分子生物学

Fork stability is key to genome DNA duplication and genetic integrity. Long non-coding RNAs (LncRNAs) may play vital roles in fork stabilization and chromatin remodeling. Existing techniques such as NCC-RNA sequencing are useful to identify LncRNAs on nascent chromatin DNA. However, there is still a lack of methods for LncRNAs purification directly from replicative forks, hindering a deep understanding of the functions of LncRNAs in fork regulation. Here, we provide a step-by-step protocol named iROND (isolate RNAs on nascent DNA). iROND was developed and modified from iPOND, a well-known method for purifying fork-associated proteins. iROND relies on click chemistry reaction of 5'-ethynyl-2'-deoxyuridine (EdU)-labeled forks and biotin. After streptavidin pull down, fork-associated LncRNAs and proteins are purified simultaneously. iROND is compatible with downstream RNA sequencing, qPCR confirmation, and immunoblotting. Integrated with functional methods such as RNA fluorescent in situ hybridization (RNA FISH) and DNA fiber assay, it is feasible to screen fork-binding LncRNAs in defined cell lines and explore their functions. In summary, we provide a purification pipeline of fork-associated LncRNAs. iROND is also useful for studying other types of fork-associated non-coding RNAs.

Key features

• Purify long non-coding RNAs (LncRNAs) directly from replication forks.

• Connects to RNA sequencing for screening easily.

• Allows testing various genotoxic stress responses.

• Provides LncRNA candidate list for downstream functional research.

Graphical overview

Schematic overview of isolate RNAs on nascent DNA (iROND) protocol. Cells were pulse-labeled with 5'-ethynyl-2'-deoxyuridine (EdU) for 10 min before paraformaldehyde fixation. EdU-positive forks were ligated with biotin through Click-IT chemistry reaction. Genomic DNA was ultrasonically cracked and crosslinked with streptavidin for pulling down. Both RNA and protein components were purified. RNA components were used for downstream RNA sequencing and qPCR validation. Protein components were used for immunoblotting to evaluate binding dynamics of fork-associated proteins such as helicase, topoisomerase, and DNA polymerases.

Many single nucleotide polymorphisms (SNPs) identified by genome-wide association studies exert their effects on disease risk as expression quantitative trait loci (eQTL) via allele-specific expression (ASE). While databases for probing eQTLs in tissues from normal individuals exist, one may wish to ascertain eQTLs or ASE in specific tissues or disease-states not characterized in these databases. Here, we present a protocol to assess ASE of two possible target genes (GPNMB and KLHL7) of a known genome-wide association study (GWAS) Parkinson’s disease (PD) risk locus in postmortem human brain tissue from PD and neurologically normal individuals. This was done using a sequence of RNA isolation, cDNA library generation, enrichment for transcripts of interest using customizable cDNA capture probes, paired-end RNA sequencing, and subsequent analysis. This method provides increased sensitivity relative to traditional bulk RNAseq-based and a blueprint that can be extended to the study of other genes, tissues, and disease states.

Key features

• Analysis of GPNMB allele-specific expression (ASE) in brain lysates from cognitively normal controls (NC) and Parkinson’s disease (PD) individuals.

• Builds on the ASE protocol of Mayba et al. (2014) and extends application from cells to human tissue.

• Increased sensitivity by enrichment for desired transcript via RNA CaptureSeq (Mercer et al., 2014).

• Optimized for human brain lysates from cingulate gyrus, caudate nucleus, and cerebellum.

