Biochemistry
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Genome-wide Quantification of Translation in Budding Yeast by Ribosome Profiling
Chapters
Summary December 21st, 2017
Translational regulation plays an important role in the control of protein abundance. Here, we describe a high-throughput method for quantitative analysis of translation in the budding yeast Saccharomyces cerevisiae.
Transcript
The overall goal of this procedure is to identify which regions of mRNA are translated into proteins quantitatively measure protein translation in budding yeast saccharomyces cerevisiae at the genome-wide level. Ribosome profiling can be used to monitor the rate of proteins and disease and to study the mechanisms of translational regulation. The main advantage of this technique is that it adds much greater sensitivity compared to other methods, and of those, measuring proteins and disease in vivo at the single contact resolution.
Though this protocol's specific for budding yeast, it is also useful for researchers who are trying to establish ribosome profiling in other model systems, such as mammalian cells and tissues. Extracts will be prepared from yeast cells grown to mid log phase. Use a glass holder filter assembly to filter the culture through 0.45 micromembrane filters.
Scrape the pellet with the spatula, flash freeze in liquid nitrogen, and store at minus 80 degrees celsius. Put three chrome steel 3.2 millimeter beads into a 1.8 milliliter stainless steel tube and pre-chill the tube in liquid nitrogen. Add the frozen pellets and cover the steel tube with a silicone rubber cap.
Homogenize the sample by cryo-grinding for 10 seconds at 4200 rpm. To keep the samples frozen during the entire grinding procedure it is critical to chill the tube in liquid nitrogen for at least 10 seconds in between each grinding. When the homogenization is done, add one milliliter of lysis buffer that contains cycloheximide to the sample.
Mix well by pipetting and transfer the sample to a new 1.5 milliliter tube. Centrifuge at 20, 000 times g at four degrees Celsius for five minutes. Transfer the supernatant into a new 1.5 milliliter tube.
Transfer 100 microliters of lysate to another new 1.5 milliliter tube for poly(A)mRNA isolation. Flash freeze the tubes with the mRNA samples on the rest of the lysate, which will be used for footprint extraction in liquid nitrogen, and store at minus 80 degrees celsius. On the night before the footprint extraction, thaw previously prepared sucrose gradients at four degrees celsius.
On the following day, thaw the cell lysate on ice. Transfer an aliquot of cell lysate containing 50 OD260 units to a new 1.5 milliliter tube, add fresh lysis buffer up to one milliliter and 10 microliters of RNase one. Incubate at room temperature with gentle rotation on a head over heels rotator for one hour.
Next, gently layer the lysate sample on top of a 10%to 50%sucrose gradient in polymer tubes. Using an SW41 ti rotor, ultra centrifuge at 35, 000 rpm at four degrees celsius for three hours. During the ultra cetrifugation, set up the gradient fractionation system.
Turn on the components and allow the UV monitor to warm up for at least 30 minutes. Fill the syringe with chase solution and remove any air bubbles inside the syringe and canula. When the ultracentrifugation is complete, install and pierce the ultracentrifuge tube containing the sucrose gradient.
Start the syringe pump at one milliliter per minute and collect one milliliter fractions while monitoring the A254 values. Fractions representing the ADS monosome peak are collected and pooled. Filter the fractions through 0.5 milliliter centrifugal filters at 12, 000 times g at four degrees celsius for 10 minutes, discard the flow-through, and repeat until the volume is less than 100 microliters.
At 400 microliters of release buffer mixed by pipetting and incubate on ice for 10 minutes, transfer the unit into a new collection tube and centrifuge at 12, 000 times g at four degrees celsius for 10 minutes. Collect the flow-through. Subsequently, purify the footprint fragments and precipitate the samples using ethanol precipitation as described in the text protocol.
To begin this procedure, centrifuge the precipitated footprint and fragmented mRNA samples at 20, 000 times g and four degrees celsius for 30 minutes. After removing ethanol and air drying the pellets, we suspend each pellet in 7.75 microliters of water. To each pellet, add one microliter of 10XT4 polynucleotide kinase buffer, one microliter of T4 polynucleotide kinase, and 0.25 microliters of RNase inhibitor.
Incubate at 37 degrees celsius for one hour. Add 10 microliters of 2X TBE urea sample buffer to 10 microliters of each footprint and mRNA sample. Incubate the samples and appropriate controls at 75 degrees celsius for three minutes, spin down, and put on ice for one minute.
