Name: exploring sequence space to identify binding sites for regulatory rna-binding proteins
Uploaded: 2019-08-09T14:00:00.000+00:00
Duration: 11 min 34 s
Description: Watch this Scientific Journal Video about Exploring Sequence Space to Identify Binding Sites for Regulatory RNA-Binding Proteins at JoVE.com

The author thanks the National Institutes of Health for the past funding.

NOTE: Figure 1 provides a summary of key steps in the iterative selection-amplification (SELEX) process.
1. Generation of a random library template
<ol>
	<li>Synthesize the forward primer 5&#8217;- GTAATACGACTCACTATAGGGTGATCAGATTCTGATCCA-3&#8217; and the reverse primer 5&#8217;- GCGACGGATCCAAGCTTCA-3&#8217; by chemical synthesis on a DNA synthesizer. 
	NOTE: The primers and the random library can be synthesized commercially.</li>
	<li>Synthesize a random library oligonucleotide template 5&#8217;- GGTGATCAGATTCTGATCCA(N1&#8230;N31)TGAAGCTTGGATCCGTCGC-3&#8217; by chemical synthesis. Use an equimolar mixture of four phosphoramidites during synthesis for the 31 randomized positions shown above as N. 
	NOTE: The sequence of the library template contains 31 random nucleotides (N1 to N31) and flanking sequences for the binding of the forward and reverse primers. The forward primer includes the T7 RNA polymerase promoter sequence (underlined) for in vitro transcription and a restriction site Bcl1 (italicized) for cloning. The reverse primer contains restriction enzyme sites BamH1 and HindIII (italicized) to facilitate cloning.</li>
</ol>
2. Generation of the DNA random library pool
<ol>
	<li>Attach the T7 RNA polymerase promoter to the library by polymerase chain reaction (PCR) containing 1 &#181;M DNA random library template, 1 &#181;M of each primer, 20 mM Tris (pH 8.0), 1.5 mM MgCl2, 50 mM KCl, 0.1 &#181;g/&#181;L acetylated bovine serum albumin, 2 units of Taq polymerase, and 200 &#181;M each of dNTPs (deoxyguanosine, deoxyadenosine, deoxycytidine, and deoxythymidine triphosphate).</li>
	<li>Use five cycles of denaturation, annealing, and extension steps (94 &#176;C for 1 min, 53 &#176;C for 1 min, and 72 &#176;C for 1 min) of PCR followed by one cycle of extension (72 &#176;C for 10 min).</li>
</ol>
3. Synthesis of pool 0 RNA
<ol>
	<li>Set up a 100 &#181;L transcription reaction12. Mix T7 transcription buffer, 1 &#181;M random library pool DNA, 5 mM dithiothreitol (DTT), 2 mM guanosine triphosphate (GTP), 1 mM each of adenosine triphosphate (ATP), cytidine triphosphate (CTP), and uridine triphosphate (UTP), and 2 units/&#181;L T7 RNA polymerase. 
	NOTE: RNA can be transcribed in vitro using commercially available kits with an option of the T7 or SP6 RNA polymerase.</li>
	<li>Incubate the above reaction mixture in a microcentrifuge tube for 2 h at 37 &#176;C.</li>
	<li>Gel purify RNA in a 10% denaturing polyacrylamide gel.</li>
	<li>Identify location of the transcripts on the gel by staining it with methylene blue or autoradiography by including traces of radioactivity (0.5 &#181;L or less of &#945;-32P UTP) in the transcription reaction.
	<ol>
		<li>Place the gel slice in a centrifuge tube and break into smaller pieces, for example, with a homogenizer tip. Add proteinase K (PK) buffer (100 mM Tris, pH 7.5, 150 mM NaCl, 12.5 mM EDTA, 1% Sodium dodecyl sulfate) to immerse the gel pieces. Leave the tube on a nutator from 2 h to overnight at room temperature.</li>
	</ol>
	</li>
	<li>Spin in a high-speed microcentrifuge (14,000 rpm or 16,873 x g) for 5 min at room temperature to remove the gel debris and recover the buffer solution.</li>
	<li>Vortex the sample two times with an equal volume of phenol-chloroform and one time with chloroform.</li>
	<li>Mix the aqueous phase from above with one-tenth volume of sodium acetate (3.0 M, pH 5.2), 10 &#181;g of tRNA or 20 &#181;g of glycogen, and ethanol (2&#8211;3 volumes, stored at -20 &#176;C). Leave the tubes at -80 &#176;C for 1 h.</li>
