column, a seafloor sediment sample, and two subseafloor sediment horizons (with stratigraphic ages of 1.5 and 5.5 million years old). We amplified both the V4 and V6 hypervariable regions of the 16S rRNA gene and clustered the sequences into operational taxonomic units (OTUs) of 97% similarity to analyze for diversity and community composition. OTU richness is much higher with the V6 tag than with the V4 tag, and subsequently OTU-level community composition is quite different between the two tags. Vertical patterns of relative diversity are broadly the same for both tags, with maximum taxonomic richness in seafloor sediment and lowest richness in subseafloor sediment at both geographic locations. Genetic dissimilarity between sample locations is also broadly the same for both tags. Community composition is very similar for both tags at the class level, but very different at the level of 97% similar OTUs. Class-level diversity and community composition of water-column samples are very similar at each water depth between the Atlantic and Pacific. However, sediment communities differ greatly from the Atlantic site to the Pacific site. Finally, for relative patterns of diversity and class-level community composition, deep sequencing and shallow sequencing provide similar results.

β-Actin mRNA interactome mapping by proximity biotinylation

The molecular function and fate of mRNAs are controlled by RNA-binding proteins (RBPs). Identification of the interacting proteome of a specific mRNA in vivo remains very challenging, however. Based on the widely used technique of RNA tagging with MS2 aptamers for RNA visualization, we developed a RNA proximity biotinylation (RNA-BioID) technique by tethering biotin ligase (BirA*) via MS2 coat protein at the 3' UTR of endogenous MS2-tagged β-actin mRNA in mouse embryonic fibroblasts. We demonstrate the dynamics of the β-actin mRNA interactome by characterizing its changes on serum-induced localization of the mRNA. Apart from the previously known interactors, we identified more than 60 additional β-actin-associated RBPs by RNA-BioID. Among these, the KH domain-containing protein FUBP3/MARTA2 has been shown to be required for β-actin mRNA localization. We found that FUBP3 binds to the 3' UTR of β-actin mRNA and is essential for β-actin mRNA localization, but does not interact with the characterized β-actin zipcode element. RNA-BioID provides a tool for identifying new mRNA interactors and studying the dynamic view of the interacting proteome of endogenous mRNAs in space and time.

NAD tagSeq reveals that NAD+-capped RNAs are mostly produced from a large number of protein-coding genes in Arabidopsis

The 5' end of a eukaryotic mRNA transcript generally has a 7-methylguanosine (m7G) cap that protects mRNA from degradation and mediates almost all other aspects of gene expression. Some RNAs in Escherichia coli, yeast, and mammals were recently found to contain an NAD+ cap. Here, we report the development of the method NAD tagSeq for transcriptome-wide identification and quantification of NAD+-capped RNAs (NAD-RNAs). The method uses an enzymatic reaction and then a click chemistry reaction to label NAD-RNAs with a synthetic RNA tag. The tagged RNA molecules can be enriched and directly sequenced using the Oxford Nanopore sequencing technology. NAD tagSeq can allow more accurate identification and quantification of NAD-RNAs, as well as reveal the sequences of whole NAD-RNA transcripts using single-molecule RNA sequencing. Using NAD tagSeq, we found that NAD-RNAs in Arabidopsis were produced by at least several thousand genes, most of which are protein-coding genes, with the majority of these transcripts coming from <200 genes. For some Arabidopsis genes, over 5% of their transcripts were NAD capped. Gene ontology terms overrepresented in the 2,000 genes that produced the highest numbers of NAD-RNAs are related to photosynthesis, protein synthesis, and responses to cytokinin and stresses. The NAD-RNAs in Arabidopsis generally have the same overall sequence structures as the canonical m7G-capped mRNAs, although most of them appear to have a shorter 5' untranslated region (5' UTR). The identification and quantification of NAD-RNAs and revelation of their sequence features can provide essential steps toward understanding the functions of NAD-RNAs.

From fluorescent proteins to fluorogenic RNAs: Tools for imaging cellular macromolecules.

