This work was supported by a grant from the Canadian Institutes for Health Research (MOP-86514).

Experimental procedures with mice were approved by the Animal Care Committee of McGill University (protocol 4966).
1. Preparation of Protein Extracts from Cultured Cells
<ol>
	<li>Wash cultured cells twice with 10&#160;ml of ice-cold phosphate buffered saline (PBS).</li>
	<li>Scrape adherent cells with either a rubber policeman or a plastic cell scraper in 1&#160;ml of ice-cold PBS, transfer suspension into a 1.5&#160;ml microcentrifuge tube.</li>
	<li>Spin in a microcentrifuge for 5&#160;min at 700&#160;x&#160;g, at 4&#160;&#176;C. Aspirate PBS.</li>
	<li>Add 100&#160;&#956;l of ice-cold cytoplasmic lysis buffer (Table 1) per 107 cells, and pipette up and down.</li>
	<li>Incubate on ice for 20&#160;min.</li>
	<li>Spin for 10&#160;min at full speed in a microcentrifuge at 4&#160;&#176;C.</li>
	<li>Discard pellet. Transfer supernatant into new 1.5&#160;ml microcentrifuge tube and keep on ice.</li>
	<li>Determine protein concentration (usually 1 - 10&#160;&#956;g/&#956;l) using the Bradford assay23.</li>
	<li>Aliquot and store cell extracts at -80&#160;&#176;C until use.</li>
</ol>
2. Preparation of Protein Extracts from Mouse Liver and Spleen
<ol>
	<li>Euthanize a mouse with CO2 inhalation.</li>
	<li>Lay the euthanized animal on a clean pad over a dissecting board. Open the abdomen with scissors.</li>
	<li>Dissect the liver and the spleen by using scissors and forceps, and rinse each tissue in approximately 50&#160;ml ice-cold PBS.</li>
	<li>Immediately cut tissues into small pieces with a scalpel (for example: approximately 1 - 2&#160;mm3).</li>
	<li>Without delay, put pieces of tissues in a fresh cryotube and then snap freeze them in liquid nitrogen. Store snap-frozen tissue aliquots at -80&#160;&#176;C until use.</li>
	<li>Homogenize one piece of frozen tissue (approximately 1 - 2&#160;mm3) in 0.25 - 0.5&#160;ml of ice-cold cytoplasmic lysis buffer (Table 1) with a tissue homogenizer for 10&#160;sec.</li>
	<li>Transfer homogenate to 1.5&#160;ml microcentrifuge tube and chill on ice for 20&#160;min.</li>
	<li>Spin for 10&#160;min at full speed in a microcentrifuge at 4&#160;&#176;C.</li>
	<li>Discard pellet and transfer supernatant into new 1.5 ml microcentrifuge tube. Keep on ice.</li>
	<li>Determine protein concentration (usually 1 - 10&#160;&#956;g/&#956;l) using the Bradford assay23.</li>
	<li>Aliquot and store cell extracts at -80&#160;&#176;C until use.</li>
</ol>
3. Preparation of Radiolabeled IRE-probe
<ol>
	<li>Linearize the IRE-containing plasmid I-12.CAT22 by incubating at 37&#160;&#176;C for 1&#160;hr with the restriction endonuclease XbaI (1&#160;U per &#956;g of plasmid), which cleaves downstream of the IRE sequence. The linearized plasmid will be used as template for in vitro transcription.</li>
	<li>Set up an in vitro transcription reaction in a total volume of 20&#160;&#956;l. Use the stock solutions shown in Table 2 and add: 1&#160;&#956;l linearized plasmid template, 4&#160;&#956;l transcription buffer; 1&#160;&#956;l mix of ATP/CTP/GTP mix, 10&#160;&#956;l [&#945;-32P]-UTP, 2&#160;&#956;l dithiothreitol, 1&#160;&#956;l RNase inhibitor and 1&#160;&#956;l T7 RNA polymerase. Mix by pipetting up and down.</li>
	<li>Incubate at 40&#160;&#176;C for 1&#160;hr24.</li>
</ol>
4. Purification of Radiolabeled IRE-probe
<ol>
	<li>Terminate in vitro transcription reaction by adding 1&#160;&#956;l of 0.5&#160;M EDTA, pH&#160;8. Mix by pipetting up and down.</li>