	<li>Spin the tubes containing the solution for 5&#8211;10 min at 4 &#176;C in a microcentrifuge (14,000 rpm or 16,873 x g). Discard the supernatant carefully. Rinse the RNA pellet with 70% ethanol and spin for 2&#8211;5 min. Aspirate ethanol carefully. Air dry the RNA pellet.</li>
	<li>Solubilize the RNA pellet in 50 &#181;L of water treated with of diethyl pyrocarbonate (DEPC). Leave the sample at -20 &#176;C for storage. 
	NOTE: Purify the RNA using commercially available spin columns, which are currently more commonly used to remove unincorporated radioactivity and serve as a quick and more convenient alternative for RNA purification. 
	CAUTION: Use an acrylic glass shield, gloves, and other precautions to protect from radioactivity.</li>
</ol>
4. Protein binding reaction and separation of bound RNA
<ol>
	<li>Carry out binding of protein and RNA in 10 mM Tris-HCl, pH 7.5 in a volume of 100 &#181;L by adding the following ingredients to these final concentrations: 50 mM KCl, 1 mM DTT, 0.09 &#181;g/&#181;L bovine serum albumin, 0.5 units/&#181;L RNasin, 0.15 &#181;g/&#181;L tRNA, 1 mM EDTA, and 30 &#181;L of appropriate recombinant protein (PTB) concentration. Add RNA from the appropriate pool. 
	NOTE: The splicing factor U2AF65 typically binds to the polypyrimidine-tract/3&#8217; splice sites of model introns with a binding affinity (equilibrium dissociation constant or Kd) of approximately 1&#8211;10 nM. Therefore, the first two rounds of binding used protein concentration 10-fold above the Kd for U2AF65; for SXL and PTB proteins, the starting concentration in this range was only our best guess. This ensured that desired RNA species that could bind, although lower affinity sequences also potentially bound. In rounds 3 and 4 (transcription, binding, and amplification), the protein concentration was reduced three-fold (Step 4.1). This was done to successively eliminate low affinity RNA species.</li>
	<li>Place the tubes containing the binding reactions for about 30 min at 25 &#176;C in a temperature block (or on ice).</li>
	<li>Fractionate the bound RNA from the unbound RNA for the first 4 rounds of selection-amplification using the following steps.
	<ol>
		<li>Filter the sample (100 &#181;L) at room temperature through a nitrocellulose filter attached to a vacuum manifold. 
		NOTE: The RNA-protein complex, but not the unbound RNA, remains on the filter.</li>
		<li>Chop the filter with retained RNA into fragments with a sterile razor blade; insert these into a centrifuge tube. Recover RNA by tumbling the tube gently for a minimum of 3 h (or overnight) with filter pieces immersed in the proteinase K (PK) buffer.</li>
		<li>Deproteinize the RNA sample by vortexing it in the presence of an equal volume of phenol-chloroform (1:1) and then of chloroform. Recover the aqueous phase each time by centrifuging the sample at high speed for 5 min at room temperature.</li>
		<li>Mix it with sodium acetate (0.1 volume of 3.0 M, pH 5.2) and ethanol (2&#8211;3 volumes of absolute ethanol, 200 proof). Leave the tube in a -80 &#176;C freezer for 30 min, centrifuge it at high speed for 10 min, and, following the washing and drying steps (step 3.8), solubilize the RNA in water treated with DEPC. These steps are outlined above (steps 3.6 to 3.9).</li>
	</ol>
	</li>
	<li>Separate the protein-bound RNA fractions from the unbound fractions for the last 2 rounds (rounds 5 and 6; transcription, binding, and amplification) as follows. Reduce the protein concentration in the binding reaction (step 4.1) by further three-fold for additional selection pressure to enrich high-affinity binding sequences and preferentially remove low-affinity sequences.
	<ol>
		<li>Pre-cast a native polyacrylamide gel (5% with 60:1 acrylamide:bis-acrylamide ratio) in 0.5x TBE buffer (Tris-Borate-EDTA) prior to setting up the above RNA:protein-binding (step 4.1) reaction. Electrophorese this gel in a cold room (4 &#176;C) by applying 250 V for 15 min.</li>
		<li>Pipette the above RNA:protein binding reactions (step 4.1) into different wells of this gel. 