Graphical overview

Information on RNA localisation is essential for understanding physiological and pathological processes, such as gene expression, cell reprogramming, host–pathogen interactions, and signalling pathways involving RNA transactions at the level of membrane-less or membrane-bounded organelles and extracellular vesicles. In many cases, it is important to assess the topology of RNA localisation, i.e., to distinguish the transcripts encapsulated within an organelle of interest from those merely attached to its surface. This allows establishing which RNAs can, in principle, engage in local molecular interactions and which are prevented from interacting by membranes or other physical barriers. The most widely used techniques interrogating RNA localisation topology are based on the treatment of isolated organelles with RNases with subsequent identification of the surviving transcripts by northern blotting, qRT-PCR, or RNA-seq. However, this approach produces incoherent results and many false positives. Here, we describe Controlled Level of Contamination coupled to deep sequencing (CoLoC-seq), a more refined subcellular transcriptomics approach that overcomes these pitfalls. CoLoC-seq starts by the purification of organelles of interest. They are then either left intact or lysed and subjected to a gradient of RNase concentrations to produce unique RNA degradation dynamics profiles, which can be monitored by northern blotting or RNA-seq. Through straightforward mathematical modelling, CoLoC-seq distinguishes true membrane-enveloped transcripts from degradable and non-degradable contaminants of any abundance. The method has been implemented in the mitochondria of HEK293 cells, where it outperformed alternative subcellular transcriptomics approaches. It is applicable to other membrane-bounded organelles, e.g., plastids, single-membrane organelles of the vesicular system, extracellular vesicles, or viral particles.

Key features

• Tested on human mitochondria; potentially applicable to cell cultures, non-model organisms, extracellular vesicles, enveloped viruses, tissues; does not require genetic manipulations or highly pure organelles.

• In the case of human cells, the required amount of starting material is ~2,500 cm² of 80% confluent cells (or ~3 × 10⁸ HEK293 cells).

• CoLoC-seq implements a special RNA-seq strategy to selectively capture intact transcripts, which requires RNases generating 5′-hydroxyl and 2′/3′-phosphate termini (e.g., RNase A, RNase I).

• Relies on nonlinear regression software with customisable exponential functions.

Graphical overview

The study of translation is important to the understanding of gene expression. While genome-wide measurements of translation efficiency (TE) rely upon ribosome profiling, classical approaches to address translation of individual genes of interest rely on biochemical methods, such as polysome fractionation and immunoprecipitation (IP) of ribosomal components, or on reporter constructs, such as luciferase reporters. Methods to investigate translation have been developed that, however, require considerable research effort, including addition of numerous features to mRNA regions, genomic integration of reporters, and complex data analysis. Here, we describe a simple biochemical reporter assay to study TE of mRNAs expressed from a transiently transfected plasmid, which we term Nascent Chain Immunoprecipitation (NC IP). The assay is based on a plasmid expressing an N-terminally Flag-tagged protein and relies on the IP of Flag-tagged nascent chains from elongating ribosomes, followed by quantitative reverse transcription polymerase chain reaction (RT-qPCR) quantification of eluted mRNA. We report that elution of mRNA following IP can be achieved by treatment with puromycin, which releases ribosome-mRNA complexes, or with purified Flag peptide, which instead releases nascent chain-ribosome-mRNA complexes. In the example described in this protocol, untranslated regions (UTRs) of a gene of interest were used to flank a FlagVenus coding sequence, with the method allowing to infer UTR-dependent regulation of TE. Importantly, our method enables discrimination of translating from non-translating mRNAs. Additionally, it requires simple procedures and standard laboratory equipment. Our method can be used to test the effect of regulators, such as microRNAs or therapeutic drugs or of various genetic backgrounds, on translation of any user-selected mRNA.

Key features

• The novel NC IP protocol builds upon a previously published method for detection of mRNA-binding proteins (Williams et al., 2022).

• The NC IP protocol is adapted for detecting mRNA actively undergoing translation.

• The method uses mammalian cell culture but could be adapted to multiple organisms, including budding yeast (S. cerevisiae).