	<li>Add 10&#160;&#956;l of 10&#160;mg/ml tRNA, as carrier for better precipitation. Mix by pipetting up and down.</li>
	<li>Add 82.5&#160;&#956;l of 3&#160;M ammonium acetate. Mix by vortexing.</li>
	<li>Add 273&#160;&#956;l of ethanol. Mix by vortexing.</li>
	<li>Let stand at RT for 5&#160;min.</li>
	<li>Spin for 10&#160;min at full speed in a microcentrifuge at RT. Discard supernatant.</li>
	<li>Wash pellet with 100&#160;&#956;l of 70% ethanol.</li>
	<li>Spin for 10&#160;min at full speed in a microcentrifuge at RT. Discard supernatant.</li>
	<li>Air dry pellet for 10&#160;min.</li>
	<li>Resuspend pellet in 100&#160;&#956;l of double distilled, previously autoclaved H2O.</li>
	<li>Quantify radioactivity in a liquid scintillation counter, aliquot radiolabeled IRE probe and store at -80&#160;&#176;C until use. Frozen aliquots can be used for up to 3&#160;weeks.</li>
</ol>
5. Preparation of a native polyacrylamide gel for EMSA
<ol>
	<li>Assemble the gel (16 x 16&#160;cm) by using 1.5&#160;mm spacers and comb.</li>
	<li>To prepare a 6% native polyacrylamide gel, use the stock solutions shown in Table 3. Mix 7.5&#160;ml of 40% acrylamide:bisacrylamide, 5&#160;ml of 5x TBE and 37.5&#160;ml double distilled H2O.</li>
	<li>Add 0.5&#160;ml of 10% freshly prepared ammonium persulfate (APS) and 25&#160;&#956;l tetramethylethylenediamine (TEMED).</li>
	<li>Immediately pour the acrylamide solution to the gel and let it polymerize. Wait for approximately 30&#160;min.</li>
	<li>Assemble the electrophoresis apparatus with the gel, fill the tanks with 0.5x TBE and connect with the power supply.</li>
</ol>
6. Electrophoretic Mobility Shift Assay
<ol>
	<li>Dilute 25&#160;&#956;g of protein extract from cells or tissues with cytoplasmic lysis buffer (Table 1) in a total volume of in 10&#160;&#956;l (lower protein concentrations may also be used). Keep on ice. If necessary, add 1&#160;&#956;l of 1:4 diluted 2-mercaptoethanol (2-ME) to activate dormant IRP1 (final concentration: 2%)25.</li>
	<li>Dilute the radiolabeled IRE probe in double distilled H2O to 200,000&#160;cpm/&#956;l, heat denature at 95&#160;&#176;C for 1&#160;min, and cool down at RT for at least 5&#160;min.</li>
	<li>Set up an EMSA reaction by adding 1&#160;&#956;l of radiolabeled IRE probe to the protein extract.</li>
	<li>Incubate for 20&#160;min at RT.</li>
	<li>Add 1&#160;&#956;l of 50&#160;mg/ml heparin to the reaction (to inhibit non-specific protein interactions with the probe9) and continue the incubation for another 10&#160;min. Aliquot the stock solution of heparin (50&#160;mg/ml) and store at -80&#160;&#176;C.</li>
	<li>When using long radiolabeled probes (&#62;60 nucleotides), add 1&#160;&#956;l RNase T1 (1&#160;U/&#956;l) and incubate for 10&#160;min at RT to reduce non-specific protein binding to the probe, and to allow better separation of the RNA/protein complex during electrophoresis.</li>
	<li>Add 3&#160;&#956;l of loading buffer (80% glycerol + bromophenol blue), mix and load on the 6% non-denaturing polyacrylamide gel.</li>
	<li>Run the gel for 60&#160;min at 130&#160;V (5&#160;V/cm).</li>
	<li>Transfer the gel onto a large filter paper and dry.</li>
	<li>Expose to a film and develop autoradiography. Exposure time may range from 1&#160;hr (or less) up to O/N.</li>
</ol>

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The authors declare that they have no competing financial interests.

