Pool design and oligo recovery
<ol>
<li>The first step is to design the pre-mRNA pool to be studied. This can be done using the UCSC Genome Browser and downloading particular genes, splice junctions, or other areas of interest.</li>
<li>Once the windows of interest have been selected, tile across them computationally using the following conditions: read length should be 30 nucleotides with a 10 nucleotide overlap, therefore each oligo is shifted 20 nucleotides from the prior one. *Oligo overlap should be increased in proximity to splice sites to ensure adequate coverage; increasing the overlap from 10 to 20 nucleotides can accomplish this.</li>
<li>Flank each 30-nucleotide oligo with universal primer sequences, which will be used to amplify the pool downstream. Tiled oligo orders should be submitted to Agilent, where the sequences will be printed on a custom oligonucleotide microarray.</li>
<li>To recover the DNA oligos from the microarray surface begin by placing the array cover slide in an array hybridization chamber and gently pipetting 500 &mu;L of dH2O onto the slide</li>
<li>Sandwich the array on top of the cover slide, taking care to avoid air bubbles that can disrupt the process. Be sure to place the array face down so the side with the oligos is touching the surface of the water. A good rule of thumb is "Agilent touches Agilent" meaning that the Agilent label on the array faces the Agilent label on the cover slide</li>
<li>Close the hybridization chamber and rotate overnight at 99&deg;C in a hybridization oven</li>
<li>The following day, carefully remove the array from the chamber and draw off the water, which now contains the oligos liberated from the array surface. This is the oligo pool</li>
<li>Place the pool in a 1.5 mL Eppendorf tube and sonicate the pool at 50% amplitude for 3-5 second pulses</li>
<li>Next, amplify the pool by low cycle PCR, using the universal primers with a T7 tag appended to the end. For the first round, denature at 94&deg;C for 1 minute, anneal at 55&deg;C for 20 seconds, and elongate for 1 minute at 72&deg;C. For subsequent rounds, do 10 seconds, 20 seconds, and 10 seconds at each respective temperature, with a final elongation step of 5 minutes at 72&deg;C. Because the number of cycles necessary to amplify the pool depends on the efficiency of oligo recovery, it may be necessary to try multiple cycle numbers. Be careful not to over-amplify the pool, which is signified by a smeared band on an acrylamide gel. PCR amplified samples can be stored at 4&deg;C until needed.</li>
<li>The final step in oligo preparation is to transcribe RNA using the MEGAshortscript&trade; Kit from Ambion. *The following protocol has been adapted from Ambion
<ol class="lalpha">
 <li>Thaw the T7 10x Reaction Buffer, four ribonucleotide solutions and water at room temperature while keeping the T7 Enzyme Mix on ice</li>
 <li>Assemble the reaction mixture in an RNase-free microcentrifuge tube at room temperature. Components of the reaction buffer can cause template DNA to precipitate if the reaction is mixed on ice.</li>
 <li>In the following order, pipette 3 &mu;L nuclease-free water, 2 &mu;L of the 10x reaction buffer, 2 &mu;L of each 75 mM nTP solution, 5 &mu;L of template DNA (the PCR amplified oligo pool) and 2 &mu;L of T7 enzyme, for a final volume of 20 &mu;L</li>
 <li>Flick the tube gently to mix the reaction then briefly microfuge to collect the mixture at the bottom of the tube.</li>
 <li>Incubate the reaction in a thermocycler at 37&deg;C for 2 hours. Incubation time will be template-dependent, therefore it may be necessary to use a time-course experiment to determine the optimal incubation time for maximal yield</li>
 <li>After 2 hours, remove the template DNA by adding 1 &mu;L of Turbo DNase to the reaction and incubate at 37&deg;C for an additional 15 minutes</li>
 <li>Terminate the reaction and recover the RNA by phenol/chloroform extraction and ethanol precipitation.
 <ol class="lalpha">
 <li>For a 20 &mu;L reaction volume, add 115 &mu;L of dH2O and 15 &mu;L of 3 M sodium acetate to the reaction. </li>
 <li>Mix thoroughly then add 75 &mu;L of acidic phenol and 75 &mu;L of chloroform. Mix the solution by vortexing, then microfuge for one minute at 13,000 rpm at room temperature. Transfer the aqueous layer to a new tube and repeat the extraction</li>
 <li>Transfer the aqueous layer to a new tube and add 150 &mu;L of chloroform. Vortex the mixture and centrifuge as before. Transfer the aqueous layer to a new tube. </li>
 </ol>
 </li>
 <li>Recover the RNA by adding two volumes of 100% cold ethanol to the aqueous layer and mixing well. Chill the mixture for a minimum of 15 minutes at -20&deg;C then centrifuge at 4&deg;C for 15 minutes at maximum speed to pellet the RNA. Carefully remove the supernatant and resuspend the pellet in 105 &mu;L of 0.1x Tris-EDTA (TE) Buffer..</li>
</ol>
</li>
<li>It is difficult to remove all of the free nucleotides from the T7 Kit reaction, so for the most accurate results, quantify the RNA using RiboGreen.
<ol class="lalpha">
 <li>To begin the quantitation, first dilute enough 1x TE, 2,300 &mu;L + 50&mu;L/sample.</li>
 <li>Next, make the RiboGreen. You will need 1000 &mu;L + 50 &mu;L/sample. For high range samples (1 &mu;g/mL to 20 ng/mL) mix the RiboGreen 1:200 in 1x TE; for the low range samples (50 ng/mL to 1 ng/mL) mix the RiboGreen 1:2000 in 1x TE; Pipette 50 &mu;L of RiboGreen solution into each well of an opaque plate, starting with A1 and moving down, not across</li>
 <li>Dilute an RNA stock solution to 2 &mu;g/mL for high range samples or 100 ng/&mu;L for low range samples</li>