Graphical overview

The transfection of microRNA (miRNA) mimics and inhibitors can lead to the gain and loss of intracellular miRNA function, helping us better understand the role of miRNA during gene expression regulation under specific physical conditions. Our previous research has confirmed the efficiency and convenience of using liposomes to transfect miRNA mimics or inhibitors. This work uses miR-424 as an example, to provide a detailed introduction for the transfection process of miRNA mimics and inhibitors in the regular SW982 cell line and primary rheumatoid arthritis synovial fibroblasts (RASF) cells from patients by using lipofection, which can also serve as a reference to miRNA transfection in other cell lines.

Key features

• MiRNA mimics and inhibitors transfection in regular SW982 cell line and primary RASF cells.

• Treatment and culture of RASF primary cells before transfection.

Using liposomes for transfection purposes.

Here, we present an approach combining fluorescence in situ hybridization (FISH) and immunolabeling for localization of pri-miRNAs in isolated nuclei of A. thaliana. The presented method utilizes specific DNA oligonucleotide probes, modified by addition of digoxigenin-labeled deoxynucleotides to its 3′ hydroxyl terminus by terminal deoxynucleotidyl transferase (TdT). The probes are then detected by immunolabeling of digoxigenin (DIG) using specific fluorescent-labeled antibodies to visualize hybridized probes. Recently, we have applied this method to localize pri-miRNA156a, pri-miRNA163, pri-miRNA393a, and pri-miRNA414 in the nuclei isolated from leaves of 4-week-old A. thaliana. The present approach can be easily implemented to analyze nuclear distribution of diverse RNA classes, including mRNAs and pri-miRNAs in isolated fixed cells or nuclei from plant.

Inflammation of the gastrointestinal tract is a prevalent pathology in diseases such as inflammatory bowel disease (IBD). Currently, there are no therapies to prevent IBD, and available therapies to treat IBD are often sub-optimal. Thus, an unmet need exists to better understand the molecular mechanisms underlying intestinal tissue responses to damage and regeneration. The recent development of single-cell RNA (sc-RNA) sequencing-based techniques offers a unique opportunity to shed light on novel signaling pathways and cellular states that govern tissue adaptation or maladaptation across a broad spectrum of diseases. These approaches require the isolation of high-quality cells from tissues for downstream transcriptomic analyses. In the context of intestinal biology, there is a lack of protocols that ensure the isolation of epithelial and non-epithelial compartments simultaneously with high-quality yield. Here, we report two protocols for the isolation of epithelial and stromal cells from mouse and human colon tissues under inflammatory conditions. Specifically, we tested the feasibility of the protocols in a mouse model of dextran sodium sulfate (DSS)-induced colitis and in human biopsies from Crohn’s patients. We performed sc-RNA sequencing analysis and demonstrated that the protocol preserves most of the epithelial and stromal cell types found in the colon. Moreover, the protocol is suitable for immunofluorescence staining of surface markers for epithelial, stromal, and immune cell lineages for flow cytometry analyses. This optimized protocol will provide a new resource for scientists to study complex tissues such as the colon in the context of tissue damage and regeneration.

Key features

• This protocol allows the isolation of epithelial and stromal cells from colon tissues.

• The protocol has been optimized for tissues under inflammatory conditions with compromised cell viability.

• This protocol is suitable for experimental mouse models of colon inflammation and human biopsies.

Graphical overview

Graphical representation of the main steps for the processing of colon tissue from dextran sodium sulfate (DSS)-treated mice (upper panel) and frozen biopsies from Crohn’s patients (lower panel)

