Molecular Biology

Transcriptional pausing dynamically regulates spatiotemporal gene expression during cellular differentiation, development, and environmental adaptation. Precise measurement of pausing duration, a critical parameter in transcriptional control, has been challenging due to limitations in resolution and confounding factors. We introduce Fast TV-PRO-seq, an optimized protocol built on time-variant precision run-on sequencing (TV-PRO-seq), which enables genome-wide, single-base resolution mapping of RNA polymerase II pausing times. Unlike standard PRO-seq, Fast TV-PRO-seq employs sarkosyl-free biotin-NTP run-on with time gradients and integrates on-bead enzymatic reactions to streamline workflows. Key improvements include (1) reducing experimental time from 4 to 2 days, (2) reducing cell input requirements, and (3) improved process efficiency and simplified command-line operations through the use of bash scripts.

Genome-wide gene expression analysis is a commonly used method to quantitatively examine the transcriptional signature of any tissue or cell state. Standard bulk cell RNA sequencing (RNA-seq) quantifies RNAs in the cells of the tissue type of interest through massive parallel sequencing of cDNA synthesized from the cellular RNA. The subsequent analysis of global RNA expression and normalization of RNA expression levels between two or more samples generally assumes that cells from all samples produce equivalent amounts of RNA per cell. This assumption may be invalid in cells where MYC or MYCN expression levels are markedly different and thus, overall mRNA expression per cell is altered. Here, we describe an approach for RNA-seq analysis of MYCN-amplified neuroblastoma cells during treatment with retinoic acid, which causes dramatic downregulation of MYCN expression and induces growth arrest and differentiation of the cells. Our procedure employs spiked-in RNA standards added in ratio to the number of cells in each sample prior to RNA extraction. In the analysis of differential gene expression, the expression level of each gene is standardized to the spiked-in RNA standard to accurately assess gene expression levels per cell in conditions of high and low MYCN expression. Our protocol thus provides a step-by-step experimental approach for normalizing RNA-seq expression data on a per-cell-number basis, allowing accurate assessment of differential gene expression in cells expressing markedly different levels of MYC or MYCN.

For obtaining insights into gene networks during plant reproductive development, having transcriptomes of specific cells from developmental stages as starting points is very useful. During development, there is a balance between cell proliferation and differentiation, and many cell and tissue types are formed. While there is a wealth of transcriptome data available, it is mostly at the organ level and not at specific cell or tissue type level. Therefore, methods to isolate specific cell and tissue types are needed. One method is fluorescent activated cell sorting (FACS), but it has limitations such as requiring marker lines and protoplasting. Recently, single-cell/nuclei isolation methods have been developed; however, a minimum amount of genetic information (marker genes) is needed to annotate/predict the resulting cell clusters in these experiments. Another technique that has been known for some time is laser-assisted microdissection (LAM), where specific cells are microdissected and collected using a laser mounted on a microscope platform. This technique has advantages over the others because no fluorescent marker lines must be made, no marker genes must be known, and no protoplasting must be done. The LAM technique consists in tissue fixation, tissue embedding and sectioning using a microtome, microdissection and collection of the cells of interest on the microscope, and finally RNA extraction, library preparation, and RNA sequencing. In this protocol, we implement the use of normal slides instead of the membrane slides commonly used for LAM. We applied this protocol to obtain the transcriptomes of specific tissues during the development of the gynoecium of Arabidopsis.