 <li>Using the stock solutions, create a standard curve by six subsequent 1:2 dilutions of the original 2 &mu;g/mL or 100 ng/mL stock; Pipette 240 &mu;L of the stock into a PCR strip tube. In the remaining 7 tubes, pipette 120 &mu;L of 1x TE. Take 120 &mu;L from the RNA stock well, and mix with tube 2. Take 120 &mu;L from tube 2 and mix with tube 3. Repeat until tube 7. Tube 8 will only have TE and will be your blank. Pipette 50 &mu;L of the standard curve solutions into the opaque plate, A1:H2 (run the curve twice)</li>
 <li>In a new tube, mix 45 &mu;L of TE with 5 &mu;L of the oligo pool and add all 50 &mu;L to a well that contains 50 &mu;L of RiboGreen.</li>
 <li>Using a plate reader set the machine to read fluorescence from the top after a one-minute mix with an excitation wavelength of 485 nm and emission wavelength of 530 nm and read the plate</li>
 <li>Use the standard curve generated to quantify the recovered RNA, which should be stored at -20&deg;C.</li>
</ol>
</li>
</ol>
Co-immunoprecipitation of the oligo pool with an RNP of interest
<ol>
<li>Prior to beginning the co-immunoprecipitation, prepare the Protein A and Protein G Dynabeads from Invitrogen by mixing them in a 1:1 ratio up to a final volume of 50 &mu;L per reaction. For example, for two reactions you would add 50 &mu;L each of each bead. </li>
<li>Use a Magnetic Separation Stand like this one from Novagen to hold the beads in place while you remove the supernatant and wash the beads twice with an equal volume of cold 1x PBS</li>
<li>After the second wash, resuspend the beads in an equal volume of cold 1x PBS and add 2 ng of the antibody per reaction. This amount may vary depending on the antibody, so experimentation is necessary to find the optimal conditions.</li>
<li>Incubate the antibody/bead mixture overnight at 4&deg;C on a rotating platform</li>
<li>In the morning, add 2 &mu;g per reaction of sonicated yeast total RNA to block non-specific binding, and rotate for an additional 30 minutes at 4&deg;C.</li>
<li>Using the magnetic rack to hold the beads, remove the supernatant and wash the beads twice with cold 1x PBS, leaving the PBS from the second wash in the tube.</li>
<li>Take out new tubes, one for each reaction and aliquot 50 &mu;L of the bead mixture into each tube. Let the beads sit while preparing a master mix</li>
<li>For each reaction add 200 ng of the oligo pool, an equimolar ratio of protection primers, 2 &mu;g of sonicated yeast total RNA, 20 &mu;L of HeLa nuclear extract (this is the RNP source), and buffer E to bring the volume up to 120 &mu;L. The protection primers are RNA versions of the universal primers. Order transcription templates of the universal primers with a T7 tag appended to the end, then transcribe RNA from the templates using the MEGAshortscript&trade; Kit .</li>
<li>Remove the supernatant from the bead aliquots and resuspend beads in 120 &mu;L of the master mix. Rock the reactions at 4&deg;C for 1-2 hours.</li>
<li>After the incubation, for each reaction, remove a small aliquot (including the beads) for a "total" RNA sample</li>
<li>Place the remainder of the reactions on the magnetic separation stand and remove the supernatant. Although it is not necessary to analyze this flow-through sample downstream, it may be useful to save.</li>
<li>Wash the beads 1-2 times with 50 &mu;L of cold 1x PBS. The RNA bound to the antibody on the beads is the "IP" sample.</li>
<li>Resuspend the "total" and the "IP" beads in 50 &mu;L of 1% SDS in 0.1x TE buffer and heat the samples at 65&deg;C for 15 minutes to elute the RNA.</li>
<li>Remove the supernatant, which now contains the RNA that was previously bound to the beads and clean by phenol/chloroform extraction followed by an ethanol precipitation. Dry the pellet and resuspend it in 50 &mu;L 0.1x TE buffer.</li>
</ol>
Reverse transcription and amplification of co-IP samples in preparation for aminoallyl labeling
<ol>
<li>Next, the RNA samples must be reverse transcribed and PCR amplified. For the reverse transcription reaction, use the reverse universal primers.</li>
<li>For each sample, begin by mixing 1 &mu;L of the reverse universal primer, with 1 &mu;L of the template RNA (from the immunoprecipitation reaction) and 10 &mu;L of dH2O. Heat the reactions at 70&deg;C for 5 minutes then chill on ice or keep at 4&deg;C.</li>
<li>While the samples cool, make the following master mix. For each reaction combine 4 &mu;L of 10 mM dNTPs, 2 &mu;L of 10x Arrayscript buffer, 1 &mu;L RNase inhibitor, and 1 &mu;L reverse transcription enzyme. Mix thoroughly and add 8 &mu;L of the master mix to each reverse transcription reaction.</li>
<li>Use a PCR machine to incubate the samples at 42&deg;C for 2 hours, followed by 95&deg;C for 5 minutes, then 4&deg;C until the samples are needed for amplification.</li>
<li>After reverse transcribing the RNA samples, PCR amplify them using the universal primers, with one of the primers containing a T7 tag. The number of cycles needed to amplify the pool will depend on the efficiency of the co-IP so it may be necessary to try multiple cycles. Each reaction requires 5 &mu;L of 10x Taq reaction buffer, 0.2 &mu;L of each primer (100 mM), 1 &mu;L of the cDNA template (from the reverse transcription reaction), 1 &mu;L of 10 mM dNTPs, 42.2 &mu;L of dH2O, and 0.4 &mu;L of Taq enzyme. Begin the PCR by incubating the samples at 94&deg;C for 3 minutes. Next, for each cycle, denature at 94&deg;C for 45 seconds, anneal at 55&deg;C for 30 seconds, and elongate at 72&deg;C for one minute, with a final elongation at 72&deg;C for 10 minutes.</li>
<li>Next, transcribe the samples again using the MEGAshortscript&trade; Kit, however, this time add 2 &mu;L of aminoallyl UTP, instead of the T7 UTP. The aminoallyl UTP will be used to label the RNA with the Cy dyes.</li>
<li>Phenol/chloroform and ethanol precipitate the samples. Do NOT use Glycoblue or ammonium acetate, both of which may interfere with hybridization of the samples to the array. Dry the pellet and resuspend in 4.5 &mu;L of 0.1 M sodium carbonate.</li>
