分子生物学

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

Design of the Nascent Chain Immunoprecipitation (NC IP) reporter and assay. Left. The construct carries a 3× Flag tag at the N-terminal end of Venus protein (FlagVenus). In this example, the reporter is adapted to study untranslated regions (UTR)-dependent expression by flanking FlagVenus coding sequence with UTRs of Aurora kinase A (AURKA) mRNA (depicted reporters refer to Cacioppo et al., 2023, Figure 3). The depicted reporters carry mutations in the proximal (p) or distal (d) polyadenylation signal (PAS). Right. Following reporter transfection, ribosomes are locked onto reporter mRNA by treating cells with cycloheximide (CHX), which prevents ribosome run-off and additional rounds of elongation, before cell lysis and immunoprecipitation (IP) of FlagVenus nascent chains via anti-Flag beads. Reporter mRNAs are then eluted, isolated, and quantified by RT-qPCR.

Cilia and flagella are microtubule-based hair-like organelles protruding from the surface of most eukaryotic cells, and play essential roles in cell locomotion, left-right asymmetry, embryo development, and tissue homeostasis. With isolated cilia and flagella, great progress has been made in understanding the composition, structure, and function of cilia. However, the current cilia/flagella isolation methods are deficient in the integrity or productivity of purified cilia when applied to mammalian motile cilia. Here, we describe a new protocol that isolates cilia shafts from mouse ependymal cells, by horizontal shear force and mild detergent. This method enables the production of virtually integral cilia with high yields and less cell body contamination. It is suitable for immunostaining, puromycin labeling assay, and proximity ligation assay of mammalian motile cilia.

Graphical abstract:

RNA granules (RGs) are membraneless intracellular compartments that play important roles in the post-transcriptional control of gene expression. Stress granules (SGs) are a type of RGs that form under environmental challenges and/or internal cellular stresses. Stress treatments lead to strong mRNAs translational inhibition and storage in SGs until the normal growth conditions are restored. Intriguingly, we recently showed that plant stress granules are associated with siRNA bodies, where the RDR6-mediated and transposon-derived siRNA biogenesis occurs (Kim et al., 2021). This protocol provides a technical workflow for the enrichment of cytoplasmic RGs from Arabidopsis seedlings. We used the DNA methylation-deficient ddm1 mutant in our study, but the method can be applied to any other plant samples with strong RG formation. The resulting RG fractions can be further tested for either RNAs or proteins using RNA-seq and mass spectrometry-based proteomics.

Plants make up by far the largest part of biomass on Earth. They are the primary source of food and the basis of most drugs used for medicinal purposes. Similarly to all eukaryotes, plant cells also use mitochondria for energy production. Among mitochondrial gene expression processes, translation is the least understood; although, recent advances have revealed the specificities of its main component, the mitochondrial ribosome (mitoribosome). Here, we present a detailed protocol to extract highly pure cauliflower mitochondria by differential centrifugation for the purification of mitochondrial ribosomes using a sucrose gradient and the preparation of cryo-electron microscopy (cryo-EM) grids. Finally, the specific bioinformatics pipeline used for image acquisition, the processing steps, and the data analysis used for cryo-EM of the plant mitoribosome are described. This protocol will be used for further analysis of the critical steps of mitochondrial translation, such as its initiation and regulation.

Here, we describe how to image and quantitate the translation dynamics of a bicistronic biosensor that we recently created to fairly compare cap-dependent and IRES-mediated translation at single-molecule resolution in live human cells. This technique employs a pair of complementary intrabodies loaded into living cells that co-translationally bind complementary epitopes in the two separate ORFs of the bicistronic biosensor. This causes the biosensor to fluoresce in different colors depending on which ORF/epitopes are translated. Using the biosensor together with high-resolution fluorescence microscopy and single-molecule tracking analysis allows for the quantitative comparison of translation dynamics between the two ORFs at a resolution of tens-of-nanometers in space and sub-seconds in time, which is not possible with more traditional GFP or luciferase reporters. Since both ORFs are on the same biosensor, they experience the same microenvironment, allowing a fair comparison of their relative translational activities. In this protocol, we describe how to get this assay up and running in cultured human cells so that translation dynamics can be studied under both normal and stressful cellular conditions. We also provide a number of useful tips and notes to help express components at appropriate levels inside cells for optimal live cell imaging.

Graphical abstract:

Steps required for 3-color single-molecule translation imaging and analysis.

