Molecular Biology

Pseudouridine (Ψ), the most prevalent modified base in cellular RNAs, has been mapped to numerous sites not only in rRNAs, tRNAs, and snRNAs but also mRNAs. Although there have been multiple techniques to identify Ψs, due to the recent development of sequencing technologies some reagents are not compatible with the current sequencer. Here, we show the updated Pseudo-seq, a technique enabling the genome-wide identification of pseudouridylation sites with single-nucleotide precision. We provide a comprehensive description of Pseudo-seq, covering protocols for RNA isolation from human cells, library preparation, and detailed data analysis procedures. The methodology presented is easily adaptable to any cell or tissue type with high-quality mRNA isolation. It can be used for discovering novel pseudouridylation sites, thus constituting a crucial initial step toward understanding the regulation and function of this modification.

N⁶-methyladenosine (m⁶A) is the most prevalent internal modification of eukaryotic messenger RNAs (mRNAs), affecting their fold, stability, degradation, and cellular interaction(s) and implicating them in processes such as splicing, translation, export, and decay. The m⁶A modification is also extensively present in non-coding RNAs, including microRNAs (miRNAs), ribosomal RNAs (rRNAs), and transfer RNAs (tRNAs). Common m6A methylation detection techniques play an important role in understanding the biological function and potential mechanism of m⁶A, mainly including the quantification and specific localization of m⁶A modification sites. Here, we describe in detail the dot blotting method for detecting m⁶A levels in RNA (mRNA as an example), including total RNA extraction, mRNA purification, dot blotting, and data analysis. This protocol can also be used to enrich specific RNAs (such as tRNA, rRNA, or miRNA) by isolation technology to detect the m⁶A level of single RNA species, so as to facilitate further studies of the role of m6A in biological processes.

RNA sequencing allows for the quantification of the transcriptome of embryos to investigate transcriptional responses to various perturbations (e.g., mutations, infections, drug treatments). Previous protocols either lack the option to genotype individual samples, or are laborious and therefore difficult to do at a large scale. We have developed a protocol to extract total nucleic acid from individual zebrafish embryos. Individual embryos are lysed in 96-well plates and nucleic acid is extracted using SPRI beads. The total nucleic acid can be genotyped and then DNase I treated to produce RNA samples for sequencing. This protocol allows for processing large numbers of individual samples, with the ability to genotype each sample, which makes it possible to undertake transcriptomic analysis on mutants at timepoints before the phenotype is visible.

Graphic abstract:

Extraction of total nucleic acid from individual zebrafish embryos for genotyping and RNA-seq.

Recent popularization of next-generation sequencing enables conducting easy transcriptome analysis. Nevertheless, substantial RNA isolation work prior to RNA sequencing, as well as the high cost involved, still makes the routine use of large-scale transcriptome analysis difficult. For example, conventional phenol-chloroform RNA extraction cannot be easily applied to hundreds of samples. Therefore, we developed Direct-TRI, a new cost-effective and high throughput RNA-extraction method that uses a commercial guanidine-phenol-based RNA extraction reagent and a 96-well silica column plate. We applied Direct-TRI to zebrafish whole larvae and juvenile samples and obtained comparable RNA qualities by several different homogenization methods such as vortexing, manual homogenizing, and freezing/crushing. Direct-TRI enabled the extraction of 192 RNA samples in an hour with a cost of less than a dollar per sample. Direct-TRI is useful for large-scale transcriptome studies, manipulating hundreds of zebrafish individuals, and may be used with other animal samples.

Many cells contain spatially defined subcellular regions that perform specialized tasks enabled by localized proteins. The subcellular distribution of these localized proteins is often facilitated by the subcellular localization of the RNA molecules that encode them. A key question in the study of this process of RNA localization is the characterization of the transcripts present at a given subcellular location. Historically, experiments aimed at answering this question have centered upon microscopy-based techniques that target one or a few transcripts at a time. However, more recently, the advent of high-throughput RNA sequencing has allowed the transcriptome-wide profiling of the RNA content of subcellular fractions. Here, we present a protocol for the isolation of cell body and neurite fractions from neuronal cells using mechanical fractionation and characterization of their RNA content.