		NOTE: The protein is stored at -80 &#176;C and diluted prior to use in 20 mM 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), pH 8.0, 20% glycerol, 0.2 mM ethylenediaminetetraacetic acid (EDTA), 0.05% NP-40, and 1 mM dithiothreitol (DTT). Addition of 0.5&#8211;1.0 mM protease inhibitor phenylmethane sulfonyl fluoride (PMSF) is optional. In the binding reaction, this buffer contributes about 6% glycerol, which allows direct loading of samples into the wells without the need for mixing them with a separate gel-loading buffer.</li>
		<li>Fractionate the bound RNA from the unbound RNA using gel electrophoresis in a cold room (4 &#176;C) at 250 V for 1 to 2 h. This process is also known as gel mobility shift assay. 
		NOTE: Duration of electrophoresis varies depending on features of the RNA and protein used for binding.</li>
		<li>Expose the gel to an X-ray film and identify the location of the bound RNA using autoradiography. Cut out the gel slice with the bound RNA and insert it into a tube.</li>
		<li>Incubate the crushed gel slice in the PK buffer used for elution for 3 h or overnight.</li>
		<li>Repeat steps 3.5 to 3.9 outlined above. Briefly, vortex the eluted RNA sample vigorously first with phenol-chloroform and then with chloroform.</li>
		<li>Mix sodium acetate and ethanol with the aqueous phase after chloroform extraction. Following incubation in -80 &#176;C freezer, spin at 4 &#176;C for 5&#8211;10 min to collect the RNA pellet. Wash the RNA pellet with ethanol and air dry it by leaving the lid of the tube open. Dissolve the RNA in water treated with DEPC. 
		NOTE: Switching to the gel mobility shift assay for fractionation allows elimination of unwanted RNA species that might have been enriched for binding, for example, to the nitrocellulose filter here (or any matrix) used in the initial rounds for fractionation.</li>
	</ol>
	</li>
</ol>
5. Reverse transcription and PCR amplification
<ol>
	<li>Synthesize cDNA from the dissolved RNA using reverse transcriptase and the reverse primer by incubating the 20 &#181;L reaction (2 &#181;L of 10x RT Buffer, 2 &#181;L of AMV reverse transcriptase, 1 &#181;M reverse primer, 10 &#181;L of RNA, RNase inhibitor optional) at 42 &#176;C for 60 min.</li>
	<li>Amplify the cDNA using 20&#8211;25 PCR cycles as described in step 2.1.</li>
</ol>
6. Transcription and protein binding
<ol>
	<li>Repeat the process of RNA synthesis, protein binding, separation of protein bound and unbound fraction, as described in section 3&#8211;5 above.</li>
</ol>
7. Analysis of RNA-protein interactions
<ol>
	<li>Use the gel mobility shift assay (step 4.4) or the filter binding assay (step 4.3) to determine binding affinity and specificity for selected pools or individual sequences within each pool (see step 8.3).</li>
	<li>Use autoradiography or a phosphor imager to detect and quantify bands in the bound fraction and unbound fraction.</li>
</ol>
8. Cloning and sequencing
<ol>
	<li>Digest the final PCR DNA product with restriction enzymes Bcl1 and HindIII for 1&#8211;2 h, ligate with the appropriately digested pGEM3 or other plasmids carrying the restriction sites from 2 h to overnight, transform the ligation product into competent bacterial cells by heat shock or electroporation using standard molecular biology procedures13.</li>
	<li>Grow bacteria overnight by plating the transformed cells on agar plates with Luria-Bertani (LB) medium and ampicillin (50 &#181;g/mL) at 37 &#176;C. Pick colonies to inoculate culture tubes containing LB liquid medium with ampicillin and grow at 37 &#176;C in a shaking incubator overnight. Purify plasmid DNAs containing DNA inserts using a standard plasmid isolation protocol13. 
	NOTE: Commercial kits are available for plasmid purification.</li>
	<li>Sequence the plasmids with DNA inserts using the dideoxy chain termination sequencing protocol, following manufacturer&#8217;s instructions for sequencing14. 
	NOTE: Sequencing can be performed in house or done commercially.</li>
</ol>
9. Sequence alignment
<ol>
	<li>Align sequences and obtain a consensus binding site(s) using available online alignment tools 
	(https://www.ebi.ac.uk/Tools/msa/).</li>
</ol>

