Ribosome Profiling Protocol

Introduction to Ribosome Profiling

Ribosome profiling, commonly referred to as Ribo-Seq, stands as a potent technique employed in the investigation of translation, the intricate process through which ribosomes orchestrate the synthesis of proteins from mRNA templates. This cutting-edge method offers a snapshot of ribosome positions along mRNAs, enabling scientists to scrutinize the translation status of individual mRNAs and assess their efficiency. Ribosome profiling encompasses the capture and sequencing of mRNA fragments safeguarded by ribosomes. Labelled as ribosome-protected footprints (RPFs), these fragments delineate the precise locations of ribosomes on mRNAs, offering valuable insights into the dynamics of translation. The principal objective is to quantify and chart ribosome presence throughout the transcriptome, ultimately disclosing the genes actively undergoing translation and the pace at which this process unfolds.

The detailed protocol for Ribosome Profiling is as follows:

Ribosome Profiling Library Preparation

Cytosolic Fractionation

Extract the cytosolic fraction (final volume of 600 μL), using the hypotonic lysis buffer without RNase Inhibitor. Maintain 4 °C in cold room to avoid unwanted mRNA degradation during the process.

RNase I Footprinting and Ribosome Recovery

1. Take the 600 μL of cytosolic fraction and add 7.5 μL of RNase I (100 U/μL).
2. Incubate for 45 min at room temperature with gentle mixing on nutator.
3. To stop the RNase I digestion, add 10 μL of SUPERase Inhibitor and tap the tubes gently.
4. Run 15-45% sucrose and confirm efficient RNase I digestion.
5. Collect and combine fractions 6, 7, and 8. Fractions 9-10 include disome-protected mRNA fragments which may represent ribosome stalling, and are excluded from our analysis.
6. Add 4.5 mL of Trizol-LS to the combined fractions (total 6 mL) and extract total RNA.

Footprint Gel Purification

1. Add 2× denaturing loading buffer to each sample.
2. Prepare 1 μL of 26 nt and 34 nt RNA marker and 1 μL of 10 bp DNA ladder as control. Add 4 μL of 10 mM Tris-HCl (pH 8.0) and 5 μL of 2× denaturing loading buffer to each.
3. Denature the sample at 80 °C for 90 s and transfer the sample to ice directly.
4. Separate the sample using 10% polyacrylamide TBE-Urea gel in 0.5× TBE buffer with constant power (5 W) for 80 min until the bromophenol blue dye reaches two third of the gel. Flushing each well with 0.5× TBE solution with syringe before sample loading helps to generate nicely shaped bands.
5. Carefully disassemble the gel cassette and stain the gel for 3 min with 1× SYBR gold in 0.5× TBE buffer.
6. Excise the ribosome footprint (24-36 nt region) and 26 nt/34 nt marker. 26 nt/34 nt footprints extracted here serve as a control for the remaining steps.
7. Overnight gel extraction. Add 400 μL of RNA gel extraction buffer and freeze the sample for 30 min on dry ice. Leave the sample overnight (15 h is recommended) with gentle mixing. Collect all solution and transfer to non-stick RNase-free eppitubes. Precipitate the footprints by adding 2 μL Glycoblue and 500 μL isopropanol. Carry out precipitation for 30 min or more on dry ice. Centrifuge at maximum speed at 4 °C for 30 min using a tabletop centrifuge. Footprints frequently end up on the side of the tubes. If this happens, resuspend footprints and centrifuge again. Completely remove the supernatant and allow air dry for 5 min. Dissolve footprints in 10 μL RNase-free water.

Ribo-Zero rRNA Depletion

1. Conduct the rRNA depletion using Ribo-zero rRNA removal kit following the manufacturer's protocol.
2. Concentrate rRNA-depleted sample according to the kit manufacturer's protocol. At the final elution step, use 8 μL of RNase-free water, which will give 7 μL final elute.

Dephosphorylation, Linker Ligation, and Purification of Linker Ligated Products

1. Dephosphorylation reaction. Mix the 7 μL of rRNA depleted footprint and 1 μL PNK buffer. Incubate at 75 °C for 1 min and cool on ice. Add 1 μL of SUPERase Inhibitor and 1 μL T4 PNK. Incubate for 1 h at 37 °C (total volume: 10 μL). Upon completion, heat-inactivate samples at 65 °C for 3 min and cool samples on ice.
2. Linker ligation reaction. Mix the 10 μL of dephosphorylated products and 1 μL of linker-1. Denature the sample for 90 s at 80 °C and immediately cool on ice for 10 min. Add 1 μL of T4 Rnl2 buffer, 1 μL of SUPERase inhibitor, 6 μL of PEG8000, and 1 μL of T4 RNA ligase 2 (truncated). Incubate for 3 h at 25 °C.
3. Add 30 μL of RNase-free water to each sample and purify ligated product using Zymo RNA Clean and Concentrator-5 kit. Note that the kit efficiently removes un-ligated linker. At the final step, elute the ligated product in 11 μL of 10 mM Tris-HCl (pH 8.0) solution. Final elute is -10 μL.