Genetic networks regulate nearly all biological processes, including cellular differentiation, homeostasis, and immune responses. Determining the precise role of each gene within a regulatory network can explain its overall, integrated function, and pinpoint mechanisms underlying misregulation in disease states. Transcriptional reporter assays are a useful tool for dissecting these genetic networks, because they link a molecular process to a measurable readout, such as the expression of a fluorescent protein. Here, we introduce a new technique that uses expressed RNA barcodes as reporters, to measure transcriptional changes induced by CRISPRi-mediated genetic perturbation across a diverse, genome-wide library of guide RNAs. We describe an exemplary reporter based on the promoter that drives His4 expression in these guidelines, which can be used as a framework to interrogate other expression phenotypes. In this workflow, a library of plasmids is assembled, encoding a CRISPRi guide RNA (gRNA) along with one or more transcriptional reporters that drive expression of guide-specific nucleotide barcode sequences. For example, when interrogating regulation of the budding yeast HIS4 promoter normalized against a control housekeeping promoter that drives Pgk1 expression, this plasmid library contains a gRNA expression cassette, a HIS4 reporter driving expression of one gRNA-specific nucleotide barcode, and a PGK1 reporter driving expression of a second, gRNA-specific barcode. Long-read sequencing is used to determine which gRNA is associated with these nucleotide barcodes. The plasmid library is then transformed into yeast cells, where each cell receives one plasmid, and experiences a genetic perturbation driven by the guide on that plasmid. The expressed RNA barcodes are extracted in bulk and quantified using high-throughput sequencing, thereby measuring the effect of their corresponding gRNA on barcoded reporter expression. In the case of the HIS4 reporter described above, guides disrupting translation elongation will increase expression of the associated HIS4 barcode specifically, without changing expression of the PGK1 control barcode. It is further possible to quantify plasmid abundance by DNA sequencing, as an additional approach to normalize for differences in plasmid abundance within the population of cells. This protocol outlines the steps to prepare barcode reporter CRISPRi plasmid libraries, link guides to barcodes with long-read sequencing, and measure expression changes through barcode RNA and DNA sequencing. This method is ideal for probing transcriptional or post-transcriptional regulation, as it measures the effects of a genetic perturbation by directly quantifying reporter RNA abundance, rather than relying on indirect growth or fluorescence readouts.

Graphic abstract:

High-throughput RNA sequencing (RNA-seq) has extraordinarily advanced our understanding of gene expression and disease etiology, and is a powerful tool for the identification of biomarkers in a wide range of organisms. However, most RNA-seq methods rely on retroviral reverse transcriptases (RTs), enzymes that have inherently low fidelity and processivity, to convert RNAs into cDNAs for sequencing. Here, we describe an RNA-seq protocol using Thermostable Group II Intron Reverse Transcriptases (TGIRTs), which have high fidelity, processivity, and strand-displacement activity, as well as a proficient template-switching activity that enables efficient and seamless RNA-seq adapter addition. By combining these activities, TGIRT-seq enables the simultaneous profiling of all RNA biotypes from small amounts of starting material, with superior RNA-seq metrics, and unprecedented ability to sequence structured RNAs. The TGIRT-seq protocol for Illumina sequencing consists of three steps: (i) addition of a 3' RNA-seq adapter, coupled to the initiation of cDNA synthesis at the 3' end of a target RNA, via template switching from a synthetic adapter RNA/DNA starter duplex; (ii) addition of a 5' RNA-seq adapter, by using thermostable 5' App DNA/RNA ligase to ligate an adapter oligonucleotide to the 3' end of the completed cDNA; (iii) minimal PCR amplification, to add capture sites and indices for Illumina sequencing. TGIRT-seq for the Illumina sequencing platform has been used for comprehensive profiling of coding and non-coding RNAs in ribodepleted, chemically fragmented cellular RNAs, and for the analysis of intact (non-chemically fragmented) cellular, extracellular vesicle (EV), and plasma RNAs, where it yields continuous full-length end-to-end sequences of structured small non-coding RNAs (sncRNAs), including tRNAs, snoRNAs, snRNAs, pre-miRNAs, and full-length excised linear intron (FLEXI) RNAs.

Graphic abstract:

Figure 1. Overview of the TGIRT-seq protocol for Illumina sequencing. Major steps are: (1) Template switching from a synthetic R2 RNA/R2R DNA starter duplex with a 1-nt 3' DNA overhang (a mixture of A, C, G, and T residues, denoted N) that base pairs to the 3' nucleotide of a target RNA, and upon initiating reverse transcription by adding dNTPs, seamlessly links an R2R adapter to the 5' end of the resulting cDNA; (2) Ligation of an R1R adapter to the 3' end of the completed cDNA; and (3) Minimal PCR amplification with primers that add Illumina capture sites (P5 and P7) and barcode sequences (indices 5 and 7). The index 7 barcode is required, while the index 5 barcode is optional, to provide unique dual indices (UDIs).

Site-specific transcription arrest is the basis of emerging technologies that assess nascent RNA structure and function. Cotranscriptionally folded RNA can be displayed from an arrested RNA polymerase (RNAP) for biochemical manipulations by halting transcription elongation at a defined DNA template position. Most transcription “roadblocking” approaches halt transcription elongation using a protein blockade that is non-covalently attached to the template DNA. I previously developed a strategy for halting Escherichia coli RNAP at a chemical lesion, which expands the repertoire of transcription roadblocking technologies and enables sophisticated manipulations of the arrested elongation complexes. To facilitate this chemical transcription roadblocking approach, I developed a sequence-independent method for preparing internally modified dsDNA using PCR and translesion synthesis. Here, I present a detailed protocol for the preparation and characterization of internally modified dsDNA templates for chemical transcription roadblocking experiments.

