Name: sample preparation for mass spectrometry-based identification of rna-binding regions
Uploaded: 2017-09-28T14:00:00.000+00:00
Duration: 10 min 52 s
Description: Watch this Scientific Journal Video about Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions at JoVE.com

R.B. was supported by the Searle Scholars Program, the W.W. Smith Foundation (C1404), and the March of Dimes Foundation (1-FY-15- 344). B.A.G acknowledges support from NIH grants R01GM110174 and NIH R01AI118891, as well as DOD grant BC123187P1. R.W.-T. was supported by NIH training grant T32GM008216.

1. Culture and Expansion of mESCs
NOTE: Mouse embryonic stem cells are easy to culture and can be quickly expanded to the large numbers required by biochemical experiments thanks to their fast cycling time. Healthy mESCs double every 12 h.
<ol>
	<li>Expand mESCs to the desired number on gelatinized plates in mESC medium (see below) in a tissue culture incubator kept at 37 &#176;C, 5% CO2, and &#62;95% humidity. 
	NOTE: Enough plates should be prepared for 3 - 5 biological replicates for the +4SU sample and the -4SU control. One 10-cm plate (i.e. 5 - 10 million cells) is more than sufficient for each biological replicate.</li>
	<li>The medium for mESCs is composed of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) supplemented with 15% fetal bovine serum (FBS), 2 mM L-glutamine, 0.5x penicillin/streptomycin, 1x non-essential amino acids, 100 &#181;M beta-mercaptoethanol, 1,000 U/mL leukemia inhibitory factor (LIF), 3 &#181;M CHIR99021, 1 &#181;M PD0325901.</li>
	<li>Before passaging mESCs, prepare the desired number of 10 cm plates.
	<ol>
		<li>Add 4 mL of 0.1% gelatin in water making sure that the entire surface of the plate is covered; incubate for 10 min at room temperature (RT); remove gelatin solution.</li>
	</ol>
	</li>
	<li>Passage cells when they reach ~10 million cells per plate, typically ~48 h following inoculation. 
	NOTE: As mESCs grow in dense colonies, plates should never be allowed to reach 100% confluency as observed for cell lines that grow in monolayers (e.g. HEK293 or 3T3 cells). An mESC plate should be passaged when ~50% confluent and definitely before the medium turns yellow.
	<ol>
		<li>To detach cells, wash plates with dense mESC cultures once in 1x phosphate buffered saline (PBS) supplemented with 2 mM EDTA, then add 0.05% trypsin dissolved in DMEM and return plate to incubator for 5 min (37 &#176;C, 5% CO2, and &#62;95% humidity).</li>
		<li>Remove plate from incubator and resuspend cells with equal volume of DMEM by pipetting up and down vigorously several times.</li>
		<li>Count cells using hemocytometer and ensure that cells form a single-cell suspension (i.e. no clusters), then centrifuge cell mixture 500 x g for 5 min at room temperature. Remove supernatant and resuspend cell pellet in 1 - 2 mL DMEM.</li>
		<li>Inoculate cells at a density of 10,000 cells/cm2 in mESC medium (i.e. ~800,000 cells per 10-cm plate) using the gelatin-coated plates prepared in step 1.3.1.</li>
	</ol>
	</li>
</ol>
2. Crosslink of Protein&#8211;RNA Interactions in Live Cells
NOTE: RNA-protein crosslinking is mediated by the photo-activatable ribonucleoside analog 4-thiouridine (4SU). 4SU has a longer absorbance maximum than endogenous nucleotides and can only be incorporated into RNA; therefore intermediate-wavelength UVB can be used to selectively crosslink RNA to proteins25,26. UVB treatment of 4SU-treated cells leads to covalent crosslinks between 4SU-containing RNA and amino acids, with a reported preference for Tyr, Trp, Met, Lys, and Cys27.
<ol>
	<li>Expand mESCs to the required number of 10-cm plates as described in step 1. 
	NOTE: Enough plates should be prepared for 3 - 5 biological replicates for the +4SU sample and the -4SU control. One 10-cm plate (i.e. 5 - 10 million cells) is sufficient for each biological replicate.</li>
	<li>Before the plates reach confluency, add 4SU directly into medium to a final concentration of 500 &#181;M. Leave -4SU control dishes untreated.</li>
	<li>Shake plates gently to distribute the 4SU homogeneously throughout the medium. Return them to the same tissue culture incubator for 2 h (37 &#176;C, 5% CO2, and &#62;95% humidity).</li>
	<li>Transfer plates to an ice bucket and discard medium.</li>
	<li>Rinse the plate gently with 5 mL of ice-cold PBS. 
	NOTE: Rinse gently or cells might detach.</li>
	<li>Add 2 mL of ice-cold PBS and place plates without lids in a crosslinker equipped with UVB-emitting bulbs (312 nm). Irradiate plates at an energy setting of 1 J/cm2. 
	NOTE: It is critical that plastic lids are removed or they might shield the cells from UV light and prevent crosslinking.</li>
	<li>Collect cells using cell lifters and transfer to ice-cold PBS in conical tubes. Centrifuge at 2,000 x g for 5 min, 4 &#176;C.</li>
	<li>Remove the supernatant and resuspend cell pellet in 5 mL ice-cold PBS with 2 mM EDTA and 0.2 mM phenylmethylsulfonyl fluoride (PMSF).</li>
	<li>Count cells using a hemocytometer. Expect 5 - 10 and 10 - 20 million cells from 10-cm and 15-cm plates respectively.</li>
	<li>Centrifuge at 2,000 x g for 5 min, 4 &#176;C. 
	NOTE: Pelleted cells can be immediately used for step 3 or they can be flash-frozen in liquid nitrogen and stored at -80 &#176;C indefinitely. This is a potential stopping point.</li>
</ol>
3. Isolation of Nuclei
NOTE: Nuclei are isolated to remove cytoplasmic proteins and increase coverage of nuclear proteins. This step can be replaced with other forms of cellular fractionation to study RBPs in different cellular compartments.
<ol>
	<li>Prepare ice cold Buffer A (10 mM Tris-HCl pH 7.9, 4 &#176;C, 1.5 mM MgCl2, 10 mM KCl). Use at least 2.5 mL per 10 million cells (as counted in step 2.9) for extraction. 
	NOTE: Stocks of Buffer A (e.g. a 500 mL bottle) can be prepared in advance and stored at 4 &#176;C for 6 months.
	<ol>
		<li>Prior to use, add 0.5 mM dithiothreitol (DTT) and 0.2 mM PMSF to the desired amount (2.5 mL per 10 million cells) of ice-cold buffer A.</li>
	</ol>
	</li>
	<li>Wash cells using 2 mL Buffer A per 10 million cells. Spin at 2,500 x g for 5 min at 4 &#176;C.</li>
	<li>Discard supernatant, add Buffer A supplemented with 0.2% of a nonionic, non-denaturing detergent, such as octyl-phenoxy-polyethoxy-ethanol (see the Table of Materials), 500 &#181;L per 10 million cells. Rotate for 5 min at 4 &#176;C.</li>
	<li>Centrifuge at 2,500 x g for 5 min.</li>
	<li>Remove the supernatant; the pellet comprises intact nuclei devoid of cytoplasm. 