The explosion in genome-wide sequencing has revealed that noncoding RNAs are ubiquitous and highly conserved in biology. New molecular tools are needed for their study in live cells. Fluorescent RNA-small molecule complexes have emerged as powerful counterparts to fluorescent proteins, which are well established, universal tools in the study of proteins in cell biology. No naturally fluorescent RNAs are known; all current fluorescent RNA tags are in vitro evolved or engineered molecules that bind a conditionally fluorescent small molecule and turn on its fluorescence by up to 5000-fold. Structural analyses of several such fluorescence turn-on aptamers show that these compact (30-100 nucleotides) RNAs have diverse molecular architectures that can restrain their photoexcited fluorophores in their maximally fluorescent states, typically by stacking between planar nucleotide arrangements, such as G-quadruplexes, base triples, or base pairs. The diversity of fluorogenic RNAs as well as fluorophores that are cell permeable and bind weakly to endogenous cellular macromolecules has already produced RNA-fluorophore complexes that span the visual spectrum and are useful for tagging and visualizing RNAs in cells. Because the ligand binding sites of fluorogenic RNAs are not constrained by the need to autocatalytically generate fluorophores as are fluorescent proteins, they may offer more flexibility in molecular engineering to generate photophysical properties that are tailored to experimental needs.

Generation of small molecule-binding RNA arrays and their application to fluorogen-binding RNA aptamers.

The discovery and engineering of more and more functions of RNA has highlighted the utility of RNA-targeting small molecules. Recently, several fluorogen-binding RNA aptamers have been developed that have been applied to live cell imaging of RNA and metabolites as RNA tags or biosensors, respectively. Although the design and application of these fluorogen-binding RNA aptamer-based devices is straightforward in theory, in practice, careful optimisation is required. For this reason, high throughput in vitro screening techniques, capable of quantifying fluorogen-RNA aptamer interactions, would be beneficial. We recently developed a method for generating functional-RNA arrays and demonstrated that they could be used to detect fluorogen-RNA aptamer interactions. Specifically, we were able to visualise the interaction between malachite green and the malachite green-binding aptamer. Here we expand this study to demonstrate that functional-RNA arrays can be used to quantify fluorogen-aptamer interactions. As proof-of-concept, we provide detailed protocols for the production of malachite green-binding RNA aptamer and DFHBI-binding Spinach RNA aptamer arrays. Furthermore, we discuss the potential utility of the technology to fluorogen-binding RNA aptamers, including application as a molecular biosensor platform. We anticipate that functional-RNA array technology will be beneficial for a wide variety of biological disciplines.

Influence of family integrated care on the intestinal microbiome of preterm infants in the neonatal intensive care unit

Objective: To investigate the influence of family integrated care (FICare) on the intestinal microbiome of preterm infants in neonatal intensive care unit (NICU). Methods: This was a prospective observational pilot study. A total of 44 preterm infants (23 boys, 52%) admitted to NICU of the Third Xiangya Hospital of Central South University from July, 2015 to June, 2017 were enrolled and divided into FICare, non-FICare groups. Totally 20 term infants (11 boys, 55%) were enrolled into control group, who were sent to the Pediatric Healthcare Clinic for regular health check on postnatal 28-31 days. All infants were free from probiotics after birth and on full enteral feeding. Clinical data of all infants were collected. Two fresh stool specimens of infants in FICare group were collected after 2 weeks of FICare implementation, without use of antibiotics during the prior 1 week. Stool specimens of infants in non-FICare group were collected at the meantime;while for the infants in control group, stool samples were collected at 4 weeks of age. All specimens were stored in-80 ℃ freezer, subsequently investigated by 16 S rRNA sequencing. The results were filtered by paired-end reads software based on RNA overlapping-splicing and tags calculation. Operational taxonomic units (OTU) were analyzed for intestinal microbiome richness. Intestinal microbiome diversity was measured with Shannon index. One-way ANOVA or Kruskal-Wallis H statistic analysis or Chi-square test was used for statistical analysis. Results: There were no significant differences among FICare, non-FICare and control groups in male proportion (52% (11/21) vs. 52% (12/23) vs. 55% (11/20), χ(2)=0.041, P=0.980), in-born ratio (90% (19/21) vs. 87% (20/23) vs. 85% (17/20), χ(2)=0.000, P=1.000), and percentage of infants with Apgar scores<7 at 5 minutes after birth (14% (3/21) vs. 9% (2/23) vs. 5% (1/20), χ(2)=0.120, P=0.729). Similarly, no significant differences were found between FICare and non-FICare groups in terms of gestational age ((29.7±1.8) vs. (29.9±1.7) weeks, t=0.378, P=0.707), birth weight ((1 266±310) vs. (1 326±318) g, t=0.631, P=0.531), median age of initiating feeds (4 vs. 4 days old, Z=0.666, P=0.505), and median age of achieving feeding volume of 120 ml/(kg·d)(13 vs. 11 days old, Z=1.014, P=0.310). However, the breast-feeding rate in FICare group (18/21, 86%) was significantly higher than that in non-FICare group (8/23, 35%) (t=11.780, P=0.001). The medium Shannon index was 0.72 (0.27,2.66), 0.61 (0.18,1.83), and 0.52 (0.08,1.71) in control, FICare, and non-FICare groups, respectively, without significant difference (H=1.823, P=0.402). The domain flora was Lactobacillus Firmicutes in all three groups, which was of the highest percentage in FICare group (71.6±5.4)%, followed by control group (65.4±6.6)% and non-FICare group (55.6±8.8)%, with a significant difference (F=27.919, P=0.000). Conclusions: FICare can improve the richness and diversity of intestinal microbiome, stimulate the establishment of flora close to those of normal breast-feeding infants in preemies in NICU, making its establishment being more similar to normal term breast-feeding infants. This effect might be caused by the increased skin-to-skin contact and increased fresh breast-milk-feeding in FICare.

