Ribosome footprinting, or Ribo-seq, has revolutionized the studies of translation. It was originally developed for yeast and mammalian cells in culture (Ingolia et al., 2009). Herein, we describe a plant-optimized hands-on ribosome footprinting protocol derived from previously published procedures of polysome isolation (Ingolia et al., 2009; Mustroph et al., 2009) and ribosome footprinting (Ingolia et al., 2009; Ingolia et al., 2013). With this protocol, we have been able to successfully isolate and analyze high-quality ribosomal footprints from different stages of in vitro grown Arabidopsis thaliana plants (dark-grown seedlings [Merchante et al., 2015] and 13-day-old plantlets in plates and plants grown in liquid culture [unpublished results]).
[Background] The central role of translation in modulating gene activity has long been recognized, yet the systematic exploration of quantitative changes in translation at a genome-wide scale in response to a specific stimulus has only recently become technically feasible. The ribosome footprinting technology (often known as the Ribo-seq), developed originally for yeast and mammalian cells in culture, has revolutionized the studies of translation regulation and gene expression, as it allows to determine the exact positions of the ribosomes at a genome-wide scale and at a single-codon resolution (Ingolia et al., 2009).
Prior to the development of Ribo-seq, the most common methods employed to study translation regulation in plants were the isolation of polysomal RNA via sucrose gradient centrifugation or translating ribosome affinity purification (TRAP) followed by Northern blotting, qRT-PCR, microarrays, or RNA-seq. The first method, known as polysome profiling, relies on resolving distinct polysomal fractions on a sucrose gradient via ultracentrifugation (Branco-Price et al., 2008; Missra and von Arnim, 2014; Li et al., 2015). By comparing different plant growth conditions or mutants, one could infer the changes in the rates of translation from observing a shift in the distribution of mRNAs between polysomal fractions. For example, if a transcript becomes more abundant in the monosomal fraction with the concomitant decrease in the higher order polysomes, the translation of this mRNA is considered as down-regulated. The key limitation of this technique, however, is its low resolution of higher-order polysomes (and thus mild quantitative changes in translation are often missed) and the inability to differentiate between polysomal RNAs that undergo active translation versus are loaded with arrested ribosomes (for example, those ‘stuck’ in the upstream open reading frames of a transcript). The second polysomal RNA isolation technique, TRAP, is based on the stable expression of an epitope-tagged ribosomal protein followed by the immunoprecipitation of entire ribosomes along with their associated mRNAs (Zanneti et al., 2005; Reynoso et al., 2015). While this latter method accommodates both global and tissue-specific studies of translation (achieved by driving the expression of a tagged ribosomal protein in a ubiquitous versus tissue-specific manner), its use is limited to transformable species where transgenic lines can be generated. Furthermore, transcriptomic analysis of TRAP samples per se does not provide a quantitative measure of translation (unless coupled with Ribo-seq [Juntawong et al., 2014]), as any mRNA with one or more ribosomes bound to it will be purified by TRAP. Also, since TRAP relies on epitope-tagging and the tag may interfere with the function of the ribosome, the regulation of translation of some mRNAs may be disrupted in the TRAP transgenic lines, e.g., due to a reduced ability of the tagged ribosome to associate with specific proteins at certain stages of translation. Another limitation of TRAP is that it typically uses a specific redundant isoform of a ribosomal protein for tagging, such as RPL18, and thus likely purifies only a subset of ribosomes that carry just that RPL18 variant. Given that there are multiple RPL18-like proteins in plant genomes, using one specific ribosomal protein isoform for tagging misses the ribosomes that utilize an alternative RPL18 isoform.
The method of choice for our studies, the Ribo-seq, does not involve transgenic line generation nor affinity purification, thus avoiding many of the limitations of the aforementioned earlier techniques. Most importantly, the single-codon resolution of the ribosome footprinting technology allows researchers to map the ribosomes on the mRNAs and thus clearly distinguish between the transcripts harboring productive ribosomes translating the main genic open reading frames versus transcripts associated with non-productive ribosomes arrested in the 5’UTRs. Not only does this method offer a snapshot of a whole-genome view of ribosomal distribution at an unprecedented resolution, it also enables the true quantitative measure of translational efficiency of every expressed gene in the genome by correlating the Ribo-seq data with the transcriptional information obtained via RNA-seq. Nonetheless, even the Ribo-Seq has its own drawbacks, as it cannot discriminate between mRNA subpopulations with different translation efficiencies, giving an average translation efficiency readout for each expressed gene.
Herein, we provide a plant-optimized Ribo-seq protocol that enables the study of translation regulation through the isolation of high-quality ribosomal footprints from different developmental stages of in vitro grown Arabidopsis thaliana seedlings. It describes step by step how to pellet and digest polysomes, isolate monosomes, extract the mRNA footprints, and generate sequencing libraries for the Illumina platform. The protocol also describes the preparation of parallel RNA-seq libraries to account for transcriptional regulation. We conclude the description of our method with a brief summary of how to analyze the sequencing results.
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