Herein, we describe a protocol that has been developed to study the IRE-binding activities of IRP1 and IRP2, and we show representative data. By using different probes, this protocol can also be adjusted for the study of other RNA-binding proteins. A critical step is the size of the probe. Usage of long probes, which is common when the exact binding site is unknown, can result in RNA/protein complexes that do not migrate differently than the free RNA. In this case, it is advisable to remove unbound RNA by treatment with ...

The EMSA was originally developed to study the association of DNA-binding proteins with target DNA sequences1,2. The principle is similar for RNA/protein interactions3, which is the focus of this article. Briefly, RNA is negatively charged and will migrate towards the anode during non-denaturing electrophoresis in polyacrylamide (or agarose) gels. Migration within the gel depends on the size of the RNA, which is proportional to its charge. Specific binding of a protein to RNA alters its mobility, and the complex migrates slower compared to the free RNA. This is mainly due to an increase in the molecular mass, but also to alterations in the charge and possibly conformation. Utilizing a labeled RNA as probe allows easy monitoring of the &#8220;gel retardation&#8221; or &#8220;bandshift&#8221;. Usage of 32P-labeled RNA probes is very common and offers high sensitivity. The migration of RNA/protein complexes and free RNA are detected by autoradiography. Drawbacks are the short half-life of 32P (14.29 days), the gradual deterioration of the quality of the probe due to radiolysis, the requirement of a radioactivity license and infrastructure for radioactivity work, and potential biosafety concerns. Therefore, alternative non-isotopic methods for labeling the RNA probe have been developed, for instance with fluorophores or biotin, which enable detection by fluorescent or chemiluminescent imaging4,5. Limitations of these methods are the higher cost and often reduced sensitivity compared to isotopic labeling, and the potential of non-isotopic labels to interfere with the RNA/protein interaction. Non-denaturing polyacrylamide gels are suitable for most EMSA applications and are commonly used. On occasion, agarose gels may pose an alternative for the analysis of large complexes.
The major advantage of EMSA is that it combines simplicity, sensitivity, and robustness4. The assay can be completed within a few hours and does not require sophisticated instrumentation. RNA/protein interactions can be detected by EMSA at concentrations as low as 0.1 nM or less, and within a broad range of binding conditions (pH&#160;4.0 - 9.5, monovalent salt concentration 1 - 300&#160;mM, and temperature 0 - 60&#160;&#176;C).
RNA/protein complex formation can also be studied by the filter-binding assay. This is a simple, fast, and inexpensive procedure based on the retention of RNA/protein complexes in a nitrocellulose filter, while a free RNA probe passes through6. Compared to EMSA, it is limited in cases where the RNA probe contains multiple binding sites, or the crude extract contains more than one RNA-binding proteins that bind to the probe at the same site. While multiple RNA/protein interactions will escape detection by the filter-binding assay, they can be readily visualized by EMSA. In some cases, visualization is even possible when two RNA/protein complexes co-migrate (for instance, human IRP1/IRE and IRP2/IRE complexes), by adding an antibody against one of the RNA-binding proteins to the EMSA reaction, yielding further retardation on the gel (&#8220;supershift&#8221;)7.
The EMSA has been widely used to study IRP1 and IRP2, which are post-transcriptional regulators of iron metabolism8-10. They operate by binding to IREs, phylogenetically conserved hairpin structures within the UTRs of several mRNAs11. IREs were first discovered in the mRNAs encoding ferritin12 and transferrin receptor 1 (TfR1)13, proteins of iron storage and uptake, respectively. Later on, IREs were found in erythroid-specific aminolevulinate synthase (ALAS2)14, mitochondrial aconitase15, ferroportin16, divalent metal transporter 1 (DMT1)17, hypoxia inducible factor 2&#945; (HIF2&#945;)18, and other mRNAs19-21. The prototype H- and L-ferritin mRNAs contain one IRE in their 5&#8217; UTR, while TfR1 mRNA contains multiple IREs in its 3&#8217; UTR. IRE/IRP interactions specifically inhibit ferritin mRNA translation by sterically blocking its association of the 43S ribosomal subunit; moreover, they stabilize TfR1 mRNA against endonucleolytic cleavage. IRP1 and IRP2 share extensive sequence similarity and exhibit high IRE-binding activity in iron-starved cells. In iron-replete cells, IRP1 assembles a cubane Fe-S cluster that converts it to cytosolic aconitase at the expense of its IRE-binding activity, while IRP2 undergoes proteasomal degradation. Thus, the IRE/IRP interactions depend upon the cellular iron status, but are also regulated by other signals, such as H2O2, nitric oxide (NO) or hypoxia. Here, we describe the protocol for assessing IRE-binding activity from crude cell and tissue extracts by EMSA. We used a 32P-labeled H-ferritin IRE probe that was generated by in vitro transcription from a plasmid DNA template (I-12.CAT), where the IRE sequence was originally introduced in sense orientation downstream of the T7 RNA polymerase site by cloning of annealed synthetic oligonucleotides 22.