Reverse Transcription and Circularization

1. Add 2 μL of 0.25 μM reverse transcription primer to 10 μL of linker-ligated product.
2. Denature the sample for 2 min at 80 °C and immediately cool on ice.
3. Add 8 μL of reverse transcription master mix to each sample to make a 20 μL reaction.
4. Hydrolyze the RNA template by adding 2.2 μL of 1 N NaOH and incubate at 90 °C for 20 min.
5. Precipitate the RT product. Add 20 μL of 3 M sodium acetate (pH 5.5), 2 μL of Glycoblue, 156 μL distilled water, 300 μL isopropanol and precipitate on dry ice for 30 min. Pellet the DNA for 30 min as described and resuspend the RT product in 10 μL distilled water.
6. Prepare 2 μL of 0.25 μM RT primer (mixed with 3 μL of 10 nM Tris-HCl, pH 8.0 and 5 μL of 2× denaturing loading buffer) and 1 μL of 10 bp DNA ladder as control. Add 10 μL of 2× loading buffer to each sample. Denature the sample at 80 °C for 90 s and immediately cool on ice.
7. Separate the sample using 7.5% polyacrylamide TBE-Urea gel in 0.5× TBE buffer with constant power (5 W) for 1 h and 30 min until the bromophenol blue dye reaches the end of the gel.
8. Excise the reverse transcription product and transfer it to nonsticky eppi-tubes. 2 mL tubes are recommended for DNA extraction. Avoid any contamination from un-extended RT primer.
9. Overnight gel extraction. Add 400 μL of DNA gel extraction buffer and freeze the sample for 30 min on dry ice. Leave the sample overnight (15 h is recommended) with gentle mixing. Collect the entire solution and transfer it to non-stick RNasefree tubes. Precipitate the footprints by adding 2 μL Glycoblue and 500 μL isopropanol. Carry out precipitation for 30 min or more on dry ice. Centrifuge at maximum speed at 4 °C for 30 min using tabletop centrifuge. Completely remove the supernatant and allow air dry for 5 min. Dissolve the footprint in 15 μL of 10 mM Tris-HCl (pH 8.0).
10. Mix 15 μL of gel extracted product, 2 μL of CircLigase buffer, 1 μL of 1 mM ATP, 1 μL of 10 nM MnCl2 and 1 μL of CircLigase per sample, incubate the reaction for 1 h at 60 °C and heat-inactivate for 10 min at 80 °C.
11. Recover the final DNA product. Add 74 μL distilled water, 6 μL of 5 M NaCl, 2 μL of Glycoblue and 150 μL of isopropanol and precipitate DNA.
12. Resuspend the circularized DNA product in 5 μL of 10 mM Tris-HCl (pH 8.0).

Indexed Library Generation

1. PCR amplification of circularized library using Phusion polymerase according to the manufacturer's instruction. Run several different cycles for comparison. Use different barcode reverse primers for different samples with a single forward primer.
2. Add 6× DNA loading dye to each PCR product and prepare 1 μL 10 bp DNA ladder as control.
3. Separate the PCR products using 8% polyacrylamide TBE gel, for 90 min at 180 V in 0.5× TBE buffer, or until the bromophenol blue dye reaches to two-third of the gel.
4. Stain the gel for 3 min in 1× SYBR Gold in 0.5× TBE buffer.
5. Excise the ~175 nt PCR product from the gel and carefully exclude slowly migrating smeared band (above 200 nt), bands from un-extended reverse transcription (~145 nt), template and primer (less than 100 nt).
6. Overnight gel extraction. Add 400 μL of DNA gel extraction buffer and freeze the sample for 30 min on dry ice. Leave the sample overnight (15 h is recommended) with gentle mixing. Collect the entire solution and transfer it to non-stick RNase-free eppitubes. Precipitate the footprints by adding 2 μL Glycoblue and 500 μL isopropanol. Carry out precipitation for 30 min or more on dry ice. Centrifuge at maximum speed at 4 °C for 30 min using tabletop centrifuge. Completely remove the supernatant and air dry for 5 min. Dissolve the footprint in 15 μL of 10 mM Tris-HCl (pH 8.0).
7. Measure DNA concentration and combine equal amount from each sample for Illumina sequencing.

Sequencing with Illumina Platforms

This protocol results in a fully functional sequencing library compatible with Illumina HiSeq, MiSeq, and NextSeq sequencing platforms. The user should follow standard procedures recommended by Illumina for adjusting the final library concentration for loading onto an Illumina flow-cell and sequencing appropriate for the sequencer used.

Conclusion

Ribosome profiling provides a high-resolution, comprehensive view of translation by precisely mapping ribosome locations on messenger RNAs (mRNAs). This technique serves as a robust instrument for quantitatively analyzing gene expression patterns, uncovering new translational phenomena, and scrutinizing the dynamics of protein synthesis. The unique capacity of ribosome profiling to monitor translation in real-time, while furnishing precise quantitative information on ribosome abundance and effectiveness, renders it an essential asset for inquiries into functional genomics, translational regulation, and the modulation of gene expression across diverse biological circumstances.

Reference:

1. Jin H Y, Xiao C. An integrated polysome profiling and ribosome profiling method to investigate in vivo translatome//Next Generation Sequencing. Humana Press, New York, NY, 2018: 1-18.