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The author declares that he has no competing financial interests.

Nucleic acid-binding proteins are important regulators of animal and plant development. A key requirement for the SELEX procedure is the development of an assay that can be used to separate protein-bound and unbound RNA fractions. In principle, this assay can be an in vitro binding assay such as the filter-binding assay, the gel mobility shift assay, or a matrix binding assay19 for recombinant proteins, purified proteins, or protein complexes. The assay can also be an enzymatic assay where the precursors and products (or intermediates) can be separated based on size or some other means20.
While mutagenesis has been widely used to characterize binding sites for proteins, it is laborious, and time consuming and longer sequences are not as easily amenable to saturation mutagenesis. The significance of the iterative binding and amplification approach described here is that not only does it overcome some of the above limitations, it can most importantly identify previously unknown binding sites and provide important information about nucleotide requirements at each position at the same time.
An important consideration for the success of iterative selection-amplification is binding affinity and specificity. Typically, 12 to 15 rounds of selection-amplification are employed and a sequence space of 1012 to 1015 molecules can be routinely sampled. The progress and eventual success of the selection-amplification protocol can be monitored using a binding assay or direct sequencing, which monitors affinity for or enrichment of specific sequences in intermediate pools, respectively. While the binding assay was traditionally used, advent of the next generation sequencing allows analysis of sequence enrichment in ways not possible by manual Sanger sequencing14.
A critical step in the success of SELEX is fold enrichment of the desired molecules at each step. The number of cycles required for SELEX varies and depends on several factors. For example, if fold-enrichment of desired or specific sequences is higher in each round, fewer rounds will be sufficient. However, if an assay allows a high proportion of undesired sequences in the bound pool, additional rounds will become necessary to enrich desired RNA sequences. A limitation of the technique or an unintended consequence of the need for additional cycles of selection-amplification that must be kept in mind is the possibility that it might introduce artefacts or enrich sequences that have unrelated properties such as their ability to amplify. Finally, while some applications benefit from the highest affinity binders, for other uses, a balance must be struck during the selection-amplification process between binding affinity and function because tightest binding sequences might not necessarily be the most functional sequences in biological contexts (e.g., if a sequence is recognized multiple times by different proteins during splicing).
Among the modifications and troubleshooting to improve the procedure, negative selection or counter selection can be employed to increase specificity. Similarly, use of different partitioning protocols, such as the filter binding assay followed by the gel mobility shift assay, can eliminate enrichment of unwanted sequences that bind, for example, to the nitrocellulose filter or a column matrix21. Given that proteins-nucleic acid interactions have both specific and non-specific components, buffer conditions such as salt and pH have effects on RNA-protein interactions. Moreover, use of appropriate protein concentration can have a direct effect on retention of strong, weak and non-specific binders. Selection pressure can be increased in successive rounds, for example, by including a competitor RNA, reducing protein concentration, or reducing the time of incubation. Thus, careful considerations and optimizing these parameters can impact the outcome of the SELEX protocol.
Recently, many variations or modifications of the original SELEX protocol have been developed which overcome some of the limitations mentioned above. These include high throughput-SELEX (HT-SELEX), which combines SELEX and massively parallel sequencing6, RNAcompete, which involves incubation with excess non-random RNA, pull-down of the bound RNA, fluorescent labeling of RNA, and analysis on microarrays5, RNA Bind-n-Seq, which combines RNA affinity analysis in a quantitative and high throughput fashion22, and RAPID-SELEX, which shortens the process and includes a non-amplification step23.
Chemically modified bases have been used to expand the repertoire of the RNA molecules for specific applications24. Diagnostics, therapeutics, as well as molecules with catalytic activities are among the many applications (including in medicine) of the selected molecules25. Aptamers complement antibody-based protocols and provide excellent tools whose potential, for example, in diagnostics, therapeutics, and other applications, remains to be fully exploited26,27,28. In the future, for example, clinical benefits are among the numerous desired applications, beyond what the first FDA-approved aptamer (Pegaptanib sodium) could deliver for the age-related macular degeneration. The scalable proteomic technology for protein measurements offers a step toward understanding health and diseases24.

In cell biology, gene regulation plays a central role. At one or multiple steps along the gene expression pathway, genes have the potential to be regulated. These steps include transcription (initiation, elongation, and termination) as well as splicing, polyadenylation or 3&#8217; end formation, RNA export, mRNA translation, and decay/localization of primary transcripts. At these steps, nucleic acid-binding proteins modulate gene regulation. Identification of binding sites for such proteins is an important aspect of studying gene control. Mutational analysis and phylogenetic sequence comparison have been used to discover regulatory sequences or protein-binding sites in nucleic acids, such as promoters, splice sites, polyadenylation elements, and translational signals1,2,3,4.
Pre-mRNA splicing is an integral step during gene expression and regulation. The majority of mammalian genes, including those in humans, have introns. A large fraction of these transcripts is alternatively spliced, producing multiple mRNA and protein isoforms from the same gene or primary transcript. These isoforms have cell-specific and developmental roles in cell biology. The 5&#8217; splice site, the branch-point, and the polypyrimidine-tract/3&#8217; splice site are critical splicing signals that are subject to regulation. In negative regulation, an otherwise strong splice site is repressed, whereas in positive regulation an otherwise weak splice site is activated. A combination of these events produces a plethora of functionally distinct isoforms. RNA-binding proteins play key roles in these alternative splicing events.
Numerous proteins are known whose binding site(s) or RNA targets remain to be identified5, 6. Linking regulatory proteins to their downstream biological targets or sequences is often a complex process. For such proteins, identification of their target RNA or binding site is an important step in defining their biological functions. Once a binding site is identified, it can be further characterized using standard molecular and biochemical analyses.
The approach described here has two advantages. First, it can identify a previously unknown binding site for a protein of interest. Second, an added advantage of this approach is that it simultaneously allows saturation mutagenesis, which would otherwise be labor intensive to obtain comparable information about sequence requirements within the binding site. Thus, it offers a quicker, easier, and less costly tool to identify protein binding sites in RNA. Originally, this approach (SELEX or Systematic Evolution of Ligands by EXponential enrichment) was used to characterize the binding site for the bacteriophage T4 DNA polymerase (gene 43 protein), which overlaps with the ribosome binding site in its own mRNA. The binding site contains an 8-base loop sequence, representing 65,536 randomized variants for analysis7. Second, the approach was also independently used to show that specific binding sites or aptamers for different dyes can be selected from a pool of approximately 1013 sequences8. In fact, this approach has been broadly used in many different contexts to identify aptamers (RNA or DNA sequences) for binding numerous ligands, such as proteins, small molecules, and cells, and for catalysis9. As an example, an aptamer can discriminate between two xanthine derivatives, caffeine and theophylline, which differ by the presence of one methyl group in caffeine10. We have extensively used this approach (SELEX or iterative selection-amplification) to study how RNA-binding proteins function in splicing or splicing regulation11, which will be the basis for the discussion below.
The random library: We used a random library of 31 nucleotides. The length consideration for the random library was loosely based on the idea that the general splicing factor U2AF65 binds to a sequence between the branch-point sequence and the 3&#8217; splice site. On average, the spacing between these splicing signals in metazoans is in the range of 20 to 40 nucleotides. Another protein Sex-lethal was known to bind to a poorly characterized regulatory sequence near the 3&#8217; splice site of its target pre-mRNA, transformer. Thus, we chose a random region of 31 nucleotides, flanked by primer binding sites with restriction enzyme sites to allow for PCR amplification and attachment of the T7 RNA polymerase promoter for in vitro transcription. The theoretical library size or complexity was 431 or approximately 1018. We used a small fraction of this library to prepare our random RNA pool (~1012-1015) for the experiments described below.