PERSIA for Direct Fluorescence Measurements of Transcription, Translation, and Enzyme Activity in Cell-Free Systems.

Quantification of biology's central dogma (transcription and translation) is pursued by a variety of methods. Direct, immediate, and ongoing quantification of these events is difficult to achieve. Common practice is to use fluorescent or luminescent proteins to report indirectly on prior cellular events, such as turning on a gene in a genetic circuit. We present an alternative approach, PURExpress-ReAsH-Spinach In-vitro Analysis (PERSIA). PERSIA provides information on the production of RNA and protein during cell-free reactions by employing short RNA and peptide tags. Upon synthesis, these tags yield quantifiable fluorescent signal without interfering with other biochemical events. We demonstrate the applicability of PERSIA in measuring cell-free transcription, translation, and other enzymatic activity in a variety of applications: from sequence-structure-function studies, to genetic code engineering, to testing antiviral drug resistance.

Comprehensive identification of RNA-protein interactions in any organism using orthogonal organic phase separation (OOPS).

Existing high-throughput methods to identify RNA-binding proteins (RBPs) are based on capture of polyadenylated RNAs and cannot recover proteins that interact with nonadenylated RNAs, including long noncoding RNA, pre-mRNAs and bacterial RNAs. We present orthogonal organic phase separation (OOPS), which does not require molecular tagging or capture of polyadenylated RNA, and apply it to recover cross-linked protein-RNA and free protein, or protein-bound RNA and free RNA, in an unbiased way. We validated OOPS in HEK293, U2OS and MCF10A human cell lines, and show that 96% of proteins recovered were bound to RNA. We show that all long RNAs can be cross-linked to proteins, and recovered 1,838 RBPs, including 926 putative novel RBPs. OOPS is approximately 100-fold more efficient than existing methods and can enable analyses of dynamic RNA-protein interactions. We also characterize dynamic changes in RNA-protein interactions in mammalian cells following nocodazole arrest, and present a bacterial RNA-interactome for Escherichia coli. OOPS is compatible with downstream proteomics and RNA sequencing, and can be applied in any organism.

Immunoprecipitation and SDS-PAGE for Cross-Linking Immunoprecipitation (CLIP).

This first part of this protocol is designed to optimize purification of the RNABP by immunoprecipitation for cross-linking immunoprecipitation (CLIP) experiments. The key variables to assess are the quality and quantity of antibody needed to immunoprecipitate most but not quite all of the RNABP (the titration will decrease nonspecific binding), and the tolerance of the antibody:antigen interaction to stringent wash conditions. The results of these experiments can be checked first by western blot, and subsequently using the pilot CLIP protocol described in the second half of this protocol. RNase-treated cross-linked RNABP:RNA complexes from mixed lysates or cell pellets are immunoprecipitated using conditions optimized in the first half of the protocol. 3' Linkers are added, the RNA is radiolabeled, and the complexes are purified on SDS-PAGE. The pilot experiment will identify the optimal RNase concentration for the particular sample and will assess the quality and purity of the RNABP-RNA complexes following labeling of the RNA tags with 32P. This can be done without ligation of the 3' linker as described in the main protocol below. The pilot experiment assesses whether sufficient RNA-protein complexes can be detected by autoradiography and whether contaminating RNA ligands are present in immunoprecipitations compared with control samples. Once it is confirmed that the signal-to-noise ratio for detection of RNA-protein complexes after immunoprecipitation is sufficient, the optimal immunoprecipitation conditions should be incorporated into the general CLIP protocol including the steps of cross-linking, RNase digestion, linker ligation, and labeling of RNA "tags," and the results analyzed by autoradiography.

