Molecular Biology

Preserving biological samples in the field is essential for ensuring high-quality nucleic acid extraction and reliable downstream molecular analyses. Broadly, two main preservation strategies are available: physical preservation, such as flash freezing in liquid nitrogen, which halts enzymatic activity by rapid cooling, and chemical preservation, using stabilizing reagents that inactivate nucleases and protect nucleic acids even at ambient temperatures. This protocol presents a comparative approach using liquid nitrogen and a commercial stabilizing reagent (DNA/RNA Shield, Zymo Research) to preserve tissue from five marine invertebrate species: two cold-water corals, two sponges, and one bivalve. Samples preserved by each method were processed with the AllPrep DNA/RNA Mini kit (Qiagen) to extract both RNA and DNA. RNA quality was assessed using RNA Integrity Number (RIN) scores. The stabilizing reagent preserved high-quality RNA in sponge and bivalve samples but did not prevent RNA degradation in coral tissues, which showed lower RIN scores compared to those preserved in liquid nitrogen. DNA yields were also consistently lower in tissues preserved with DNA/RNA Shield across all species. These findings suggest that DNA/RNA Shield can be a viable alternative to liquid nitrogen for some marine invertebrates, particularly in field conditions where cryopreservation is impractical. However, for cold-water corals, liquid nitrogen remains essential to ensure RNA integrity for transcriptomic analyses and other sensitive molecular applications (e.g., RT-qPCR).

N⁶-methyladenosine (m⁶A) is the most abundant internal modification in mRNA and is regulated primarily by the balance between the METTL3 methylase complex and two demethylases, FTO (fat mass and obesity-associated protein) and ALKBH5 (α-ketoglutarate-dependent dioxygenase alkB homolog). Reflecting this prevalence, m⁶A participates in virtually every step of RNA metabolism, influencing a wide range of physiological and pathological processes. The first step in studying m⁶A is genome-wide mapping, typically performed by m⁶A-seq, which sequences RNA fragments immunoprecipitated with an m⁶A-specific antibody. This is followed by identification of RRACH motifs (R = A or G; H = A, C, or U) within these sequences, with m⁶A being located at the third nucleotide. The second step involves mutating the putative m⁶A sites to establish a causal link between the modification and downstream biological effects. Since the mapping step has been covered in several detailed protocols, this article focuses on the second step—mutagenesis of RRACH motifs and subsequent functional analysis of the mutations by ectopic expression. The 3′ untranslated region (UTR) of the mouse Runx2 gene is used as an example. The mutant and wild-type sequences are inserted into a luciferase reporter vector and transfected into 293FT cells to evaluate how loss of m⁶A affects luciferase protein levels. The same reporter plasmids are also used in an RNA stability assay with a transcription inhibitor. Although site-specific demethylation of endogenous mRNA would be preferable, it remains technically challenging despite many attempts. Thus, ectopic expression of the mutated target gene remains a widely used and practical alternative.

Synthetic trans-acting small interfering RNAs (syn-tasiRNAs) are 21-nucleotide small RNAs designed to induce highly specific and efficient gene silencing in plants. Traditional approaches rely on the transgenic expression of ~1 kb TAS precursors, which limits their use in non-model species, under strict GMO regulations, and in size-constrained expression or delivery systems. This protocol describes a rapid workflow for the design, assembly, and delivery of syn-tasiRNAs derived from much shorter precursors, referred to as minimal precursors. The pipeline includes in silico design of highly specific syn-tasiRNA sequences, cloning of minimal precursors into plant expression or potato virus X (PVX)-based viral vectors through Golden Gate or Gibson assembly, and delivery to plants through Agrobacterium-mediated expression or by spraying crude extracts containing recombinant PVX expressing the minimal precursors. These methodologies make syn-tasiRNA-based tools more accessible and broadly applicable for plant research and biotechnology across diverse species and experimental contexts.

Long noncoding RNAs (lncRNAs) are increasingly understood to play important roles in cell biology, development, and disease, though the vast majority of annotated lncRNAs have yet to be functionally characterized. Disrupting lncRNAs is often challenging owing to their tolerance for mutations (e.g., single-nucleotide polymorphisms and short indels) along with the limitations of other genetic knockdown strategies such as RNA interference (RNAi). Here, we describe a protocol to achieve robust knockdown of lncRNAs in the fruit fly Drosophila using a self-cleaving ribozyme. The 111-bp ribozyme cassette, which consists of the N79 hammerhead ribozyme flanked by flexible linker sequences, is inserted into transcript regions of lncRNA genes using CRISPR/Cas9-mediated homology-directed repair (HDR). The fluorescent eye transformation marker is then removed using a piggyBac transposase, leaving no other modifications at the lncRNA locus save the ribozyme cassette insertion. When transcribed as part of the lncRNA, the ribozyme folds and catalyzes its own self-cleavage, resulting in two RNA cleavage fragments. The efficacy of lncRNA knockdown is then evaluated using reverse transcription quantitative PCR (RT-qPCR) and single-molecule RNA fluorescence in situ hybridization (smFISH). This approach has resulted in efficient knockdown of both nuclear and cytoplasmic lncRNAs in Drosophila, with knockdown of steady-state RNA levels in 3' cleavage fragments typically exceeding 90% and no evidence of off-target effects. The method can also be applied to protein-coding genes in order to knock down specific mRNA isoforms. Thus, self-cleaving ribozymes are a valuable addition to the genetic toolkit in Drosophila.