Graphic abstract:

Precise transcription roadblocking using functionalized DNA lesions

Primary somatosensory neurons, whose cell bodies reside in the dorsal root ganglion (DRG) and trigeminal ganglion, are specialized to transmit sensory information from the periphery to the central nervous system. Our molecular understanding of peripheral sensory neurons has been limited by both their heterogeneity and low abundance compared with non-neuronal cell types in sensory ganglia. We describe a protocol to isolate nuclei from mouse DRGs using iodixanol density gradient centrifugation, which enriches for neuronal nuclei while still sampling non-neuronal cells such as satellite glia and Schwann cells. This protocol is compatible with a range of downstream applications such as single-nucleus transcriptional and epigenomic assays.

DNA transcription by RNA polymerases has always interested the scientific community as it is one of the most important processes involved in genome expression. This has led scientists to come up with different protocols allowing analysis of this process in specific locations across the genome by quantitating the amount of RNA polymerases transcribing that genomic site in a cell population. This can be achieved by either detecting the total number of polymerases in contact with that region (i.e., by chromatin immunoprecipitation (ChIP) with anti-RNA polymerase antibodies) or by measuring the number of polymerases that are effectively engaged in transcription in that position. This latter strategy is followed using transcription run-on (TRO), also known as nuclear run-on (NRO), which was first developed in mammalian cells over 40 years ago and has since been adapted to many other different organisms and high-throughput methods. Here, we detail the procedure for performing TRO in Saccharomyces cerevisiae for single genomic regions to study active transcription on a single gene scale. To do so, we wash the cells in the detergent sarkosyl, which prevents new initiations at the promoter level, and then perform an in situ reaction, leading to the radiolabeling of transcripts by RNA polymerases that were already engaged in transcription at the moment of harvesting. By subsequently quantitating the signal of these transcripts, we can determine the level of active transcription in a single gene. This presents a major advantage over other forms of transcription quantitation such as RNA polymerase ChIP, since in the latter, both active and inactive polymerases are measured. By combining both ChIP and TRO, the amount of inactive or paused polymerases on a particular gene can be estimated.

Graphic abstract:

Transcriptional run-on scheme

Transcription errors can substantially affect metabolic processes in organisms by altering the epigenome and causing misincorporations in mRNA, which is translated into aberrant mutant proteins. Moreover, within eukaryotic genomes there are specific Transcription Error-Enriched genomic Loci (TEELs) which are transcribed by RNA polymerases with significantly higher error rates and hypothesized to have implications in cancer, aging, and diseases such as Down syndrome and Alzheimer’s. Therefore, research into transcription errors is of growing importance within the field of genetics. Nevertheless, methodological barriers limit the progress in accurately identifying transcription errors. Pro-Seq and NET-Seq can purify nascent RNA and map RNA polymerases along the genome but cannot be used to identify transcriptional mutations. Here we present background Error Model-coupled Precision nuclear run-on Circular-sequencing (EmPC-seq), a method combining a nuclear run-on assay and circular sequencing with a background error model to precisely detect nascent transcription errors and effectively discern TEELs within the genome.

The 5′ cap is a ubiquitous feature of eukaryotic mRNAs. It is added in the nucleus onto newly synthesized pre-mRNA, and in the cytoplasm onto mRNAs after decapping or endonuclease cleavage. Cytoplasmic recapping can occur after loss of the cap at the native 5′ end, or downstream within the body of the mRNA. The identification and location of recapping events is key to understanding the functional consequences of this process. Here we present an approach that addresses this problem, using the Lexogen TeloPrime^® cDNA synthesis kit to tag recapped 5′ ends. TeloPrime uses a proprietary DNA ligase to add a double stranded DNA oligonucleotide onto the 3′ end of cDNA while it is base paired with mRNA. Specificity for capped ends is obtained by the oligonucleotide having an unpaired C residue that base pairs weakly with ^m7G on the mRNA 5′ end. This is followed by PCR amplification of double-stranded cDNA using primers to the appended oligonucleotide and the mRNA of interest. The resulting products are gel purified and sequenced directly (if a single band) or cloned and sequenced. The sequence at the junction between the ligated oligonucleotide and the target mRNA provides the location of the cap on the corresponding transcript. This assay is applicable to all capped transcripts. It can be used with Sanger sequencing for small numbers of transcripts or adapted for use with Illumina library sequencing.