<table><tbody><tr><td>Gel Electrophoresis equipment</td><td>Standard</td><td>Standard</td><td></td></tr><tr><td>Glass Plates</td><td>Standard</td><td>Standard</td><td></td></tr><tr><td>Nitrocellulose</td><td>Millipore</td><td>HAWP</td><td></td></tr><tr><td>Nitrocellulose</td><td>Schleicher &amp;amp; Schuell</td><td>PROTRAN</td><td></td></tr><tr><td>polyacrylamide gel solutions</td><td>Standard</td><td>Standard</td><td></td></tr><tr><td>Proteinase K</td><td>NEB</td><td>P8107S</td><td></td></tr><tr><td>Recombinant PTB</td><td>Laboratory Preparation</td><td>Not applicable</td><td></td></tr><tr><td>Reverse Transcriptase</td><td>NEB</td><td>M0277S</td><td></td></tr><tr><td>Vacuum manifold</td><td>Fisher Scientific</td><td>XX1002500</td><td>Millipore 25mm Glass Microanalysis Vacuum Filter</td></tr><tr><td>Vacuum manifold</td><td>Millipore</td><td>XX2702552</td><td>1225 Sampling Vacuum Manifold</td></tr><tr><td>X-ray films</td><td>Standard</td><td>Standard</td><td></td></tr></tbody></table>

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.