In eukaryotic cells, RNA biogenesis generally requires processing of the nascent transcript as it is being synthesized by RNA polymerase. These processing events include endonucleolytic cleavage, exonucleolytic trimming, and splicing of the growing nascent transcript. Endonucleolytic cleavage events that generate an exposed 5′-monophosphorylated (5′-PO₄) end on the growing nascent transcript occur in the maturation of rRNAs, tRNAs, and mRNAs. These 5′-PO₄ ends can be a target of further processing or be subjected to 5′-3′ exonucleolytic digestion that may result in termination of transcription. Here, we describe how to identify 5′-PO₄ ends of intermediates in nascent RNA metabolism. We capture these species via metabolic labeling with bromouridine followed by immunoprecipitation and specific ligation of 5′-PO₄ RNA ends with the 3′-hydroxyl group of a 5′ adaptor (5′-PO₄ Bru-Seq) using RNA ligase I. These ligation events are localized at single nucleotide resolution via highthroughput sequencing, which identifies the position of 5′-PO₄ groups precisely. This protocol successfully detects the 5′monophosphorylated ends of RNA processing intermediates during production of mature ribosomal, transfer, and micro RNAs. When combined with inhibition of the nuclear 5′-3′ exonuclease Xrn2, 5′-PO₄ Bru-Seq maps the 5′ splice sites of debranched introns and mRNA and tRNA 3′ end processing sites cleaved by CPSF73 and RNaseZ, respectively.

Key features

• Metabolic labeling for brief periods with bromouridine focuses the analysis of 5′-PO₄ RNA ends on the population of nascent transcripts that are actively transcribed.

• Detects 5′-PO₄ RNA ends on nascent transcripts produced by all RNA polymerases.

• Detects 5′-PO₄ RNA ends at single nucleotide resolution.

Circular RNA (circRNA) is an intriguing class of non-coding RNA that exists as a continuous closed loop. With the improvements in high throughput sequencing, biochemical analysis, and bioinformatic algorithms, studies on circRNA expression became abundant in recent years. However, functional studies of circRNA are still limited. Subcellular localization of circRNA may provide some clues in elucidating its biological functions by performing subcellular fractionation assay. Notably, circRNAs that are predominantly found in the cytoplasm are more likely to be involved in post-transcriptional gene regulation, e.g., acting as micoRNA sponge, whereas nuclear-retained circRNAs are predicted to play a role in transcriptional regulation. Subcellular fractionation could help researchers to narrow down and prioritize downstream experiments. The majority of the currently available protocols describe the steps for subcellular fractionation followed by western blot analysis for protein molecules. Here, we present a protocol for the subcellular fractionation of cells to detect circRNA via RT-qPCR with divergent primers. Moreover, detailed steps for the generation of specific circRNAs-enriched cDNA included in this protocol will enhance the amplification and detection of low-abundance circRNAs. This will be useful for researchers studying low-abundance circRNAs.

Key features

• This protocol builds upon the method developed by Gagnon et al. (2014) and extends its application to circRNA study.

• Protocol for amplification of low levels of circRNA expression.

• Analysis takes into consideration the ratio of cytoplasmic RNA concentration to nuclear RNA concentration.

Graphical overview

When performing expression analysis either for coding RNA (e.g., mRNA) or non-coding RNA (e.g., miRNA), reverse transcription quantitative real-time polymerase chain reaction (RT-qPCR) is a widely used method. To normalize these data, one or more stable endogenous references must be identified. RefFinder is an online web-based tool using four almost universally used algorithms for assessing candidate endogenous references—delta-Ct, BestKeeper, geNorm, and Normfinder. However, the online interface is presently cumbersome and time consuming. We developed an R package, RefSeeker, which performs easy and straightforward RefFinder analysis by enabling raw data import and calculation of stability from each of the algorithms and provides data output tools to create graphs and tables. This protocol uses RefSeeker R package for fast and simple RefFinder stability analysis.

Key features

• Perform stability analysis using five algorithms: Normfinder, geNorm, delta-Ct, BestKeeper, and RefFinder.

• Identification of endogenous references for normalization of RT-qPCR data.

• Create publication-ready graphs and tables output.

• Step-by-step guide dialog window for novice R users.

Graphical overview

Simple workflow diagram. Two main workflow paths are presented. A) Using the RefSeeker wizard allows non-R programmers to easily load data and choose between selected output formats. B) Command line interface provides more options to control input and output formats and to automate analysis.