</ol>
Amino-allyl labeling of RNA with Cy Dyes
<ol>
<li>To begin the labeling protocol dissolve each of the fluorescent dyes, cy3 and cy5, in 45 &mu;L of DMSO. To avoid light, wrap the tubes in aluminum foil and store at -20&deg;C in the dark</li>
<li>Add 2.5 &mu;L of nuclease-free water to RNA samples and mix thoroughly</li>
<li>Next, add 3 &mu;L of Cy3 to the "total" sample, and 3 &mu;L of Cy5 to the "IP" sample. Incubate the samples in the dark for 1 hour at room temperature; The Cy-3 sample will turn red or pink while the Cy-5 will turn blue.</li>
<li>To terminate the reaction, add 6 uL of 4 M hydroxylamine-HCl to each sample, mix thoroughly and spin briefly. Incubate for 15 minutes at room temperature.</li>
<li>Purify the labeled samples by phenol/chloroform extraction followed by ethanol precipitation and resuspend pellet in 50 &mu;L of 0.1x TE</li>
</ol>
Hybridize samples to the microarray using the Agilent Array Hybridization Kit
<ol>
<li>Begin the hybridization by gently mixing 50 &mu;L of 10x blocking buffer, 140 &mu;L of nuclease-free H2O, 25 &mu;L of the Cy3-labeled "total" pool, 25 &mu;L of the Cy5-labeled "IP" pool, 10 &mu;L of the 25x fragmentation buffer and 250 &mu;L of the 2x hybridization buffer, taking care to avoid introducing bubbles. Microfuge the sample briefly to collect the solution in the bottom of the tube and also to reduce bubbles.</li>
<li>Place the o-ring cover slide in the hybridization chamber and carefully pipette the solution onto the slide.</li>
<li>Place the detection array on top of the cover slide to "sandwich" the solution. Remember to orient the array slide so that the Agilent label on the array faces the Agilent label on the cover slide.</li>
<li>Clamp the slide/array sandwich in the hybridization chamber and rotate at 50&deg;C for 3 hours.</li>
<li>While the array is rotating, make the following 50 mL solutions in 50-mL conical tubes: one conical filled with 50 mL of dH2O, two 1x SCC buffer solutions, each in a separate tube, and one 2x SCC buffer.</li>
<li>After the three-hour incubation, carefully unclamp the array/slide sandwich and place it in the 50 mL conical containing the 2x SCC buffer. Using a pair of tweezers, gently separate the array from the cover slide and pull out the cover slide. Be careful not to touch the array as the slide is pulled out. Close the conical and gently invert for 60 seconds.</li>
<li>Using the tweezers, transfer the array to the 1x SCC solution, cap the tube, and gently invert for 1 minute then transfer the array to the second 1x SCC solution and again gently invert for another minute.</li>
<li>Lastly, transfer the array to the tube of dH2O and gently invert for 30 seconds.</li>
<li>Dry the array by placing the edges of the slide on a Kimwipe. Continue to rotate the array on the wipe making sure to not disrupt the side containing the oligos.</li>
<li>Place the dry array in an empty 50-mL conical tube and scan with a Microarray scanner</li>
</ol>
Interpreting the Microarray
<ol>
<li>The microarray scanner will create a TIF image of the array. Run this image through the Agilent Feature Extraction software, using the grid file (supplied by Agilent) which is unique to your specific pool.</li>
<li>The Agilent software will interpret the signal at each feature to create a file, which is parsed with a Perl script. The signal at each feature is normalize to the total intensity at that channel. For example at a particular feature in the green channel, the intensity of the green channel for the individual feature is divided by the total intensity of the green channel over all features. The final output is the log ratio of the red channel over the green channel at each feature on the array.</li>
<li>The oligo pool data from the microarray can be re-mapped back to the genome to get a picture of where, and how strongly, the RNP of interest bound.</li>
</ol>
Figure 1. Experimental schematic
<img src="/files/ftp_upload/1622/1622fig1a.jpg" alt="Figure 1a" /> A. Tiling scheme. After selecting and downloading pre-mRNA sequences of interest, the selecting area is tiled through in 30 nucleotide windows that shift in 10 nucleotide increments. Universal primer binding sites flank each side of the window.
<img src="/files/ftp_upload/1622/1622fig1b.jpg" alt="Figure 1b" /> B. Synthesis Array. Tiled oligo orders should be submitted to Agilent, where the sequences will be printed on a custom oligonucleotide microarray. 
<img src="/files/ftp_upload/1622/1622fig1c.jpg" alt="Figure 1c" /> C. Co-immunoprecipitation. After boiling the oligos off the array, incubate in HeLa nuclear extract, then add the extract to magnetic beads that are prepared with an antibody against the RNP of interest. Label the starting oligo pool with Cy3 and the bound IP pool with Cy5 and apply to the detection array and scan.
Figure 2. Annotating the genome with and analysis of enrichment data.
<img src="/files/ftp_upload/1622/1622fig2.jpg" alt="Figure 2" /> A. Convert the average score at each genomic coordinate from the array data to base-ten log and map that score to the given coordinates. An illustration of this averaging step is given for 3 overlapping 30-nt oligos with scores of 2, 4, and 0.5, where the average enrichment score for each 10-nt window is graphed above. The selected genomic regions can be visualized using a Custom Track in the USCS Genome Browser. The example browser window given is in the clta gene, where gene features (exons/introns/alternative splicing etc.) are given along the bottom and log average enrichment scores from the PTB-bound oligos are represented by dark red vertical bars. PTB strongly binds just upstream of exon 2 in the polypyrimidine tract.