	NOTE: Nuclei can be used immediately for step 4 or flash-frozen in liquid nitrogen and stored at -80 &#176;C indefinitely. This is a potential stopping point.</li>
</ol>
4. Lysis of Nuclei
NOTE: Crosslinked nuclei are lysed in a mass spectrometry-compatible buffer to release proteins and protein-RNA complexes.
<ol>
	<li>Add lysis buffer (9 M urea, 100 mM Tris-HCl pH 8.0, 24 &#176;C), 200 &#181;L per 10 million cells. Sonicate to shear chromatin using a 1/8&#34; probe at 50% amplitude setting for 5 - 10 s.</li>
	<li>Spin at 12,000 x g for 5 min and collect 175 &#181;L of supernatant to avoid the pellet. 
	NOTE: 100 &#181;L will be used for the next RBR-ID steps and the remaining 75 &#181;L of lysate can be stored at -80 &#176;C for later use and/or western blot analysis.</li>
	<li>Measure protein concentration via Bradford assay or similar technique28. Dilute protein concentration in the different samples to 1 mg/mL or the lowest sample concentration using appropriate amounts of lysis buffer. 
	NOTE: From the nuclei of 10 million mESCs expect 100 -&#160;200 &#181;g of protein.</li>
</ol>
5. Trypsin Digestion
NOTE: Proteins are digested to generate peptides suitable for bottom-up mass spectrometry (MS) analysis.
<ol>
	<li>Add DTT solution to nuclear lysate (5 mM final); incubate for 1 h at RT.</li>
	<li>Add iodoacetamide from fresh stock (14 mM final); incubate for 30 min at RT in the dark.</li>
	<li>Dilute lysate with 5 times volume of 50 mM NH4HCO3</li>
	<li>Add trypsin (&#181;g protein : &#181;g trypsin = 200 : 1); incubate at 37 &#176;C overnight.</li>
</ol>
6. Desalting of peptides
<ol>
	<li>Prepare one custom-made stage tip per sample.
	<ol>
		<li>6.1.1. Place a disk of solid phase extraction C18 material into the bottom of a 200 &#181;L pipette tip.</li>
		<li>Add 50 &#181;L of oligo R3 reversed phase resin on top of the C18 disk. Place stage tip inside centrifugation adaptor and the place tip with adaptor inside a 1.5 mL microfuge tube.</li>
		<li>Add 100 &#181;L 100% acetonitrile to tips to wash. Centrifuge at 1000 x g for 1 min to remove the solvent.</li>
		<li>Equilibrate with 50 &#181;L 0.1% trifluoroacetic acid (TFA). Centrifuge at 1000 x g for 1 min.</li>
	</ol>
	</li>
	<li>Add 100% formic acid to digested protein sample to decrease the pH to 2 - 3. 
	NOTE: This should require ~5 &#181;L per 1 mL of diluted peptide sample, but pH should be measured and more formic acid added if necessary.</li>
	<li>Load sample onto custom-made stage tip, centrifuge at 1000 x g for 1 min until all of sample goes through the tip.</li>
	<li>Wash bound peptides with 50 &#181;L 0.1% TFA. Centrifuge at 1000 x g for 1 min.</li>
	<li>Elute peptides by adding 50 &#181;L elution buffer (75% acetonitrile, 0.025% TFA) to stage tip and centrifuge at 1000 x g for 1 min to force the elution buffer out of the tip.</li>
	<li>Dry the sample using a vacuum centrifuge set at 150 x g and 1 - 3 kPa for 1 h. 
	NOTE: Dried peptides can be stored at -80 &#176;C indefinitely. This is a potential stopping point.</li>
</ol>
7. Removal of Crosslinked RNA
NOTE: Treat peptides with nuclease to remove crosslinked RNA.
<ol>
	<li>Resuspend the pellet in 50 &#181;L 2x nuclease buffer (100 mM NH4HCO3, 4 mM MgCl2).</li>
	<li>Measure peptide concentration using absorbance at 280 nm assuming 1 mg/mL of peptides for an A280 of 1. 
	NOTE: The expected concentration is 1 mg/mL.</li>
	<li>Adjust all samples to 1 mg/mL or to the lowest concentration using 2x nuclease buffer.</li>
	<li>Prepare a master mix of 50 &#181;L H2O + 1 &#181;L high purity nuclease (&#8805;250 U) (refer to the Table of Materials) per sample.</li>
	<li>Take 50 &#181;L of peptide sample and add 50 &#181;L of H2O + nuclease master mix.</li>
	<li>Incubate at 37 &#176;C for 1 h. Proceed to perform mass spectrometry. 
	NOTE: The peptides are now ready for mass spectrometry. They can be analyzed immediately or stored at -80 &#176;C indefinitely. This is a potential stopping point.</li>
</ol>
8. Nano Liquid Chromatography, Mass Spectrometry, and Raw Data Processing
NOTE: Because 4SU-crosslinking changes the mass of the peptide, their ions do not count toward the intensity of the non-crosslinked peptide during LC-MS/MS, which therefore appears to be decreased by the crosslinking. The degree and consistency of this decrease reflects the degree of protein-RNA crosslinking for each peptide24.
<ol>
	<li>Prepare HPLC buffers as follows: 0.1% formic acid in HPLC-grade water (buffer A); 0.1% formic acid in HPLC-grade acetonitrile (Buffer B).</li>
	<li>If a nano-HPLC is available, connect a nano-column with the following specifications: internal diameter 75 &#181;m; packed with C18-AQ particles; length 20 - 25 cm. 
	NOTE: If the HPLC available runs at higher flow rates use a proper column size for the indicated flow rate. Higher flow chromatography decreases sensitivity.</li>
	<li>Program the HPLC method as follows: flow rate 250 - 300 nL/min; gradient from 2 to 28% buffer B in 120 min, from 28 to 80% B for the next 5 min and isocratic 80% B for 10 min. 
	NOTE: If the HPLC does not have an automated column equilibration prior sample loading, start the program with 10 min at 0% B before loading the sample.</li>
	<li>Program the MS acquisition method to perform data-dependent acquisition (DDA) as standard shotgun proteomics experiments. The instrument duty cycle should alternate full MS scans with tandem MS scans (or MS/MS) of the most intense signals. Use the optimal settings for proteomics MS runs. 
	NOTE: For confident results, use instruments that can perform at least the full MS scan in high resolution mode (e.g. orbitrap, time-of-flight). For accurate quantification, make sure that intervals between full MS scans are no longer than 3 s. Longer duty cycles might prevent accurate definition of extracted ion chromatograms.</li>
	<li>Load 1 - 2 &#181;g of sample onto the HPLC column and run the HPLC-MS/MS method as programmed. For each biological replicate perform at least two independent runs and treat them as technical replicates.</li>
	<li>After MS, import the raw files into a proteomics software package for MS/MS spectra identification and peptide quantification via extracted ion chromatography. We recommend software packages that can perform chromatographic alignment of multiple MS runs (see the Table of Materials)29,30.</li>
	<li>If the option is available in the software suite, enable &#34;Match Between Runs&#34; prior to starting data processing.</li>
	<li>Once the analysis is finished, export the peptide list together with the quantification values.</li>
</ol>