Isolation of the RNA Cross-Linking Immunoprecipitation (CLIP) Tags, 5'-Linker Ligation, Reverse Transcription-Polymerase Chain Reaction (RT-PCR) Amplification, and Sequencing.

This protocol describes purification of RNA cross-linking immunoprecipitation (CLIP) tags by proteinase K digestion of the cross-linked protein, addition of a 5' linker to the RNA tags, and amplification of the product by transcription-polymerase chain reaction (RT-PCR). Use of this protocol adds another important purification step: sizing of the PCR products to enrich for those derived from RNA originally cross-linked to the desired RNABP. Finally, sequencing of the PCR products is described. There are two strategies for sequencing the PCR products of "CLIPed" RNA. Low-throughput sequencing involves cloning of PCR products, conventional minipreps, and sequencing. This can be performed on the PCR products generated here using standard protocols for A-tailing the PCR product and TA-cloning. This may be a worthwhile strategy when analyzing a small number of clones. In general, particularly in light of falling costs, high-throughput sequencing is the preferred method for sequencing the products of CLIPed RNA. This protocol describes a method for reamplifying PCR products with primers suitable for use on Illumina's Solexa platform. Although this protocol is specific to the Illumina deep-sequencing platform, similar schemes for reamplification of the initial PCR products can be used to add platform-specific sequences to the termini of the PCR-amplified DNA.

3'-Linker Ligation and Size Selection by SDS-PAGE for Cross-Linking Immunoprecipitation (CLIP).

This protocol describes the purification by denaturing polyacrylamide gel electrophoresis of RNA linkers for cross-linking immunoprecipitation (CLIP). Purification is necessary because if the 3' linker loses the puromycin blocking group, concatemerization of the 3' linker will occur during the 3' linker ligation reaction. In addition, truncated linkers make bioinformatic processing of the sequencing results more difficult than it need be. Additionally, this protocol describes the treatment of coimmunoprecipitated RNA tags for CLIP with alkaline phosphatase to remove the 3' phosphate remaining after RNase digestion. Dephosphorylation prevents intramolecular circularization of RNA during subsequent ligation to the linker. The purified RNA linker, blocked with puromycin at its 3' end to prevent linker-linker multimerization, is then ligated to the 3' end of the RNA tag. Removal of free linker is accomplished by performing the ligation while the RNABP:RNA complex is associated, via antibody, to protein A Dynabeads, allowing thorough washing and linker removal. Additional purification is achieved by SDS-PAGE and transfer of the size-selected RNABP:RNA complexes to nitrocellulose.

RNA Structure and Cellular Applications of Fluorescent Light‐Up Aptamers

AbstractThe cellular functions of RNA are not limited to their role as blueprints for protein synthesis. In particular, noncoding RNA, such as, snRNAs, lncRNAs, miRNAs, play important roles. With increasing numbers of RNAs being identified, it is well known that the transcriptome outnumbers the proteome by far. This emphasizes the great importance of functional RNA characterization and the need to further develop tools for these investigations, many of which are still in their infancy. Fluorescent light‐up aptamers (FLAPs) are RNA sequences that can bind nontoxic, cell‐permeable small‐molecule fluorogens and enhance their fluorescence over many orders of magnitude upon binding. FLAPs can be encoded on the DNA level using standard molecular biology tools and are subsequently transcribed into RNA by the cellular machinery, so that they can be used as fluorescent RNA tags (FLAP‐tags). In this Minireview, we give a brief overview of the fluorogens that have been developed and their binding RNA aptamers, with a special focus on published crystal structures. A summary of current and future cellular FLAP applications with an emphasis on the study of RNA–RNA and RNA–protein interactions using split‐FLAP and Förster resonance energy transfer (FRET) systems is given.

RNA Structure and Cellular Applications of Fluorescent Light-Up Aptamers.