<table><tbody><tr><td>leupeptin</td><td>SIGMA</td><td>L2884</td><td></td></tr><tr><td>PMSF</td><td>SIGMA</td><td>78830</td><td></td></tr><tr><td>BioRad Protein Assay</td><td>BIORAD</td><td>500-0006</td><td></td></tr><tr><td>T7 RNA polymerase</td><td>Thermoscientific</td><td>EPO111</td><td></td></tr><tr><td>RNase Inhibitor</td><td>Invitrogen</td><td>15518-012</td><td></td></tr><tr><td>UTP [alpha-&lt;sup&gt;32&lt;/sup&gt;P]</td><td>Perkin-Elmer</td><td>NEG507H</td><td></td></tr><tr><td>Scintillation liquid</td><td>Beckman Coulter</td><td>141349</td><td></td></tr><tr><td>heparin</td><td>SIGMA</td><td>H0777</td><td></td></tr><tr><td>Rnase T1</td><td>Thermoscientific</td><td>EN0541</td><td></td></tr><tr><td>Name of the Equipment</td><td></td><td></td><td></td></tr><tr><td>Tissue Ruptor</td><td>Qiagen</td><td>9001271</td><td></td></tr><tr><td>Scintillation counter</td><td>Beckman Coulter</td><td>LS6500</td><td></td></tr><tr><td>Protean II xi Cell</td><td>BIORAD</td><td>165-1834</td><td></td></tr><tr><td>20 wells combs</td><td>BIORAD</td><td>165-1868</td><td>1.5 mm thick</td></tr><tr><td>1.5 mm spacers</td><td>BIORAD</td><td>165-1849</td><td></td></tr><tr><td>PowerPac</td><td>BIORAD</td><td>164-5070</td><td></td></tr></tbody></table>

electrophoretic mobility shift assay (emsa) for the study of rna-protein interactions: the ire/irp example