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M., Nagai, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystal+structure+of+human+U1+snRNP,+a+small+nuclear+ribonucleoprotein+particle,+reveals+the+mechanism+of+5'+splice+site+recognition.">Crystal structure of human U1 snRNP, a small nuclear ribonucleoprotein particle, reveals the mechanism of 5' splice site recognition.</a> Elife. 4, (2015).</li><li>Hafner, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PAR-CliP--a+method+to+identify+transcriptome-wide+the+binding+sites+of+RNA+binding+proteins.">PAR-CliP--a method to identify transcriptome-wide the binding sites of RNA binding proteins.</a> J Vis Exp. (41), (2010).</li></ol>

The authors have nothing to disclose.

We describe a detailed experimental protocol to perform RBR-ID in mESCs and, with appropriate modifications, in any cell that can incorporate 4SU into RNA. Other cell types may require optimization of the approach to ensure a sufficient signal to noise ratio. Additionally, while the protocol described herein focuses on the examination of nuclear RBPs, the RBR-ID technology should be easily adapted to different cellular compartments, such as the cytosol or specific organelles, by use of different fractionation strategies. Parameters that may require optimization include 4SU concentration and incorporation time as well as the energy of UV crosslinking. These parameters are important to ensure efficient formation of RNA-protein crosslinks and yield a satisfactory signal to noise ratio. We have shown that 312 nm UVB light is much more efficient at crosslinking 4SU-containing RNAs to proteins compared to the 365 nm UVA typically used in PAR-CLIP32. Direct comparison of RBR-ID performed with 312 nm and 365 nm demonstrated that 312 nm UVB resulted in the identification of a much larger portion of known RBPs24.
Two other methods are available to perform proteome-wide RBP identification. One, called RBDmap, is similar to RBR-ID in that it utilizes UV crosslinking and MS. RBDmap was used to identify RBPs and map their RNA-binding activities in HeLa cells23. This method relies on positive identification of peptides adjacent to the RNA-binding sites and relies on two sequential oligo-dT pull-downs, suggesting that it might have a lower false positive rate than RBR-ID. However, the pull-downs allow only the identification of RBPs that bind to polyA+ RNA and require large amounts of input material, up to 10 - 100 times more than RBR-ID. RBR-ID can be performed with as little as 5 &#181;g of protein per biological replicate (~2 &#181;g per technical replicate), which can be collected from as few as 250,000 cells.
The second method, termed SONAR, relies on data mining of large protein-protein interaction databases to detect proteins that frequently co-purify with known RBPs and whose interaction with these RBPs might therefore be mediated by RNA18. This clever approach is very powerful and allows for the identification of novel RBPs without performing any wet lab experiment, provided that suitable databases are available; however, it cannot reveal the interaction sites and therefore is complementary but not alternative to RBR-ID.
In summary, our RBR-ID technique can identify novel RBPs and map their RBRs with limited input sample amounts and without requiring any biochemical purification of the protein-RNA crosslinked complexes. The technique makes no assumption on the type of RNA bound by the identified RBPs and therefore can map the RNA binding activity of proteins that bind to non-polyadenylated, many of which are noncoding, suggesting that RBR-ID might prove a useful technique to study the regulatory roles of noncoding RNAs.