The cellular functions of RNA are not limited to their role as blueprints for protein synthesis. In particular, noncoding RNA, such as, snRNAs, lncRNAs, miRNAs, play important roles. With increasing numbers of RNAs being identified, it is well known that the transcriptome outnumbers the proteome by far. This emphasizes the great importance of functional RNA characterization and the need to further develop tools for these investigations, many of which are still in their infancy. Fluorescent light‐up aptamers (FLAPs) are RNA sequences that can bind nontoxic, cell‐permeable small‐molecule fluorogens and enhance their fluorescence over many orders of magnitude upon binding. FLAPs can be encoded on the DNA level using standard molecular biology tools and are subsequently transcribed into RNA by the cellular machinery, so that they can be used as fluorescent RNA tags (FLAP‐tags). In this Minireview, we give a brief overview of the fluorogens that have been developed and their binding RNA aptamers, with a special focus on published crystal structures. A summary of current and future cellular FLAP applications with an emphasis on the study of RNA–RNA and RNA–protein interactions using split‐FLAP and Förster resonance energy transfer (FRET) systems is given.

Maintenance of grafting‐induced epigenetic variations in the asexual progeny of Brassica oleracea and B. juncea chimera

Grafting-induced variations have been observed in many plant species, but the heritability of variation in progeny is not well understood. In our study, adventitious shoots from the C cell lineage of shoot apical meristem (SAM) grafting chimera TCC (where the origin of the outmost, middle and innermost cell layers, respectively, of SAM is designated by 'T' for tuber mustard and 'C' for red cabbage) were induced and identified as r-CCC (r=regenerated). To investigate the maintenance of grafting variations during cell propagation and regeneration, different generations of asexual progeny (r-CCCn, n=generation) were established through successive regeneration of axillary shoots from r-CCC. The fourth generation of r-CCC (r-CCC4) was selected to perform whole genome bisulfite sequencing for comparative analysis of hetero-grafting-induced global methylation changes relative to r-s-CCC4 (s=self-grafting). Increased CHH methylation levels and proportions were observed in r-CCC4, with substantial changes occurring in the repeat elements. Small RNA sequencing revealed 1135 specific small interfering RNA (siRNA) tags that were typically expressed in r-CCC, r-CCC2 and r-CCC4. Notably, 65% of these specific siRNAs were associated with repeat elements, termed RE siRNAs. Subsequent analysis revealed that the CHH methylation of RE siRNA-overlapping regions was mainly hypermethylation in r-CCC4, indicating that they were responsible for directing and maintaining grafting-induced CHH methylation. Moreover, the expression of 13 differentially methylated genes (DMGs) correlated with the phenotypic variation, showing differential expression levels between r-CCC4 and r-s-CCC4. These DMGs were predominantly CG hypermethylated, their methylation modifications corresponded to the transcription of relative methyltransferase.

A multicolor riboswitch-based platform for imaging of RNA in live mammalian cells.

RNAs directly regulate a vast array of cellular processes, emphasizing the need for robust approaches to fluorescently label and track RNAs in living cells. Here, we develop an RNA imaging platform using the cobalamin riboswitch as an RNA tag and a series of probes containing cobalamin as a fluorescence quencher. This highly modular ‘Riboglow’ platform leverages different colored fluorescent dyes, linkers and riboswitch RNA tags to elicit fluorescent turn-on upon binding RNA. We demonstrate the ability of two different Riboglow probes to track mRNA and small non-coding RNA in live mammalian cells. A side-by-side comparison revealed that Riboglow outperformed the dye binding aptamer Broccoli and performed on par with the gold standard RNA imaging system, the MS2-fluorescent protein system, while featuring a much smaller RNA tag. Together, the versatility of the Riboglow platform and ability to track diverse RNAs suggest broad applicability for a variety of imaging approaches.

The Lsm1-7/Pat1 complex binds to stress-activated mRNAs and modulates the response to hyperosmotic shock.