RNA/protein interactions are critical for post-transcriptional regulatory pathways. Among the best-characterized cytosolic RNA-binding proteins are iron regulatory proteins, IRP1 and IRP2. They bind to iron responsive elements (IREs) within the untranslated regions (UTRs) of several target mRNAs, thereby controlling the mRNAs translation or stability. IRE/IRP interactions have been widely studied by EMSA. Here, we describe the EMSA protocol for analyzing the IRE-binding activity of IRP1 and IRP2, which can be generalized to assess the activity of other RNA-binding proteins as well. A crude protein lysate containing an RNA-binding protein, or a purified preparation of this protein, is incubated with an excess of32 P-labeled RNA probe, allowing for complex formation. Heparin is added to preclude non-specific protein to probe binding. Subsequently, the mixture is analyzed by non-denaturing electrophoresis on a polyacrylamide gel. The free probe migrates fast, while the RNA/protein complex exhibits retarded mobility; hence, the procedure is also called &#8220;gel retardation&#8221; or &#8220;bandshift&#8221; assay. After completion of the electrophoresis, the gel is dried and RNA/protein complexes, as well as free probe, are detected by autoradiography. The overall goal of the protocol is to detect and quantify IRE/IRP and other RNA/protein interactions. Moreover, EMSA can also be used to determine specificity, binding affinity, and stoichiometry of the RNA/protein interaction under investigation.

A radiolabeled IRE probe was prepared, as described in sections 3 and 4 of the protocol. The sequence of the probe was 5&#39;-GGGCGAAUUC GAGCUCGGUA CCCGGGGAUC CUGCUUCAAC AGUGCUUGGA CGGAUCCU-3&#39;; the bolded nucleotides represent an unpaired C residue and the loop, which are critical IRE features. The specific radioactivity of the probe was 4.5 x 109 cpm/&#956;g of RNA.
To assess the effects of iron perturbations on IRE-binding activity, murine RAW...

Watch this Scientific Journal Video about Electrophoretic Mobility Shift Assay EMSA for the Study of RNA-Protein Interactions: The IRE/IRP Example at JoVE.com

Electrophoretic Mobility Shift Assay EMSA for the Study of RNA-Protein Interactions: The IRE/IRP Example

Here we present a protocol to analyze RNA/protein interactions. The electrophoretic mobility shift assay (EMSA) is based on the differential migration of RNA/protein complexes and free RNA during native gel electrophoresis. By using a radiolabeled RNA probe, RNA/protein complexes can be visualized by autoradiography.

Electrophoretic Mobility Shift Assay (EMSA) for the Study of RNA-Protein Interactions: The IRE/IRP Example

RNA/protein interactions are critical for post-transcriptional regulatory pathways. Among the best-characterized cytosolic RNA-binding proteins are iron ...

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Biology

53.5K Views.  Jewish General Hospital. Here we present a protocol to analyze RNA/protein interactions. The electrophoretic mobility shift assay (EMSA) is based on the differential migration of RNA/protein complexes and free RNA during native gel electrophoresis. By using a radiolabeled RNA probe, RNA/protein complexes can be visualized by autoradiography.

Video: Electrophoretic Mobility Shift Assay EMSA for the Study of RNA-Protein Interactions: The IRE/IRP Example

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

An Optimized Protocol for Electrophoretic Mobility Shift Assay Using Infrared Fluorescent Dye-labeled Oligonucleotides

Electrophoretic Mobility Shift Assays (EMSA) are an instrumental tool to characterize the interactions between proteins and their target DNA sequences. Radioactivity has been the predominant method of DNA labeling in EMSAs. However, recent advances in fluorescent dyes and scanning methods have prompted the use of fluorescent tagging of DNA as an alternative to radioactivity for the advantages of easy handling, saving time, reducing cost, and improving safety. We have recently used fluorescent EMSA (fEMSA) to successfully address an important biological question. Our fEMSA analysis provides mechanistic insight into the effect of a missense mutation, G73E, in the highly conserved HMG transcription factor SOX-2 on olfactory neuron type diversification. We found that mutant SOX-2G73E protein alters specific DNA binding activity, thereby causing olfactory neuron identity transformation. Here, we present an optimized and cost-effective step-by-step protocol for fEMSA using infrared fluorescent dye-labeled oligonucleotides containing the LIM-4/SOX-2 adjacent target sites and purified SOX-2 proteins (WT and mutant SOX-2G73E proteins) as a biological example.