The following observations demonstrate successful selection-amplification (SELEX). First, we analyzed pool 0 and the selected sequences for binding to the protein used for the iterative selection-amplification approach. Figure 2 shows that the mammalian polypyrimidine-tract binding protein (PTB) shows barely detectable binding to the pool 0 sequence but high affinity for the selected sequence pool. There was barely detectable binding to pool 0 when we used about 300-fold higher protein concentration for binding than used for the selected pool. Thus, there was at least a several hundred-fold difference in protein binding affinities between the random or starting pool and the selected pool. This observation experimentally confirms that the selection-amplification protocol described here is successful.
Second, we sequenced the selected pool and determined a consensus binding site. The consensus sequence obtained from alignment of the majority of selected sequences from the mammalian PTB-selected pool was: GCCUG(Y/G)UGCYYYYCYYYG(Y/G)CCC. This shows that we have selected unique pyrimidine-rich sequences that bind PTB11. When we performed iterative selection-amplification for the RNA-binding domain of the Drosophila PTB, we enriched CU-rich sequences interrupted by guanosines. Among the high affinity sequences that the Drosophila PTB selected was an 84% pyrimidine-rich sequence: GCUUUCCUCUGUCGCCCUUCUUCGUCCCCUG. In fact, this sequence is similar to the pyrimidine-rich sequence present in the alpha-tropomyosin intron which binds with high affinity to and is regulated by the mammalian PTB15. We have successfully used this approach repeatedly to study RNA-binding properties and functions splicing regulators and a splicing factor11, 15, 16. Table 1 shows successful examples of RNA-binding proteins for which SELEX was used to identify their preferred or consensus binding site(s).
Third, an in vitro splicing assay, which is based on alternative 3&#39; splice site choice, shows functional relevance of distinct but overlapping RNA-binding specificities of polypyrimidine-tract binding proteins. Whereas an upstream 3&#39; splice site is used by default, addition of the recombinant PTB leads to activation of the alternative or downstream 3&#39; splice site (Figure 3). In contrast, addition of recombinant hnRNP C17 leads to repression of both 3&#39; splice sites. Addition of the recombinant general splicing factor U2AF65 reverses the hnRNP C1-mediated 3&#39; splice site repression (Figure 3) as well as the PTB-mediated effect on downstream 3&#39; splice site activation (data not shown). A simple explanation for these effects is a direct competition between the binding of the general splice factor U2AF65 and PTB (also called hnRNP I), which preferentially binds to and represses certain 3&#39; splices sites, or between U2AF65 and hnRNP C, which binds to and represses both 3&#39; splice sites.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59635/59635fig1.jpg" /> 
Figure 1: Summary of key steps in iterative selection-amplification process (SELEX). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59635/59635fig1large.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/59635/59635fig2.jpg" /> 
Figure 2: Enrichment of PTB-binding RNAs. Increasing concentration (filled triangles) of recombinant PTB was used with either radiolabeled pool 0 RNA or the selected pool obtained following six rounds of selection and amplification. Positions of unbound RNA and the RNA:protein complex are indicated. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59635/59635fig2large.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/59635/59635fig3.jpg" /> 
Figure 3: Splice site switching assay validates distinct binding specificities of pyrimidine-binding proteins. (A-Top) Schematics of the splicing substrate. The splicing substrate contains a 5&#39; splice site and two alternative 3&#39; splice sites flanking the intron. Rectangles (open, with horizontal lines, and solid) are exons and the line is an intron. (A-Bottom) hnRNP C1 represses the upstream 3&#39; splice site (without activation of the downstream 3&#39; splice site), whereas PTB leads to activation of the downstream 3&#39; splice site. The splicing substrate was incubated in a HeLa cell nuclear extract.The splicing products (shown on the sides) were analyzed using a primer extension assay18 with splice-junction primers (arrows), which recognize splicing of the common 5&#39; splice site to either the upstream or the downstream 3&#39; splice site. (B) Recombinant U2AF65 (rU2AF65) reverses the repressive effect of hnRNP C1. Addition of the recombinant hnRNP C1, PTB, or rU2AF65 proteins to the splicing reaction is indicated by the + symbols. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59635/59635fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Protein</td>
			<td>Preferred sequence(s)</td>
		</tr>
		<tr>
			<td>U2AF65</td>
			<td>U-rich containing Cs</td>
		</tr>
		<tr>
			<td>SXL</td>
			<td>U-rich containing 2-4 Gs</td>
		</tr>
		<tr>
			<td>PTB</td>
			<td>UCUUC-rich with some Gs</td>
		</tr>
		<tr>
			<td>hnRNP C1</td>
			<td>U-rich (5-6 long)</td>
		</tr>
		<tr>
			<td>CstF64</td>
			<td>GU-rich</td>
		</tr>
		<tr>
			<td>hnRNP E1/E2 and K</td>
			<td>C-rich</td>
		</tr>
		<tr>
			<td>U2AF65/U2AF35 heterodimer</td>
			<td>UUUYYYYUNUAGGU</td>
		</tr>
	</tbody>
</table>
Table 1: Preferred binding sites for some RNA-binding proteins.

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

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 ...

exploring-sequence-space-to-identify-binding-sites-for-regulatory-rna

Nanyang Technological University

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Genetics

6.6K Views.  University of Colorado at Boulder. 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.

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

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 ...

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

A Novel Saturation Mutagenesis Approach: Single Step Characterization of Regulatory Protein Binding Sites in RNA Using Phosphorothioates

Gene regulation plays an important role in development. Numerous DNA- and RNA-binding proteins bind their target sequences with high specificity to control gene expression. These regulatory proteins control gene expression either at the level of DNA (transcription) or at the level of RNA (pre-mRNA splicing, polyadenylation, mRNA transport, decay, and translation). Identification of regulatory sequences helps understand not only how a gene is switched on or off, but also which downstream genes are regulated by a particular regulatory protein. Here, we describe a one-step approach that allows saturation mutagenesis of a protein binding site in RNA. It involves doping DNA template with non-wild-type nucleotides within the binding site, synthesis of separate RNAs with each phosphorothioate nucleotide, and isolation of the bound fraction following incubation with protein. Interference from non-wild-type nucleotides results in their preferential exclusion from the protein-bound fraction. This is monitored by gel electrophoresis following selective chemical cleavage with iodine of phosphodiester bonds containing phosphorothioates (phosphorothioate mutagenesis or PTM). This single-step saturation mutagenesis approach is applicable to the characterization of any protein binding site in RNA.

Proteins that bind specific RNA sequences play critical roles in gene expression. Detailed characterization of these binding sites is crucial for our understanding of gene regulation. Here, a single-step approach for saturation mutagenesis of protein-binding sites in RNA is described. This approach is relevant for all protein-binding sites in RNA.

Gene regulation plays an important role in development. Numerous DNA- and RNA-binding proteins bind their target sequences with high specificity to ...