Circular RNAs (circRNAs) are covalently closed RNA molecules known for their increased stability compared to linear RNAs. Synthetic circRNAs are being developed as RNA therapeutics, while natural circRNAs are being investigated for their biological roles in eukaryotes and their potential as disease biomarkers. Consequently, the accurate detection and validation of circRNAs is crucial for advancements in both fundamental RNA research and biotechnological applications. Common methods for circRNA validation involve RT-PCR using divergent primers, followed by sequencing across the circRNA junction. However, most described methods are high-throughput approaches that require time-consuming RNA processing steps, and they are unable to detect highly structured circRNAs. Additionally, methods for low-throughput sequencing of small circRNAs (<150 nt) require cloning prior to sequencing. A simplified protocol for the validation of circRNA sequences irrespective of structure, sequence complexity, and length has not yet been described. In this method, we describe an improved RT-PCR protocol for circRNA detection by using UltraMarathonRT^® (uMRT), a highly processive reverse transcriptase. Unlike other reverse transcriptases, uMRT can reverse-transcribe large, structured circRNAs of varying sizes, at ambient temperatures, enabling sequencing of the resulting concatemeric amplicons generated by RT-PCR and other methods. Using this method, we sequenced circRNAs containing highly structured internal ribosome entry sites commonly utilized in synthetic circRNAs, natural circRNAs containing repetitive elements, and small circRNAs, all without the need for cloning. With this new platform, we offer a protocol for the precise detection of nearly any circRNA species.

Translation is a key step in decoding the genetic information stored in DNA. Regulation of translation is an important step in gene expression control and is essential for healthy organismal development and behavior. Despite the importance of translation regulation, its impact and dynamics remain only partially understood. One reason is the lack of methods that enable the real-time visualization of translation in the context of multicellular organisms. To overcome this critical gap, microscopy-based methods that allow visualization of translation on single mRNAs in living cells and animals have been developed. A powerful approach is the SunTag system, which enables real-time imaging of nascent peptide synthesis with high spatial and temporal resolution. This protocol describes the implementation and use of the SunTag translation imaging system in the small round worm Caenorhabditis elegans. The protocol provides details on how to design, carry out, and interpret experiments to image translation dynamics of an mRNA of interest in a cell type of choice of living C. elegans. The ability to image translation live enables better understanding of translation and reveals the mechanisms underlying the dynamics of cell type–specific and subcellular localization of translation in development.

Chromatin-associated RNAs (caRNAs) have been increasingly recognized as key regulators of gene expression and genome architecture. A few technologies, such as ChRD-PET and RedChIP, have emerged to assess protein-mediated RNA–chromatin interactions, but each has limitations. Here, we describe the TaDRIM-seq (targeted DNA-associated RNA and RNA–RNA interaction mapping by sequencing) technique, which combines Protein G (PG)-Tn5-targeted DNA tagmentation with in situ proximity ligation to simultaneously profile caRNAs across genomic regions and capture global RNA–RNA interactions within intact nuclei. This approach reduces the required cell input, shortens the experimental duration compared to existing protocols, and is applicable to both mammalian and plant systems.

Adult muscle stem cells (MuSCs) are the key cellular source for regenerating skeletal muscle in vertebrates. MuSCs are typically identified in skeletal muscle by the expression of the paired box protein 7 (PAX7) protein. Here, we developed a combined RNA fluorescent in situ hybridization (FISH) using RNAscope technology and an immunofluorescence (IF) protocol for the simultaneous detection of Pax7 mRNA and PAX7 protein in individual MuSCs in vivo. Interestingly, we show that while most PAX7⁺ (protein) MuSCs express Pax7 mRNA, there is a subset of Pax7⁺ (mRNA) cells that do not express PAX7 protein. Altogether, we developed a combined FISH/IF protocol that allows for the co-detection of mRNA and protein in MuSCs in vivo, a strategy that can be applied to any target gene. The functional significance of the Pax7-expressing subset of cells lacking PAX7 protein prior to injury remains unknown.

Regulated IRE1-dependent decay (RIDD) is a critical cellular mechanism mediated by the endoplasmic reticulum (ER) stress sensor IRE1α, which cleaves a variety of RNA targets to regulate ER homeostasis. Current in vitro assays to study IRE1α activity largely rely on synthetic or in vitro transcribed RNA substrates, which may not fully replicate the physiological complexities of native RNA molecules. Here, we present a comprehensive protocol to assess IRE1α-dependent RNA cleavage activity using total RNA isolated directly from mouse tissues. This protocol provides a step-by-step guide for tissue collection, RNA isolation, an ex vivo RIDD assay, cDNA synthesis, and subsequent RT-PCR analysis of target mRNA cleavage products. Key reagents include active IRE1α protein, the RIDD-specific inhibitor 4μ8C, and target-specific primers for RIDD-regulated genes such asBloc1s1 and Col6a1. Quantitative assessment is achieved using agarose gel electrophoresis and imaging software. This methodology enables the study of IRE1α's RNA cleavage activity under conditions that closely mimic in vivo environments, providing a more physiologically relevant approach to understanding the role of RIDD in cellular and tissue-specific contexts.

Thousands of RNAs are localized to specific subcellular locations, and these localization patterns are often required for optimal cell function. However, the sequences within RNAs that direct their transport are unknown for almost all localized transcripts. Similarly, the RNA content of most subcellular locations remains unknown. To facilitate the study of subcellular transcriptomes, we developed the RNA proximity labeling method OINC-seq. OINC-seq utilizes photoactivatable, spatially restricted RNA oxidation to specifically label RNA in proximity to a subcellularly localized bait protein. After labeling, these oxidative RNA marks are then read out via high-throughput sequencing due to their ability to induce predictable misincorporation events by reverse transcriptase. These induced mutations are then quantitatively assessed for each gene using our software package PIGPEN. The observed mutation rate for a given RNA species is therefore related to its proximity to the localized bait protein. This protocol describes procedures for assaying RNA localization via OINC-seq experiments as well as computational procedures for analyzing the resulting data using PIGPEN.