The purpose of the RBR-ID method is to identify novel RNA-binding proteins (RBPs) and map their RNA-binding regions (RBRs) with peptide-level resolution to facilitate the design of RNA-binding mutants and the investigation of the biological and biochemical functions of protein-RNA interactions.
RNA is unique among biomolecules as it can both act as a messenger carrying genetic information and also fold into complex three-dimensional structures with biochemical functions more akin to those of proteins1,2. A growing body of evidence suggests that noncoding RNAs (ncRNAs) play important roles in various gene regulatory and epigenetic pathways3,4,5 and, typically, these regulatory functions are mediated in concert with proteins that interact specifically with a given RNA. Of particular relevance, a set of interacting proteins was recently identified for the intensely studied long ncRNA (lncRNA) Xist, providing valuable insight into how this lncRNA mediates X-chromosome inactivation in female cells6,7,8. Notably, several of these Xist-interacting proteins do not contain any canonical RNA-binding domains9, and therefore their RNA-binding activity could not be predicted in silico based on their primary sequence alone. Considering that thousands of lncRNAs are expressed in any given cell10, it is reasonable to assume that many of them might act via interactions with yet to be discovered RNA-binding proteins (RBPs). An experimental strategy to identify these novel RBPs would therefore greatly facilitate the task of dissecting the biological function of ncRNAs.
Previous attempts to identify RBPs empirically have relied on polyA+ RNA selection coupled to mass spectrometry (MS)11,12,13,14,15. Although these experiments added many proteins to the list of putative RBPs, by design they could only detect proteins bound to polyadenylated transcripts. However, most small RNAs and many lncRNAs are not polyadenylated.16,17 and their interacting proteins would likely have been missed in these experiments. A recent study applied machine learning to protein-protein interactome databases to identify proteins that co-purified with multiple known RBPs and showed that these recurrent RBP partners were more likely to possess RNA-binding activities18. However, this approach relies on mining existing large interaction databases and can only identify proteins that can be co-purified in non-denaturing conditions with known RBPs, thus excluding from the analysis insoluble, membrane-embedded, and scarce proteins.
The identification of a protein as a bona fide RBP often does not automatically yield information on the biological and/or biochemical function of the protein-RNA interaction. To address this point, it is typically desirable to identify the protein domain and amino acid residues involved in the interaction so that specific mutants can be designed to test the function of RNA binding in the context of each novel RBP19,20. Previous efforts by our group and others have used recombinant protein fragments and deletion mutants to identify RNA binding regions (RBRs)19,20,21,22; however, such approaches are labor-intensive and incompatible with high-throughput analyses. More recently, a study described an experimental strategy to map RNA-binding activities in a high-throughput fashion using mass spectrometry23; however, this approach relied on a double polyA+ RNA selection, and thus carried the same limitations as the RBP identification approaches described above.
We developed a technique, termed RNA binding region identification (RBR-ID), which exploits protein-RNA photocrosslinking and quantitative mass spectrometry to identify proteins and protein regions interacting with RNA in live cells without making assumption on the RNA polyadenylation status, thus including RBPs bound to polyA- RNAs24. Moreover, this method relies exclusively on crosslinking and has no requirements on protein solubility or accessibility and is thus suitable to map RNA-binding activities within membranes (e.g. the nuclear envelope) or poorly soluble compartments (e.g. the nuclear matrix). We describe the experimental steps to perform RBR-ID for the nuclei of mouse embryonic stem cells (mESCs) but with minor modifications this protocol should be suitable for a variety of cell types, provided that they can efficiently incorporate 4SU from the culture medium.

<table><tbody><tr><td>KnockOut DMEM</td><td>Fisher Scientific</td><td>10829018</td><td></td></tr><tr><td>Fetal bovine serum, qualified, US origin</td><td>Fisher Scientific</td><td>26140079</td><td></td></tr><tr><td>L-Glutamine solution 200 mM</td><td>Sigma</td><td>G7513</td><td></td></tr><tr><td>Penicillin-Streptomycin solution</td><td>Sigma</td><td>P0781</td><td></td></tr><tr><td>MEM Non-essential Amino Acid Solution (100&amp;times;)</td><td>Sigma</td><td>M7145</td><td></td></tr><tr><td>2-Mercaptoethanol</td><td>Sigma</td><td>M3148</td><td></td></tr><tr><td>ESGRO Leukemia Inhibitory Factor (LIF)</td><td>EMD Millipore</td><td>ESG1106</td><td></td></tr><tr><td>CHIR99021</td><td>Tocris</td><td>4423</td><td></td></tr><tr><td>PD0325901</td><td>Sigma</td><td>PZ0162</td><td></td></tr><tr><td>Gelatin solution,2% in water</td><td>Sigma</td><td>G1393</td><td></td></tr><tr><td>4-thiouridine</td><td>Sigma</td><td>T4509</td><td>50 mM stock in water</td></tr><tr><td>Spectrolinker XL-1500</td><td>Fisher Scientific</td><td>11-992-90</td><td></td></tr><tr><td>Phenylmethanesulfonyl fluoride</td><td>Sigma</td><td>78830</td><td></td></tr><tr><td>IGEPAL CO-630</td><td>Sigma</td><td>542334</td><td>Commercial form of octyl-phenoxy-polyethoxy-ethanol detergent</td></tr><tr><td>Iodoacetamide</td><td>Sigma</td><td>I6125</td><td></td></tr><tr><td>Trypsin, sequencing grade</td><td>Promega</td><td>V5111</td><td></td></tr><tr><td>Empore solid phase extraction disk</td><td>3M</td><td>66883</td><td></td></tr><tr><td>OLIGO R3 Reversed - Phase Resin</td><td>Fisher Scientific</td><td>1133903</td><td></td></tr><tr><td>Benzonase</td><td>Sigma</td><td>E8263</td><td>High purity nuclease</td></tr><tr><td>Sonic Dismembrator Model 100</td><td>Fisher Scientific</td><td>discontinued</td><td>updated with FB505110</td></tr><tr><td>HPLC grade acetonitrile</td><td>Fisher Chemical</td><td>A955-4</td><td></td></tr><tr><td>HPLC grade water</td><td>Fisher Scientific</td><td>W6 4</td><td></td></tr><tr><td>TFA</td><td>Fisher Scientific</td><td>A11650</td><td></td></tr><tr><td>Ammonium Bicarbonate</td><td>Sigma</td><td>A6141</td><td></td></tr><tr><td>Acetic Acid</td><td>Sigma</td><td>49199</td><td></td></tr><tr><td>Formic Acid</td><td>Sigma</td><td>F0507</td><td></td></tr><tr><td>ReproSil-Pur 18-AQ</td><td>Dr. Maisch GmbH HPLC</td><td>r13.aq.0003</td><td>Packing material for HPLC column</td></tr><tr><td>Capillary for nano columns (75 &amp;micro;m)</td><td>Molex</td><td>1068150017</td><td></td></tr><tr><td>MaxQuant software</td><td>Max Planck Institute for Biochemistry</td><td></td><td>Can perform chromatographic alignment of multiple MS runs</td></tr></tbody></table>