An Assay for Quantifying Protein-RNA Binding in Bacteria

In the initiation step of protein translation, the ribosome binds to the initiation region of the mRNA. Translation initiation can be blocked by binding of an RNA binding protein (RBP) to the initiation region of the mRNA, which interferes with ribosome binding. In the presented method, we utilize this blocking phenomenon to quantify the binding affinity of RBPs to their cognate and non-cognate binding sites. To do this, we insert a test binding site in the initiation region of a reporter mRNA and induce the expression of the test RBP. In the case of RBP-RNA binding, we observed a sigmoidal repression of the reporter expression as a function of RBP concentration. In the case of no-affinity or very low affinity between binding site and RBP, no significant repression was observed. The method is carried out in live bacterial cells, and does not require expensive or sophisticated machinery. It is useful for quantifying and comparing between the binding affinities of different RBPs that are functional in bacteria to a set of designed binding sites. This method may be inappropriate for binding sites with high structural complexity. This is due to the possibility of repression of ribosomal initiation by complex mRNA structure in the absence of RBP, which would result in lower basal reporter gene expression, and thus less-observable reporter repression upon RBP binding.

In this method, we quantify the binding affinity of RNA binding proteins (RBPs) to cognate and non-cognate binding sites using a simple, live, reporter assay in bacterial cells. The assay is based on repression of a reporter gene.

In the initiation step of protein translation, the ribosome binds to the initiation region of the mRNA. Translation initiation can be blocked by binding ...

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 ...

Identification of Footprints of RNA:Protein Complexes via RNA Immunoprecipitation in Tandem Followed by Sequencing (RIPiT-Seq)

RNA immunoprecipitation in tandem (RIPiT) is a method for enriching RNA footprints of a pair of proteins within an RNA:protein (RNP) complex. RIPiT employs two purification steps. First, immunoprecipitation of a tagged RNP subunit is followed by mild RNase digestion and subsequent non-denaturing affinity elution. A second immunoprecipitation of another RNP subunit allows for enrichment of a defined complex. Following a denaturing elution of RNAs and proteins, the RNA footprints are converted into high-throughput DNA sequencing libraries. Unlike the more popular ultraviolet (UV) crosslinking followed by immunoprecipitation (CLIP) approach to enrich RBP binding sites, RIPiT is UV-crosslinking independent. Hence RIPiT can be applied to numerous proteins present in the RNA interactome and beyond that are essential to RNA regulation but do not directly contact the RNA or UV-crosslink poorly to RNA. The two purification steps in RIPiT provide an additional advantage of identifying binding sites where a protein of interest acts in partnership with another cofactor. The double purification strategy also serves to enhance signal by limiting background. Here, we provide a step-wise procedure to perform RIPiT and to generate high-throughput sequencing libraries from isolated RNA footprints. We also outline RIPiT&#39;s advantages and applications and discuss some of its limitations.

Identification of Footprints of RNA:Protein Complexes via RNA Immunoprecipitation in Tandem Followed by Sequencing RIPiT-Seq

Here, we present a protocol to enrich endogenous RNA binding sites or &#34;footprints&#34;&#160;of RNA:protein (RNP) complexes from mammalian cells. This approach involves two immunoprecipitations of RNP subunits and is therefore dubbed RNA immunoprecipitation in tandem (RIPiT).

RNA immunoprecipitation in tandem (RIPiT) is a method for enriching RNA footprints of a pair of proteins within an RNA:protein (RNP) complex. RIPiT ...

MS2-Affinity Purification Coupled with RNA Sequencing in Gram-Positive Bacteria

Although small regulatory RNAs (sRNAs) are widespread among the bacterial domain of life, the functions of many of them remain poorly characterized notably due to the difficulty of identifying their mRNA targets. Here, we described a modified protocol of the MS2-Affinity Purification coupled with RNA Sequencing (MAPS) technology, aiming to reveal all RNA partners of a specific sRNA in vivo. Broadly, the MS2 aptamer is fused to the 5&#8217; extremity of the sRNA of interest. This construct is then expressed in vivo, allowing the MS2-sRNA to interact with its cellular partners. After bacterial harvesting, cells are mechanically lysed. The crude extract is loaded into an amylose-based chromatography column previously coated with the MS2 protein fused to the maltose binding protein. This enables the specific capture of MS2-sRNA and interacting RNAs. After elution, co-purified RNAs are identified by high-throughput RNA sequencing and subsequent bioinformatic analysis. The following protocol has been implemented in the Gram-positive human pathogen Staphylococcus aureus and is, in principle, transposable to any Gram-positive bacteria. To sum up, MAPS technology constitutes an efficient method to deeply explore the regulatory network of a particular sRNA, offering a snapshot of its whole targetome. However, it is important to keep in mind that putative targets identified by MAPS still need to be validated by complementary experimental approaches.