When doing this procedure it&#8217;s important to remember that creation of the pool is flexible and open to modification at either the RNA or protein level. At the RNA level orthologous regions, disease mutations, polymorphisms, or random mutations can be introduced into the oligonucleotide sequence. At the protein level the RNA binding factor can be modified by phosphorylation or other post translational modifications. Additionally, the binding environment can be manipulated to either increase or decrease the levels ...

a rapid high-throughput method for mapping ribonucleoproteins (rnps) on human pre-mrna

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding location often influences function of a splicing factor.  However, as with any sampling strategy the chance of identifying an RNA bound to a splicing factor is proportional to its cellular abundance.  We have developed a novel in vitro approach for surveying binding specificity on otherwise transient pre-mRNA. This approach utilizes a specifically designed oligonucleotide pool that tiles across introns, exons, splice junctions, or other pre-mRNA.  The pool is subjected to some kind of molecular selection. Here, we demonstrate the method by separating the oligonucleotide into a bound and unbound fraction and utilize a two color array strategy to record the enrichment of each oligonucleotide in the bound fraction. The array data generates high-resolution maps with the ability to identify sequence-specific and structural determinates of ribonucleoprotein (RNP) binding on pre-mRNA.  A unique advantage to this method is its ability to avoid the sampling bias towards mRNA associated with current IP and SELEX techniques, as the pool is specifically designed and synthesized from pre-mRNA sequence. The flexibility of the oligonucleotide pool is another advantage since the experimenter chooses which regions to study and tile across, tailoring the pool to their individual needs.  Using this technique, one can assay the effects of polymorphisms or mutations on binding on a large scale or clone the library into a functional splicing reporter and identify oligonucleotides that are enriched in the included fraction.  This novel in vitro high-resolution mapping scheme provides a unique way to study RNP interactions with transient pre-mRNA species, whose low abundance makes them difficult to study with current in vivo techniques.  