The nucleotides involved in RNA-RNA interaction can be tagged by chemical- or UV-induced crosslinking, and further identified by classical or modern high throughput techniques. The contacts of mRNA with 18S rRNA that occur along the mRNA channel of 40S subunit have been mapped by site-specific UV crosslinking followed by reverse transcriptase termination sites (RTTS) using radioactive or fluorescent oligonucleotides. However, the sensitivity of this technique is restricted to the detection of those fragments that resulted from the most frequent crosslinkings. Here, we combined RTTS with RNAseq to map the mRNA-18S rRNA contacts with a much deeper resolution. Although aimed to detect the interaction of mRNA with the ES6S region of 18S rRNA, this technique can also be applied to map the interaction of mRNA with other non-coding RNA molecules (e.g., snRNAs, microRNAs and lncRNAs) during transcription, splicing or RNA-mediated postranscriptional regulation.

RNAs and RNA-binding proteins (RBPs) can interact dynamically in ribonucleoprotein (RNP) complexes that play important roles in controlling gene expression programs. One of the powerful ways to investigate changes in the association of RNAs with an RBP of interest is by immunoprecipitation (IP) analysis of native RNPs. RIP (RNP immunoprecipitation) analysis enables the rapid identification of endogenous RNAs bound to an RBP and to monitor time-dependent changes in this association, as well as changes in response to different metabolic and stress conditions. The protocol is based on the use of an antibody, typically an anti-RBP antibody, to immunoprecipitate the RNP complex. The RNA within the immunoprecipitated complex can then be isolated and further studied using different approaches such as PCR, microarray, Northern blot, and sequencing analyses. Among other advantages, RIP analysis (i) measures RNP associations in many samples relatively quickly, (ii) can be adapted easily to different endogenous RBPs, and (iii) provides extensive information at low cost. Among its limitations, RIP analysis does not inform on the specific sites of interaction of an RBP with a given target RNAs, although recent adaptations of RIP have been developed to overcome this problem. Here we provide an optimized protocol for RIP analysis that can be used to study RNA-protein interactions relevant to many areas of biology.

Ribosome profiling provides information on the position of ribosomes on mRNA on a genomic scale. Although this information is often used to detect changes in gene expression under different conditions, it also has great potential for yielding insight into the mechanism and regulation of protein synthesis itself. First developed in yeast, ribosome profiling involves the isolation and sequencing of ribosome-protected mRNA fragments generated by nuclease treatment. Since the application of ribosome profiling in bacteria has been problematic, we report here a systematically optimized protocol for E. coli that we have used with success for other bacteria as well. Cells are harvested by flash-freezing cultures directly in liquid nitrogen. After lysis, translation is arrested by high magnesium concentration without the use of antibiotics. These improvements eliminate artifacts induced by harvesting cells by centrifugation or filtration and by use of chloramphenicol to arrest translation. These improvements are especially appropriate for studies where the exact position of the ribosome is critical, and not merely the number of ribosomes per message, such as studies aimed at monitoring differences in local elongation rates.

Macrophages are immune cells that contribute to host defense through various mechanisms including phagocytosis and antigen presentation. Their antimicrobial capacity is subverted by clinically important intracellular pathogens such as Mycobacterium tuberculosis. The study of host-pathogen interactions using these cells is therefore of considerable interest. Such studies often seek to express tagged proteins to characterize their activities, localizations, and protein-protein interactions. Here, we describe a robust method for transient protein expression in macrophages using mRNA lipoplex transfections.

The efficiency with which proteins are produced from mRNA molecules can vary widely across transcripts, cell types, and cellular states. Methods that accurately assay the translational efficiency of mRNAs are critical to gaining a mechanistic understanding of post-transcriptional gene regulation. One way to measure translational efficiency is to determine the number of ribosomes associated with an mRNA molecule, normalized to the length of the coding sequence. The primary method for this analysis of individual mRNAs is sucrose gradient fractionation, which physically separates mRNAs based on the number of bound ribosomes. Here, we describe a streamlined protocol for accurate analysis of mRNA association with ribosomes. Compared to previous protocols, our method incorporates internal controls and improved buffer conditions that together reduce artifacts caused by non-specific mRNA–ribosome interactions. Moreover, our direct-from-fraction qRT-PCR protocol eliminates the need for RNA purification from gradient fractions, which greatly reduces the amount of hands-on time required and facilitates parallel analysis of multiple conditions or gene targets. Additionally, no phenol waste is generated during the procedure. We initially developed the protocol to investigate the translationally repressed state of the HAC1 mRNA in S. cerevisiae, but we also detail adapted procedures for mammalian cell lines and tissues.