sample preparation for mass spectrometry-based identification of rna-binding regions

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most cases, the action of noncoding RNAs is mediated by proteins whose functions are in turn regulated by these interactions with noncoding RNAs. Consistent with this, a growing number of proteins involved in nuclear functions have been reported to bind RNA and in a few cases the RNA-binding regions of these proteins have been mapped, often through laborious, candidate-based methods.
Here, we report a detailed protocol to perform a high-throughput, proteome-wide unbiased identification of RNA-binding proteins and their RNA-binding regions. The methodology relies on the incorporation of a photoreactive uridine analog in the cellular RNA, followed by UV-mediated protein-RNA crosslinking, and mass spectrometry analyses to reveal RNA-crosslinked peptides within the proteome. Although we describe the procedure for mouse embryonic stem cells, the protocol should be easily adapted to a variety of cultured cells.

Figure 1 depicts the RBR-ID workflow. Due to the relatively low crosslinking efficiency of this technique, it is very important to consider both the depletion level and consistency of the observed effect (P-value) across biological replicates. Figure 2 shows a volcano plot of RBR-ID result. Peptides that overlapped RNA recognition motif (RRM) domain show highly consistent depletion level. RRM domains can be used as a positive control for RBR-ID analysis.
RBR-ID can be used to map RBRs in live cells. An RBR-ID score can be calculated for each peptide to estimate RNA-binding potential: RBR-ID score = -log2(normalized +4SU intensity/normalized -4SU intensity) x (log10(P-value))2. Figure 3 shows a heatmap of RBR-ID scores projected on the surface of the spliceosomal subunit U1-70k. The bright red color in vicinity of the RNA contact, as determined from the crystal structure, indicates correct identification of the protein-RNA interaction.
Upon identification of novel RBPs and their RBRs, it is highly recommended that their RNA-binding activity be confirmed by an independent method. In our previous study24, we identified TET2 as a novel RBP and mapped the RNA-binding activity to a C-terminal region, as shown in Figure 4A by plotting the RBR-ID score along the primary sequence of the protein. Figure 4B shows that the requirement for this novel RBR could be verified by performing PAR-CLIP26 using the WT sequence and comparing the signal to that of a mutant lacking the predicted RBR. Additional control and more detailed explanation of this validation experiment are available in He et al. 201624.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56004/56004fig1.jpg" /> 
Figure 1: RBR-ID overview. Mouse ESCs are treated with 4SU or not (1-2) and irradiated with 312 nm UV (3). Nuclei are isolated (4) and extracts digested with protease and nuclease, yielding both crosslinked and uncrosslinked peptides (5). Covalent RNA adducts at crosslink sites alter the peptide mass, leading to a corresponding decrease in intensity of the uncrosslinked peptide&#39;s mass spectrum (6, arrow). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56004/56004fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56004/56004fig2.jpg" /> 
Figure 2: Proteome-wide RBR-ID in mESCs. Volcano plots showing log-fold changes in peptide intensities on the x axis and Student&#39;s t-test P-values on the y axis for &#177; 4SU treatments (312 nm). Peptides overlapping annotated RRM domains are in blue. An RNA-binding peptide from HNRNPC is highlighted in red. Previously published in He et al. 201624. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56004/56004fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56004/56004fig3.jpg" /> 
Figure 3: RBR-ID maps the sites of protein&#8211;RNA interactions. Zoomed-in regions of the crystal structure of U1 snRNP (PDB ID: 4PJO31) showing protein surfaces color-coded according to their RBR-ID score and interacting RNAs for U1-70K. Previously published in He et al. 201624. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56004/56004fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56004/56004fig4.jpg" /> 
Figure 4: Validation of the RBR of TET2. (A) Primary sequence and known domains for TET2; smoothed residue-level RBR-ID score plotted along the primary sequence (middle); and scheme of epitope-tagged catalytic domain fragment (CD) and RBR-deleted (CD&#916;RBR) constructs used for validation (bottom). (B) PAR-CLIP of transiently expressed TET2 CD and &#916;CD&#916;RBR in HEK293 cells. Autoradiography for 32P-labeled RNA (top) and control western blot (bottom). Previously published in He et al. 201624. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56004/56004fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions at JoVE.com

Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

We describe a protocol to identify RNA-binding proteins and map their RNA-binding regions in live cells using UV-mediated photocrosslinking and mass spectrometry.