Load each sample into two wells of a 15%TBE urea gel and separate the RNA fragments by gel electrophoresis at 180 volts for one hour. Stain the gel with a nucleic acid gel stain, protected from light for five minutes. While visualizing the gel under a blue light transilluminator, excise the proper band between 28 and 32 nucleotides for footprint samples and around 50 to 70 nucleotides for mRNA samples.
Freeze the excised polyacrylamide gel pieces at minus 80 degrees celsius for at least 10 minutes before extracting RNA from the polyacrylamide gel. Prior to reverse transcription, perform three prime adapter ligation and precipitate the samples as described in the text protocol. Resuspend each pellet in 11.5 microliters of nuclease-free water.
Then add 0.5 microliters of eight micromolar reverse transcription primer and one microliter of dNTP mix. Incubate at 65 degrees celsius for five minutes and then place on ice. To each sample, add four microliters of 5x first strand buffer, two microliters of 0.1 molar DTT, 0.5 microliters of RNase inhibitor, and 0.5 microliters of reverse transcriptase.
Incubate at 48 degrees celsius for 30 minutes, 65 degrees celsius for one minute, and 80 degrees celsius for five minutes. Proceed immediately to the hydrolysis of RNA by adding 0.8 microliters of two molar sodium hydroxide to each sample and incubating at 98 degrees celsius for 30 minutes. Add 0.8 microliters of two molar hydrogen chloride to neutralize the reaction.
After precipitating the samples as described in the text protocol, resuspend each pellet in five microliters of nuclease-free water and add five microliters of 2x TBE urea sample buffer. Incubate at 75 degrees celsius for three minutes, spin down, and ice for one minute. Load the sample in one well of a 10%TBE urea gel.
Separate the products of the reverse transcription reaction by gel electrophoresis 180 volts for 50 minutes. Stain the gel with a nucleic acid gel stain protected from light for five minutes. Using a blue light transilluminator, excise the band around 128 nucleotides and higher.
Freeze the polyacrylamide gel pieces at minus 80 degrees celsius for at least 10 minutes before extracting DNA from the polyacrylamide gel. To circularize the DNA, resuspend each DNA pellet in 16.75 microliters of nuclease-free water. Add two microliters of 10x single-stranded DNA lygase buffer, one microliter of 50 millimolar manganese chloride, and 0.25 microliters of single-stranded DNA Ligase.
Incubate at 60 degrees celsius for one hour, followed by 80 degrees celsius for 10 minutes. Without precipitating, store the single-stranded DNA ligation reaction product at minus 20 degrees celsius. Subsequently, perform PCR library amplification followed by library quantification and high throughput sequencing as described in the text protocol.
Representative bio analyzer profiles of the PCR amplified footprint library and the sequencing library obtained for mRNA samples are shown. The expected average size of the footprint library is 148 to 152 nucleotides, and the expected size of the mRNA library is 170 to 190 nucleotides. This graph shows the combined number of mRNA, footprint, and rRNA reads obtained for two independent biological replicates of a wild type sample and the hbs1 deletion mutant.
50%to 60%of all sequenced reads in the footprint libraries corresponded to ribosome-protected footprint fragments. In contrast, only 3%of rRNA fragments were observed in mRNA libraries, demonstrating that poly(A)mRNA isolation allows effective elimination of rRNA reads. Both footprint and mRNA sample libraries show good reproducibility as indicated by a Pearson correlation coefficient of about 0.99 between matched samples.
When combined with mRNA abundance measurements, ribosome profiling allows measuring changes in translation efficiency between experimental conditions. Significantly up-regulated and down-regulated genes in the hbs1 deletion mutant are grouped in accordance to whether they are affected by a change in mRNA transcription, translation efficiency, or by a common effect. Once mastered, the entire protocol can be performed in approximately 11 days.
While attempting this procedure, it is important to remember to wear gloves and perform all steps using RNase free reagents and consumables to avoid RNase contamination and lower the risk of RNA degradation in the samples. After watching this video, you should have a good understanding of how to generate ribosome footprints using nuclease digestion, isolate intact ribosome footprint complexes by sucrose gradient fractionation, and prepare DNA libraries for deep sequencing. Don't forget that working with liquid nitrogen can be extremely hazardous, and precautions such as wearing safety glasses and cryo-gloves should always be taken while handling extremely cold materials.
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