Watch this Scientific Journal Video about A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA at JoVE.com

A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding ...

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Biology

11.7K Views.  Brown University. Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

Video: A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

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

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.

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 ...

Biochemistry

An Oligonucleotide-based Tandem RNA Isolation Procedure to Recover Eukaryotic mRNA-Protein Complexes

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein complexes is essential to understanding mRNA regulation inferred by interacting proteins or non-coding RNAs. Herein, we describe a tandem RNA isolation procedure (TRIP) that enables the purification of endogenously formed mRNA-protein complexes from cellular extracts. The two-step protocol involves the isolation of polyadenylated mRNAs with antisense oligo(dT) beads and subsequent capture of an mRNA of interest with 3&#39;-biotinylated 2&#39;-O-methylated antisense RNA oligonucleotides, which can then be isolated with streptavidin beads. TRIP was used to recover in vivo crosslinked mRNA-ribonucleoprotein (mRNP) complexes from yeast, nematodes and human cells for further RNA and protein analysis. Thus, TRIP is a versatile approach that can be adapted to all types of polyadenylated RNAs across organisms to study the dynamic re-arrangement of mRNPs imposed by intracellular or environmental cues.

A tandem RNA isolation procedure (TRIP) for recovery of endogenously formed mRNA-protein complexes is described. Specifically, RNA-protein complexes are crosslinked in vivo, polyadenylated RNAs are isolated from extracts with oligo(dT) beads, and particular mRNAs are captured with modified RNA antisense oligonucleotides. Proteins bound to mRNAs are detected by immunoblot analysis.

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein ...