MAPS technology has been developed to scrutinize the targetome of a specific regulatory RNA in vivo. The sRNA of interest is tagged with a MS2 aptamer enabling the co-purification of its RNA partners and their identification by RNA sequencing. This modified protocol is particularly suited for Gram-positive bacteria.&#160;

Although small regulatory RNAs (sRNAs) are widespread among the bacterial domain of life, the functions of many of them remain poorly characterized ...

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

An Optimized Quantitative Pull-Down Analysis of RNA-Binding Proteins Using Short Biotinylated RNA

Protein-RNA interactions regulate gene expression and cellular functions at transcriptional and post-transcriptional levels. For this reason, identifying the binding partners of an RNA of interest remains of high importance to unveil the mechanisms behind many cellular processes. However, RNA molecules might interact transiently and dynamically with some RNA-binding proteins (RBPs), especially with non-canonical ones. Hence, improved methods to isolate and identify such RBPs are greatly needed.
To identify the protein partners of a known RNA sequence efficiently and quantitatively, we developed a method based on the pull-down and characterization of all interacting proteins, starting from cellular total protein extract. We optimized the protein pull-down using biotinylated RNA pre-loaded on streptavidin-coated beads. As a proof of concept, we employed a short RNA sequence known to bind the neurodegeneration-associated protein TDP-43 and a negative control of a different nucleotide composition but the same length. After blocking the beads with yeast tRNA, we loaded the biotinylated RNA sequences on the streptavidin beads and incubated them with the total protein extract from HEK 293T cells. After incubation and several washing steps to remove nonspecific binders, we eluted the interacting proteins with a high-salt solution, compatible with the most commonly used protein quantification reagents and with sample preparation for mass spectrometry. We quantified the enrichment of TDP-43 in the pull-down performed with the known RNA binder compared to the negative control by mass spectrometry. We used the same technique to verify the selective interactions of other proteins computationally predicted to be unique binders of our RNA of interest or of the control. Finally, we validated the protocol by western blot via the detection of TDP-43 with an appropriate antibody. 
This protocol will allow the study of the protein partners of an RNA of interest in near-to-physiological conditions, helping uncover unique and unpredicted protein-RNA interactions.

Here, we present an optimized in vitro method to uncover, quantify, and validate protein interactors of specific RNA sequences, using total protein extract from human cells, streptavidin beads coated with biotinylated RNA, and mass spectrometry analysis.

Protein-RNA interactions regulate gene expression and cellular functions at transcriptional and post-transcriptional levels. For this reason, identifying ...

Detection of RNA-binding Proteins by In Vitro RNA Pull-down in Adipocyte Culture

RNA-binding proteins (RBPs) are emerging as a regulatory layer in the development and function of adipose. RBPs play a key role in the gene expression regulation at posttranscriptional levels by affecting the stability and translational efficiency of target mRNAs. RNA pull-down technique has been widely used to study RNA-protein interaction, which is necessary to elucidate the mechanism underlying RBPs&#39; as well as long non-coding RNAs&#39; (lncRNAs) function. However, the high lipid abundance in adipocytes poses a technical challenge in conducting this experiment. Here a detailed RNA pull-down protocol is optimized for primary adipocyte culture. An RNA fragment from androgen receptor&#39;s (AR) 3&#39; untranslated region (3&#39;UTR) containing an adenylate-uridylate-rich elementwas used as an example to demonstrate how to retrieve its RBP partner, HuR protein, from adipocyte lystate. The method described here can be applied to detect the interactions between RBPs and noncoding RNAs, as well as between RBPs and coding RNAs.

Detection of RNA-binding Proteins by In Vitro RNA Pull-down in Adipocyte Culture

An RNA pull-down protocol is optimized here for detection of interactions between RNA-binding proteins (RBPs) and noncoding as well as coding RNAs. An RNA fragment from androgen receptor (AR) was used as an example to demonstrate how to retrieve its RBP from lystate of primary brown adipocytes.

RNA-binding proteins (RBPs) are emerging as a regulatory layer in the development and function of adipose. RBPs play a key role in the gene expression ...

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

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Novel RNA-Binding Proteins Isolation by the RaPID Methodology

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

A Novel Saturation Mutagenesis Approach: Single Step Characterization of Regulatory Protein Binding Sites in RNA Using Phosphorothioates

An Assay for Quantifying Protein-RNA Binding in Bacteria

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

Identification of Footprints of RNA:Protein Complexes via RNA Immunoprecipitation in Tandem Followed by Sequencing (RIPiT-Seq)

MS2-Affinity Purification Coupled with RNA Sequencing in Gram-Positive Bacteria

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

An Optimized Quantitative Pull-Down Analysis of RNA-Binding Proteins Using Short Biotinylated RNA

Detection of RNA-binding Proteins by In Vitro RNA Pull-down in Adipocyte Culture