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most ...

sample-preparation-for-mass-spectrometry-based-identification-rna

Nanyang Technological University

Research

JoVE Journal

Biochemistry

8.1K Views.  University of Pennsylvania Perelman School of Medicine. We beschrijven een protocol te identificeren van RNA-bindende proteÃ¯nen en kaart van de regio's hun RNA-bindende in levende cellen met behulp van UV-gemedieerde photocrosslinking en massaspectrometrie.

Video: Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

Novel RNA-Binding Proteins Isolation by the RaPID Methodology

RNA-binding proteins (RBPs) play important roles in every aspect of RNA metabolism and regulation. Their identification is a major challenge in modern biology. Only a few in vitro and in vivo methods enable the identification of RBPs associated with a particular target mRNA. However, their main limitations are the identification of RBPs in a non-cellular environment (in vitro) or the low efficiency isolation of RNA of interest (in vivo). An RNA-binding protein purification and identification (RaPID) methodology was designed to overcome these limitations in yeast and enable efficient isolation of proteins that are associated in vivo. To achieve this, the RNA of interest is tagged with MS2 loops, and co-expressed with a fusion protein of an MS2-binding protein and a streptavidin-binding protein (SBP). Cells are then subjected to crosslinking and lysed, and complexes are isolated through streptavidin beads. The proteins that co-purify with the tagged RNA can then be determined by mass spectrometry. We recently used this protocol to identify novel proteins associated with the ER-associated PMP1 mRNA. Here, we provide a detailed protocol of RaPID, and discuss some of its limitations and advantages.

RNA-protein interactions lie at the heart of many cellular processes. Here, we describe an in vivo method to isolate specific RNA and identify novel proteins that are associated with it. This could shed new light on how RNAs are regulated in the cell.

RNA-binding proteins (RBPs) play important roles in every aspect of RNA metabolism and regulation. Their identification is a major challenge in modern ...

Genetics

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies have revealed that these reversible RNA modifications affect RNA splicing, translation, degradation, and localization. Multiple physiological processes, like circadian rhythms, stem cell pluripotency, fibrosis, triglyceride metabolism, and obesity are also controlled by m6A modifications. Immunoprecipitation/sequencing, mass spectrometry, and modified northern blotting are some of the methods commonly employed to measure m6A modifications. Herein, we present a northeastern blotting technique for measuring m6A modifications. The current protocol provides good size separation of RNA, better accommodation and standardization for various experimental designs, and clear delineation of m6A modifications in various sources of RNA. While m6A modifications are known to have a crucial impact on human physiology relating to circadian rhythms and obesity, their roles in other (patho)physiological states are unclear. Therefore, investigations on m6A modifications have immense possibility to provide key insights into molecular physiology.

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

A modified northern blotting method for measuring N6-methyladenosine (m6A) modifications in RNA is described. The current method can detect modifications in diverse RNAs and controls under various experimental designs.

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies ...

RNA Pull-down Procedure to Identify RNA Targets of a Long Non-coding RNA

Long non-coding RNA (lncRNA), which are sequences of more than 200 nucleotides without a defined reading frame, belong to the regulatory non-coding RNA&#39;s family. Although their biological functions remain largely unknown, the number of these lncRNAs has steadily increased and it is now estimated that humans may have more than 10,000 such transcripts. Some of these are known to be involved in important regulatory pathways of gene expression which take place at the transcriptional level, but also at different steps of RNA co- and post-transcriptional maturation. In the latter cases, RNAs that are targeted by the lncRNA have to be identified. That&#39;s the reason why it is useful to develop a method enabling the identification of RNAs associated directly or indirectly with a lncRNA of interest.
This protocol, which was inspired by previously published protocols allowing the isolation of a lncRNA together with its associated chromatin sequences, was adapted to permit the isolation of associated RNAs. We determined that two steps are critical for the efficiency of this protocol. The first is the design of specific anti-sense DNA oligonucleotide probes able to hybridize to the lncRNA of interest. To this end, the lncRNA secondary structure was predicted by bioinformatics and anti-sense oligonucleotide probes were designed with a strong affinity for regions that display a low probability of internal base pairing. The second crucial step of the procedure relies on the fixative conditions of the tissue or cultured cells that have to preserve the network between all molecular partners. Coupled with high throughput RNA sequencing, this RNA pull-down protocol can provide the whole RNA interactome of a lncRNA of interest.

This RNA pull-down method allows identifying the RNA targets of a long non-coding RNA (lncRNA). Based on the hybridization of home-made, designed anti-sense DNA oligonucleotide probes specific to this lncRNA in an appropriately fixed tissue or cell line, it efficiently allows the capture of all RNA targets of the lncRNA.

Long non-coding RNA (lncRNA), which are sequences of more than 200 nucleotides without a defined reading frame, belong to the regulatory non-coding ...

Exploring Sequence Space to Identify Binding Sites for Regulatory RNA-Binding Proteins

Gene regulation plays an important role in all cells. Transcriptional, post-transcriptional (or RNA processing), translational, and post-translational steps are used to regulate specific genes. Sequence-specific nucleic acid-binding proteins target specific sequences to control spatial or temporal gene expression. The binding sites in nucleic acids are typically characterized by mutational analysis. However, numerous proteins of interest have no known binding site for such characterization. Here we describe an approach to identify previously unknown binding sites for RNA-binding proteins. It involves iterative selection and amplification of sequences starting with a randomized sequence pool. Following several rounds of these steps-transcription, binding, and amplification-the enriched sequences are sequenced to identify a preferred binding site(s). Success of this approach is monitored using in vitro binding assays. Subsequently, in vitro and in vivo functional assays can be used to assess the biological relevance of the selected sequences. This approach allows identification and characterization of a previously unknown binding site(s) for any RNA-binding protein for which an assay to separate protein-bound and unbound RNAs exists.