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture (CARIC) Strategy

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used strategy for RBP capture exploits the polyadenylation [poly(A)] of target RNAs, which mostly occurs on eukaryotic mature mRNAs, leaving most binding proteins of non-poly(A) RNAs unidentified. Here we describe the detailed procedures of a recently reported method termed click chemistry-assisted RNA-interactome capture (CARIC), which enables the transcriptome-wide capture of both poly(A) and non-poly(A) RBPs by combining the metabolic labeling of RNAs, in vivo UV cross-linking, and bioorthogonal tagging.

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture CARIC Strategy

A detailed protocol for applying the click chemistry-assisted RNA-interactome capture (CARIC) strategy to identify proteins binding to both coding and noncoding RNAs is presented.

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used ...

RiboTag Immunoprecipitation in the Germ Cells of the Male Mouse

Quantifying differences in mRNA abundance is a classic approach to understand the impact of a given gene mutation on cell physiology. However, characterizing differences in the translatome (the whole of translated mRNAs) provides an additional layer of information particularly useful when trying to understand the function of RNA regulating or binding proteins. A number of methods for accomplishing this have been developed, including ribosome profiling and polysome analysis. However, both methods carry significant technical challenges and cannot be applied to specific cell populations within a tissue unless combined with additional sorting methods. In contrast, the RiboTag method is a genetic-based, efficient, and technically straightforward alternative that allows the identification of ribosome associated RNAs from specific cell populations without added sorting steps, provided an applicable cell-specific Cre driver is available. This method consists of breeding to generate the genetic models, sample collection, immunoprecipitation, and downstream RNA analyses. Here, we outline this process in adult male mouse germ cells mutant for an RNA binding protein required for male fertility. Special attention is paid to considerations for breeding with a focus on efficient colony management and the generation of correct genetic backgrounds and immunoprecipitation in order to reduce background and optimize output. Discussion of troubleshooting options, additional confirmatory experiments, and potential downstream applications is also included. The presented genetic tools and molecular protocols represent a powerful way to describe the ribosome-associated RNAs of specific cell populations in complex tissues or in systems with aberrant mRNA storage and translation with the goal of informing on the molecular drivers of mutant phenotypes.

Here, we describe the immunoprecipitation of ribosomes and associated RNA from select populations of adult male mouse germ cells using the RiboTag system. Strategic breeding and careful immunoprecipitation result in clean, reproducible results that inform on the germ cell translatome and provide insight into the mechanisms of mutant phenotypes.

Quantifying differences in mRNA abundance is a classic approach to understand the impact of a given gene mutation on cell physiology. However, ...

Developmental Biology

iCLIP - Transcriptome-wide Mapping of Protein-RNA Interactions with Individual Nucleotide Resolution

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional regulation1. Therefore, an essential step towards understanding transcript regulation at the molecular level is to gain positional information on the binding sites of RBPs2.
Protein-RNA interactions can be studied using biochemical methods, but these approaches do not address RNA binding in its native cellular context. Initial attempts to study protein-RNA complexes in their cellular environment employed affinity purification or immunoprecipitation combined with differential display or microarray analysis (RIP-CHIP)3-5. These approaches were prone to identifying indirect or non-physiological interactions6. In order to increase the specificity and positional resolution, a strategy referred to as CLIP (UV cross-linking and immunoprecipitation) was introduced7,8. CLIP combines UV cross-linking of proteins and RNA molecules with rigorous purification schemes including denaturing polyacrylamide gel electrophoresis. In combination with high-throughput sequencing technologies, CLIP has proven as a powerful tool to study protein-RNA interactions on a genome-wide scale (referred to as HITS-CLIP or CLIP-seq)9,10. Recently, PAR-CLIP was introduced that uses photoreactive ribonucleoside analogs for cross-linking11,12.
Despite the high specificity of the obtained data, CLIP experiments often generate cDNA libraries of limited sequence complexity. This is partly due to the restricted amount of co-purified RNA and the two inefficient RNA ligation reactions required for library preparation. In addition, primer extension assays indicated that many cDNAs truncate prematurely at the crosslinked nucleotide13. Such truncated cDNAs are lost during the standard CLIP library preparation protocol. We recently developed iCLIP (individual-nucleotide resolution CLIP), which captures the truncated cDNAs by replacing one of the inefficient intermolecular RNA ligation steps with a more efficient intramolecular cDNA circularization (Figure 1)14. Importantly, sequencing the truncated cDNAs provides insights into the position of the cross-link site at nucleotide resolution. We successfully applied iCLIP to study hnRNP C particle organization on a genome-wide scale and assess its role in splicing regulation14.