Graphic abstract:

Fractionation of neuronal cells and analysis of subcellular RNA contents

Transcriptional analysis has become a cornerstone of biological research, and with the advent of cheaper and more efficient sequencing technology over the last decade, there exists a need for high-yield and efficient RNA extraction techniques. Fungi such as the human pathogen Candida albicans present a unique obstacle to RNA purification in the form of the tough cell wall made up of many different components such as chitin that are resistant to many common mammalian or bacterial cell lysis methods. Typical in vitro C. albicans cell harvesting methods can be time consuming and expensive if many samples are being processed with multiple opportunities for product loss or sample variation. Harvesting cells via vacuum filtration rather than centrifugation cuts down on time before the cells are frozen and therefore the available time for the RNA expression profile to change. Vacuum filtration is preferred for C. albicans for two main reasons: cell lysis is faster on non-pelleted cells due to increased exposed surface area, and filamentous cells are difficult to pellet in the first place unlike yeast or bacterial cells. Using mechanical cell lysis, by way of zirconia/silica beads, cuts down on time for processing as well as overall cost compared to enzymatic treatments. Overall, this method is a fast, efficient, and high-yield way to extract total RNA from in vitro cultures of C. albicans.

Parasites of the genus Leishmania infect the mammalian hosts, including mice and humans and cause cutaneous or visceral leishmaniasis depending upon the parasite species transmitted by the vector sandfly. Leishmania amazonensis is one of the Leishmania species responsible for the cutaneous form of the disease. We have inoculated with these parasites the ear dermis of mice. RNA preparations were performed from fragmented tissues using a buffer containing guanidin isothiocynate (RLT buffer, RNeasy Mini Kit, Qiagen, SAS, France) and β-mercaptoethanol. Both reagents facilitate the isolation of intact RNA from tissues and the use of the RNeasy Kits present with several advantages that facilitate the isolation of pure non-degraded total RNA: i) This method allows to avoid the presence of phenol in the RNA extraction buffer, commonly used in alternative protocols; ii) Moreover Diethylpyrocarbonate (DEPC) treatment of glassware, to avoid RNAses contamination of the samples, is not required with this protocol; iii) Finally, it is a fast procedure and the isolated total RNA may be concentrated in a small volume thus facilitating its use for downstream experimental procedures.

Adaptation is thought to proceed in part through spatial and temporal changes in gene expression. Fish species such as the threespine stickleback are powerful vertebrate models to study the genetic architecture of adaptive changes in gene expression since divergent adaptation to different environments is common, they are abundant and easy to study in the wild and lab, and have well-established genetic and genomic resources. Fish gills, due to their respiratory and osmoregulatory roles, show many physiological adaptations to local water chemistry, including differences in gene expression. However, obtaining high-quality RNA using popular column-based extraction methods can be challenging from small tissue samples high in cartilage and bone such as fish gills. Here, we describe a bead-based mRNA extraction and transcriptome RNA-seq protocol that does not use purification columns. The protocol can be readily scaled according to sample size for the purposes of diverse gene expression experiments using animal or plant tissue.

Tissues are comprised of different cell types whose interactions elicit distinct gene expression patterns that regulate tissue formation, regeneration, homeostasis and repair. Analysis of these gene expression patterns require methods that can capture as closely as possible the transcriptomes of cells of interest in their tissue microenvironment. Current technologies designed to study in situ transcriptomics are limited by their low sensitivity that require cell types to represent more than 1% of the total tissue, making it challenging to transcriptionally profile rare cell populations rapidly isolated from their native microenvironment. To address this problem, we developed fluorouracil-tagged RNA sequencing (Flura-seq) that utilizes cytosine deaminase (CD) to convert the non-natural pyrimidine fluorocytosine to fluorouracil. Expression of S. cerevisiae CD and exposure to fluorocytosine generates fluorouracil and metabolically labels newly synthesized RNAs specifically in cells of interest. Fluorouracil-tagged RNAs can then be immunopurified and used for downstream analysis. Here, we describe the detailed protocol to perform Flura-seq both in vitro and in vivo. The robustness, simplicity and lack of toxicity of Flura-seq make this tool broadly applicable to many studies in developmental, regenerative, and cancer biology.

The purification of nucleic acids is one of the most common procedures employed in modern molecular biology laboratories. Typically, commercial column-based protocols are utilized to isolate DNA or RNA from various sources. However, these methods not only require specialized equipment, but are also extremely expensive for high-throughput applications. Although an elegant answer to this issue can be provided by paramagnetic beads, bead-based open-source protocols have been limited in the past. Here, we provide an easy to follow step-by-step manual for the synthesis of paramagnetic beads, as well as their functionalization with either a silica- or a carboxyl-surface that can be used to replace the commercial columns with self-made magnetic beads. Together with a variety of detailed protocols for their use in high-throughput nucleic acids extractions, this bead synthesis method forms the recently published open platform Bio-On-Magnetic-Beads (BOMB), which is available on PLOS Biology (Oberacker et al., 2019). Updated protocols can be found on the associated webpage (https://bomb.bio).