RNA-binding proteins (RBPs) establish the cellular fate of a transcript, but an understanding of these processes has been limited by a lack of identified specific interactions between RNA and protein molecules. Using MS2 RNA tagging, we have purified proteins associated with individual mRNA species induced by osmotic stress, STL1 and GPD1. We found members of the Lsm1-7/Pat1 RBP complex to preferentially bind these mRNAs, relative to the non-stress induced mRNAs, HYP2 and ASH1. To assess the functional importance, we mutated components of the Lsm1-7/Pat1 RBP complex and analyzed the impact on expression of osmostress gene products. We observed a defect in global translation inhibition under osmotic stress in pat1 and lsm1 mutants, which correlated with an abnormally high association of both non-stress and stress-induced mRNAs to translationally active polysomes. Additionally, for stress-induced proteins normally triggered only by moderate or high osmostress, in the mutants the protein levels rose high already at weak hyperosmosis. Analysis of ribosome passage on mRNAs through co-translational decay from the 5’ end (5P-Seq) showed increased ribosome accumulation in lsm1 and pat1 mutants upstream of the start codon. This effect was particularly strong for mRNAs induced under osmostress. Thus, our results indicate that, in addition to its role in degradation, the Lsm1-7/Pat1 complex acts as a selective translational repressor, having stronger effect over the translation initiation of heavily expressed mRNAs. Binding of the Lsm1-7/Pat1p complex to osmostress-induced mRNAs mitigates their translation, suppressing it in conditions of weak or no stress, and avoiding a hyperresponse when triggered.

Cross platform validation of a unique method for deep genomic and proteomic analysis of rare immune cell populations

Abstract Advances in flow and mass cytometry expand the number of cell parameters that can be interrogated improving our understanding of the myriad immune cell types. These technologies, however, are limited in the types of analytes profiled in a single sample. In addition to efforts to understand the immune system for the treatment of autoimmune diseases, the immune system is increasingly a target for cancer therapeutics. These efforts require deep characterization to understand rare cell populations, requiring next-generation methods that expand on flow cytometry. 3D Flow™ Analysis integrates flow cytometry cell sorting and the nCounter® Vantage 3D™ RNA:Protein Immune Cell Profiling Assay* to characterize sorted immune cell populations with direct digital counting of 770 RNA and 30 proteins. To determine if the method is agnostic to the cell sorting technology used, 3D Flow analysis was run with the Becton Dickenson FACSARIA IIu, Sony SH800z, and Beckman Coulter MoFloXDP. Stimulated and unstimulated PBMC were co-stained with flow antibodies for CD3, CD8, and CD4 plus 30 DNA-labeled antibodies for NanoString readout. 5,000 CD3+/CD4+ and CD3+/CD8+ cells were sorted on each platform and immediately lysed to release cellular RNA and DNA tags from NanoString antibodies. RNA and DNA tags were directly hybridized to NanoString barcodes and run on the nCounter system. Expression of 30 proteins and 770 RNA across cell populations correlated between the three sorting platforms, showing 3D Flow is agnostic to the sorting technology used. Importantly, high-plex RNA data was obtained without additional molecular biology methods, such as RNA purification or library construction, simplifying incorporation into a flow cytometry laboratory.

Uncovering cell type-specific complexities of gene expression and RNA metabolism by TU-tagging and EC-tagging.

Cell type-specific transcription is a key determinant of cell fate and function. An ongoing challenge in biology is to develop robust and stringent biochemical methods to explore gene expression with cell type specificity. This challenge has become even greater as researchers attempt to apply high-throughput RNA analysis methods under in vivo conditions. TU-tagging and EC-tagging are in vivo biosynthetic RNA tagging techniques that allow spatial and temporal specificity in RNA purification. Spatial specificity is achieved through targeted expression of pyrimidine salvage enzymes (uracil phosphoribosyltransferase and cytosine deaminase) and temporal specificity is achieved by controlling exposure to bioorthogonal substrates of these enzymes (4-thiouracil and 5-ethynylcytosine). Tagged RNAs can be purified from total RNA extracted from an animal or tissue and used in transcriptome profiling analyses. In addition to identifying cell type-specific mRNA profiles, these techniques are applicable to noncoding RNAs and can be used to measure RNA transcription and decay. Potential applications of TU-tagging and EC-tagging also include fluorescent RNA imaging and selective definition of RNA-protein interactions. TU-tagging and EC-tagging hold great promise for supporting research at the intersection of RNA biology and developmental biology. This article is categorized under: Technologies > Analysis of the Transcriptome.

RNA Tagging: Preparation of High-Throughput Sequencing Libraries.

Protein-RNA networks, in which a single protein binds and controls multiple mRNAs, are central in biological control. As a result, methods to identify protein-RNA interactions that occur in vivo are valuable. The "RNA Tagging" approach enables the investigator to unambiguously identify global protein-RNA interactions in vivo and is independent of protein purification, cross-linking, and radioactive labeling steps. Here, we provide a protocol to prepare high-throughput sequencing libraries for RNA Tagging experiments.