The spatial arrangement of RNA-binding proteins on a transcript is a key determinant of post-transcriptional regulation. Therefore, we developed individual-nucleotide resolution UV crosslinking and immunoprecipitation (iCLIP) that allows precise genome-wide mapping of the binding sites of an RNA-binding protein.

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional ...

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Development of the lower urogenital tract (LUT) is an intricate process. This complexity is evidenced during formation of the prostate from the fetal male urethra, which relies on androgenic signals and epithelial-mesenchymal interactions1,2. Understanding the molecular mechanisms responsible for prostate development may reveal growth mechanisms that are inappropriately reawakened later in life to give rise to prostate diseases such as benign prostatic hyperplasia and prostate cancer.
The developing LUT is anatomically complex. By the time prostatic budding begins on 16.5 days post conception (dpc), numerous cell types are present. Vasculature, nerves and smooth muscle reside within the mesenchymal stroma3. This stroma surrounds a multilayered epithelium and gives rise to the fetal prostate through androgen receptor-dependent paracrine signals4. The identity of the stromal androgen receptor-responsive genes required for prostate development and the mechanism by which prostate ductal epithelium forms in response to these genes is not fully understood. The ability to precisely identify cell types and localize expression of specific factors within them is imperative to further understand prostate development. In situ hybridization (ISH) allows for localization of mRNAs within a tissue. Thus, this method can be used to identify pattern and timing of expression of signaling molecules and their receptors, thereby elucidating potential prostate developmental regulators.
Here, we describe a high throughput ISH technique to identify mRNA expression patterns in the fetal mouse LUT using vibrating microtome-cut sections. This method offers several advantages over other ISH protocols. Performing ISH on thin sections adhered to a slide is technically difficult; cryosections frequently have poor structural quality while both cryosections and paraffin sections often result in weak signal resolution. Performing ISH on whole mount tissues can result in probe trapping. In contrast, our high throughput technique utilizes thick-cut sections that reveal detailed tissue architecture. Modified microfuge tubes allow easy handling of sections during the ISH procedure. A maximum of 4 mRNA transcripts can be screened from a single 17.5dpc LUT with up to 24 mRNA transcripts detected in a single run, thereby reducing cost and maximizing efficiency. This method allows multiple treatment groups to be processed identically and as a single unit, thereby removing any bias for interpreting data. Most pertinently for prostate researchers, this method provides a spatial and temporal location of low and high abundance mRNA transcripts in the fetal mouse urethra that gives rise to the prostate ductal network.

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Here, we describe an efficient high throughput in situ hybridization (ISH) method for visualizing patterns of mRNA expression in developing fetal mouse prostate tissue sections. The method can be easily adapted to visualize mRNA expression patterns in other mouse tissues or in tissues from other species.

Development of the lower urogenital tract (LUT) is an intricate process. This complexity is evidenced during formation of the prostate from the fetal male ...

Method for the Isolation and Identification of mRNAs, microRNAs and Protein Components of Ribonucleoprotein Complexes from Cell Extracts using RIP-Chip

As a result of the development of high-throughput sequencing and efficient microarray analysis, global gene expression analysis has become an easy and readily available form of data collection. In many research and disease models however, steady state levels of target gene mRNA does not always directly correlate with steady state protein levels. Post-transcriptional gene regulation is a likely explanation of the divergence between the two. Driven by the binding of RNA Binding Proteins (RBP), post-transcriptional regulation affects mRNA localization, stability and translation by forming a Ribonucleoprotein (RNP) complex with target mRNAs. Identifying these unknown de novo mRNA targets from cellular extracts in the RNP complex is pivotal to understanding mechanisms and functions of the RBP and their resulting effect on protein output. This protocol outlines a method termed RNP immunoprecipitation-microarray (RIP-Chip), which allows for the identification of specific mRNAs associated in the ribonucleoprotein complex, under changing experimental conditions, along with options to further optimize an experiment for the individual researcher. With this important experimental tool, researchers can explore the intricate mechanisms associated with post-transcriptional gene regulation as well as other ribonucleoprotein interactions.