Sequence specificity is critical for gene regulation. Regulatory proteins that recognize specific sequences are important for gene regulation. Defining functional binding sites for such proteins is a challenging biological problem. An iterative approach for identification of a binding site for an RNA-binding protein is described here and is applicable to all RNA-binding proteins.

Gene regulation plays an important role in all cells. Transcriptional, post-transcriptional (or RNA processing), translational, and post-translational ...

Improving Small RNA-seq: Less Bias and Better Detection of 2'-O-Methyl RNAs

The study of small RNAs (sRNAs) by next-generation sequencing (NGS) is challenged by bias issues during library preparation. Several types of sRNA such as plant microRNAs (miRNAs) carry a 2&#39;-O-methyl (2&#39;-OMe) modification at their 3&#39;&#160;terminal nucleotide. This modification adds another difficulty as it inhibits 3&#39;&#160;adapter ligation. We previously demonstrated that modified versions of the &#39;TruSeq (TS)&#39;&#160;protocol have less bias and an improved detection of 2&#39;-OMe RNAs. Here we describe in detail protocol &#39;TS5&#39;, which showed the best overall performance. TS5 can be followed either using homemade reagents or reagents from the TS kit, with equal performance.

We present a detailed small RNA library reparation protocol with less bias than standard methods and an increased sensitivity for 2&#39;-O-methyl RNAs. This protocol can be followed using homemade reagents to save cost or using kits for convenience.

The study of small RNAs (sRNAs) by next-generation sequencing (NGS) is challenged by bias issues during library preparation. Several types of sRNA such as ...

2D-HELS MS Seq: A General LC-MS-Based Method for Direct and de novo Sequencing of RNA Mixtures with Different Nucleotide Modifications

Mass spectrometry (MS)-based sequencing approaches have been shown to be useful in&#160;direct&#160;sequencing&#160;RNA without the need for a complementary DNA (cDNA) intermediate. However, such approaches are rarely applied as a de novo RNA sequencing method, but used mainly as a tool that can assist in quality assurance for confirming known sequences of purified single-stranded RNA samples. Recently, we developed&#160;a direct RNA sequencing method by integrating a 2-dimensional mass-retention time hydrophobic end-labeling strategy into MS-based sequencing (2D-HELS MS Seq). This method is capable of accurately sequencing single RNA sequences as well as mixtures containing up to 12 distinct RNA sequences. In addition to the four canonical ribonucleotides (A, C, G, and U), the method has the capacity to sequence RNA oligonucleotides containing modified nucleotides. This is possible because the modified nucleobase either has an intrinsically unique mass that can help in its identification and its location in the RNA sequence, or can be converted into a product with a unique mass. In this study, we have used RNA, incorporating two representative modified nucleotides&#160;(pseudouridine (&#936;) and 5-methylcytosine (m5C)), to illustrate the application of the method for the de novo sequencing of a single RNA oligonucleotide&#160;as well as a mixture of RNA oligonucleotides, each with a different sequence and/or modified nucleotides. The procedures and protocols described here to sequence these model RNAs will be applicable to other short RNA samples (&#60;35 nt) when using a standard high-resolution LC-MS system, and can also be used for&#160;sequence verification of modified therapeutic RNA oligonucleotides.&#160;In the future, with the development of more robust algorithms and with better instruments, this method could allow sequencing of more complex biological samples.&#160;

Here, we describe a detailed protocol for an LC-MS-based sequencing method that can be used as a direct method to sequence short RNA (&#60;35 nt per run) without a cDNA intermediate, and as a general method to sequence different nucleotide modifications in a single study at single-base precision.

Mass spectrometry (MS)-based sequencing approaches have been shown to be useful in&#160;direct&#160;sequencing&#160;RNA without the need for a ...

Identification of RNAs Engaged in Direct RNA-RNA Interaction with a Long Non-Coding RNA

The growing role attributed nowadays to long non-coding RNAs (lncRNA) in physiology and pathophysiology makes it crucial to characterize their interactome by identifying their molecular partners, DNA, proteins and/or RNAs. The latter can interact with lncRNA through networks involving proteins, but they can also be engaged in direct RNA/RNA interactions. We, therefore, developed an easy-to-use RNA pull-down procedure that allowed identification of RNAs engaged in direct RNA/RNA interaction with a lncRNA using psoralen, a molecule that cross-links only RNA/RNA interactions. Bioinformatics modeling of the lncRNA secondary structure allowed the selection of several specific antisense DNA oligonucleotide probes with a strong affinity for regions displaying a low probability of internal base pairing. Since the specific probes that were designed targeted accessible regions throughout the length of the lncRNA, the RNA-interaction zones could be delineated in the sequence of the lncRNA. When coupled with a high throughput RNA sequencing, this protocol can be used for the whole direct RNA interactome studies of a lncRNA of interest.

An easy-to-use RNA pull-down protocol is designed for the identification of RNAs engaged in direct RNA/RNA interaction with a long non-coding RNA. The protocol uses psoralen as a fixative to cross-link only RNA/RNA interactions and provides the whole direct RNA interactome of a long non-coding RNA when coupled with RNA sequencing.

The growing role attributed nowadays to long non-coding RNAs (lncRNA) in physiology and pathophysiology makes it crucial to characterize their interactome ...