A step by step protocol to isolating and identifying RNA associated complexes through RIP-Chip.

As a result of the development of high-throughput sequencing and efficient microarray analysis, global gene expression analysis has become an easy and ...

Simultaneous Mapping and Quantitation of Ribonucleotides in Human Mitochondrial DNA

Established approaches to estimate the number of ribonucleotides present in a genome are limited to the quantitation of incorporated ribonucleotides using short synthetic DNA fragments or plasmids as templates and then extrapolating the results to the whole genome. Alternatively, the number of ribonucleotides present in a genome may be estimated using alkaline gels or Southern blots. More recent in vivo approaches employ Next-generation sequencing allowing genome-wide mapping of ribonucleotides, providing the position and identity of embedded ribonucleotides. However, they do not allow quantitation of the number of ribonucleotides which are incorporated into a genome. Here we describe how to simultaneously map and quantitate the number of ribonucleotides which are incorporated into human mitochondrial DNA in vivo by Next-generation sequencing. We use highly intact DNA and introduce sequence specific double strand breaks by digesting it with an endonuclease, subsequently hydrolyzing incorporated ribonucleotides with alkali. The generated ends are ligated with adapters and these ends are sequenced on a Next-generation sequencing machine. The absolute number of ribonucleotides can be calculated as the number of reads outside the recognition site per average number of reads at the recognition site for the sequence specific endonuclease. This protocol may also be utilized to map and quantitate free nicks in DNA and allows adaption to map other DNA lesions that can be processed to 5&#180;-OH ends or 5&#180;-phosphate ends. Furthermore, this method can be applied to any organism, given that a suitable reference genome is available. This protocol therefore provides an important tool to study DNA replication, 5&#180;-end processing, DNA damage, and DNA repair.

Here we describe a method amenable to simultaneously quantitate and genome-wide map ribonucleotides in highly intact DNA at single-nucleotide resolution, combining enzymatic cleavage of genomic DNA with its alkaline hydrolysis and subsequent 5&#180;-end sequencing.

Established approaches to estimate the number of ribonucleotides present in a genome are limited to the quantitation of incorporated ribonucleotides using ...

Global Identification of Co-Translational Interaction Networks by Selective Ribosome Profiling

In recent years, it has become evident that ribosomes not only decode our mRNA but also guide the emergence of the polypeptide chain into the crowded cellular environment. Ribosomes provide the platform for spatially and kinetically controlled binding of membrane-targeting factors, modifying enzymes, and folding chaperones. Even the assembly into high-order oligomeric complexes, as well as protein-protein network formation steps, were recently discovered to be coordinated with synthesis.
Here, we describe Selective Ribosome Profiling, a method developed to capture co-translational interactions in vivo. We will detail the various affinity purification steps required for capturing ribosome-nascent-chain complexes together with co-translational interactors, as well as the mRNA extraction, size exclusion, reverse transcription, deep-sequencing, and big-data analysis steps, required to decipher co-translational interactions in near-codon resolution.

Co-translational interactions play a crucial role in nascent-chain modifications, targeting, folding, and assembly pathways. Here, we describe Selective Ribosome Profiling, a method for in vivo, direct analysis of these interactions in the model eukaryote Saccharomyces cerevisiae.

In recent years, it has become evident that ribosomes not only decode our mRNA but also guide the emergence of the polypeptide chain into the crowded ...

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Summary

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Chapters in this video

Novel RNA-Binding Proteins Isolation by the RaPID Methodology

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

An Oligonucleotide-based Tandem RNA Isolation Procedure to Recover Eukaryotic mRNA-Protein Complexes

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture (CARIC) Strategy

RiboTag Immunoprecipitation in the Germ Cells of the Male Mouse

iCLIP - Transcriptome-wide Mapping of Protein-RNA Interactions with Individual Nucleotide Resolution

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Method for the Isolation and Identification of mRNAs, microRNAs and Protein Components of Ribonucleoprotein Complexes from Cell Extracts using RIP-Chip

Simultaneous Mapping and Quantitation of Ribonucleotides in Human Mitochondrial DNA

Global Identification of Co-Translational Interaction Networks by Selective Ribosome Profiling