Practical Aspects of Sample Preparation and Setup of 1H R1&#961; Relaxation Dispersion Experiments of RNA

RNA is a highly flexible biomolecule, wherein changes in structures play crucial roles in the functions that RNA molecules execute as cellular messengers and modulators. While these dynamic states remain hidden to most structural methods, R1&#961; relaxation dispersion (RD) spectroscopy allows the study of conformational dynamics in the micro- to millisecond regime at atomic resolution. The use of 1H as the observed nucleus further expands the time regime covered and gives direct access to hydrogen bonds and base pairing.
The challenging steps in such a study are high-purity and high-yield sample preparation, potentially 13C- and 15N-labeled, as well as setup of experiments and fitting of data to extract population, exchange rate, and secondary structure of the previously invisible state. This protocol provides crucial hands-on steps in sample preparation to ensure the preparation of a suitable RNA sample and setup of 1H R1&#961; experiments with both isotopically labeled and unlabeled RNA samples.

We present a protocol to measure micro- to millisecond dynamics on 13C/15N-labeled and unlabeled RNA with 1H R1&#961; relaxation dispersion nuclear magnetic resonance (NMR) spectroscopy. The focus of this protocol lies in high-purity sample preparation and setup of NMR experiments.

RNA is a highly flexible biomolecule, wherein changes in structures play crucial roles in the functions that RNA molecules execute as cellular messengers ...

PAR-CliP - A Method to Identify Transcriptome-wide the Binding Sites of RNA Binding Proteins

RNA transcripts are subjected to post-transcriptional gene regulation by interacting with hundreds of RNA-binding proteins (RBPs) and microRNA-containing ribonucleoprotein complexes (miRNPs) that are often expressed in a cell-type dependently. To understand how the interplay of these RNA-binding factors affects the regulation of individual transcripts, high resolution maps of in vivo protein-RNA interactions are necessary1.
A combination of genetic, biochemical and computational approaches are typically applied to identify RNA-RBP or RNA-RNP interactions. Microarray profiling of RNAs associated with immunopurified RBPs (RIP-Chip)2 defines targets at a transcriptome level, but its application is limited to the characterization of kinetically stable interactions and only in rare cases3,4 allows to identify the RBP recognition element (RRE) within the long target RNA. More direct RBP target site information is obtained by combining in vivo UV crosslinking5,6 with immunoprecipitation7-9 followed by the isolation of crosslinked RNA segments and cDNA sequencing (CLIP)10. CLIP was used to identify targets of a number of RBPs11-17. However, CLIP is limited by the low efficiency of UV 254 nm RNA-protein crosslinking, and the location of the crosslink is not readily identifiable within the sequenced crosslinked fragments, making it difficult to separate UV-crosslinked target RNA segments from background non-crosslinked RNA fragments also present in the sample.
We developed a powerful cell-based crosslinking approach to determine at high resolution and transcriptome-wide the binding sites of cellular RBPs and miRNPs that we term PAR-CliP (Photoactivatable-Ribonucleoside-Enhanced Crosslinking and Immunoprecipitation) (see Fig. 1A for an outline of the method). The method relies on the incorporation of photoreactive ribonucleoside analogs, such as 4-thiouridine (4-SU) and 6-thioguanosine (6-SG) into nascent RNA transcripts by living cells. Irradiation of the cells by UV light of 365 nm induces efficient crosslinking of photoreactive nucleoside-labeled cellular RNAs to interacting RBPs. Immunoprecipitation of the RBP of interest is followed by isolation of the crosslinked and coimmunoprecipitated RNA. The isolated RNA is converted into a cDNA library and deep sequenced using Solexa technology. One characteristic feature of cDNA libraries prepared by PAR-CliP is that the precise position of crosslinking can be identified by mutations residing in the sequenced cDNA. When using 4-SU, crosslinked sequences thymidine to cytidine transition, whereas using 6-SG results in guanosine to adenosine mutations. The presence of the mutations in crosslinked sequences makes it possible to separate them from the background of sequences derived from abundant cellular RNAs.
Application of the method to a number of diverse RNA binding proteins was reported in Hafner et al.18

RNA transcripts are subject to extensive posttranscriptional regulation that is mediated by a multitude of trans-acting RNA-binding proteins (RBPs). Here we present a generalizable method to identify precisely and on a transcriptome-wide scale the RNA binding sites of RBPs.

RNA transcripts are subjected to post-transcriptional gene regulation by interacting with hundreds of RNA-binding proteins (RBPs) and microRNA-containing ...

Biology

Probing RNA Structure with Dimethyl Sulfate Mutational Profiling with Sequencing In Vitro and in Cells

The role of RNA structure in virtually any biological process has become increasingly evident, especially in the past decade. However, classical approaches to solving RNA structure, such as RNA crystallography or cryo-EM, have failed to keep up with the rapidly evolving field and the need for high-throughput solutions. Mutational profiling with sequencing using dimethyl sulfate (DMS) MaPseq is a sequencing-based approach to infer the RNA structure from a base&#39;s reactivity with DMS. DMS methylates the N1 nitrogen in adenosines and the N3 in cytosines at their Watson-Crick face when the base is unpaired. Reverse-transcribing the modified RNA with the thermostable group II intron reverse transcriptase (TGIRT-III) leads to the methylated bases being incorporated as mutations into the cDNA. When sequencing the resulting cDNA and mapping it back to a reference transcript, the relative mutation rates for each base are indicative of the base&#39;s &#34;status&#34; as paired or unpaired. Even though DMS reactivities have a high signal-to-noise ratio both in vitro and in cells, this method is sensitive to bias in the handling procedures. To reduce this bias, this paper provides a protocol for RNA treatment with DMS in cells and with in vitro transcribed RNA.

Probing RNA Structure with Dimethyl Sulfate Mutational Profiling with Sequencing In Vitro and in Cells

The protocol provides instruction for modifying RNA with dimethyl sulfate for mutational profiling experiments. It includes in vitro and in vivo probing with two alternative library preparation methods.

Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

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