We describe here an optimized protocol of fluorescent Electrophoretic Mobility Shift Assays (fEMSA) using purified SOX-2 proteins together with infrared fluorescent dye-labeled DNA probes as a case study to tackle an important biological question.

Electrophoretic Mobility Shift Assays (EMSA) are an instrumental tool to characterize the interactions between proteins and their target DNA sequences. ...

Biochemistry

Horizontal Gel Electrophoresis for Enhanced Detection of Protein-RNA Complexes

Native polyacrylamide gel electrophoresis is a fundamental tool of molecular biology that has been used extensively for the biochemical analysis of RNA-protein interactions. These interactions have been traditionally analyzed with polyacrylamide gels generated between two glass plates and samples electrophoresed vertically. However, polyacrylamide gels cast in trays and electrophoresed horizontally offers several advantages. For example, horizontal gels used to analyze complexes between fluorescent RNA substrates and specific proteins can be imaged multiple times as electrophoresis progresses. This provides the unique opportunity to monitor RNA-protein complexes at several points during the experiment. In addition, horizontal gel electrophoresis makes it possible to analyze many samples in parallel. This can greatly facilitate time course experiments as well as analyzing multiple reactions simultaneously to compare different components and conditions. Here we provide a detailed protocol for generating and using horizontal native gel electrophoresis for analyzing RNA-Protein interactions.

Native polyacrylamide gel electrophoresis is a fundamental tool for analyzing RNA-protein interactions. Traditionally most experiments have used vertical gels. However, horizontal gels provide several advantages, such as the opportunity to monitor complexes during electrophoresis. We provide a detailed protocol for generating and using horizontal native gel electrophoresis.

Native polyacrylamide gel electrophoresis is a fundamental tool of molecular biology that has been used extensively for the biochemical analysis of ...

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 ...

Studying RNA Interactors of Protein Kinase RNA-Activated during the Mammalian Cell Cycle

Protein kinase RNA-activated (PKR) is a member of the innate immune response proteins and recognizes the double-stranded secondary structure of viral RNAs. When bound to viral double-stranded RNAs (dsRNAs), PKR undergoes dimerization and subsequent autophosphorylation. Phosphorylated PKR (pPKR) becomes active and induces phosphorylation of the alpha subunit of eukaryotic initiation factor 2 (eIF2&#945;) to suppress global translation. Increasing evidence suggests that PKR can be activated under physiological conditions such as during the cell cycle or under various stress conditions without infection. However, our understanding of the RNA activators of PKR is limited due to the lack of a standardized experimental method to capture and analyze PKR-interacting dsRNAs. Here, we present an experimental protocol to specifically enrich and analyze PKR bound RNAs during the cell cycle using HeLa cells. We utilize the efficient crosslinking activity of formaldehyde to fix PKR-RNA complexes and isolate them via immunoprecipitation. PKR co-immunoprecipitated RNAs can then be further processed to generate a high-throughput sequencing library. One major class of PKR-interacting cellular dsRNAs is mitochondrial RNAs (mtRNAs), which can exist as intermolecular dsRNAs through complementary interaction between the heavy-strand and the light-strand RNAs. To study the strandedness of these duplex mtRNAs, we also present a protocol for strand-specific qRT-PCR. Our protocol is optimized for the analysis of PKR-bound RNAs, but it can be easily modified to study cellular dsRNAs or RNA-interactors of other dsRNA binding proteins.

We present experimental approaches for studying RNA-interactors of double-stranded RNA binding protein kinase RNA-activated (PKR) during the mammalian cell cycle using HeLa cells. This method utilizes formaldehyde to crosslink RNA-PKR complexes and immunoprecipitation to enrich PKR-bound RNAs. These RNAs can be further analyzed through high-throughput sequencing or qRT-PCR.

Protein kinase RNA-activated (PKR) is a member of the innate immune response proteins and recognizes the double-stranded secondary structure of viral ...

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.  



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.

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

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 ...

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 ...

Electrophoretic Mobility Shift Assay for Detection of Transcription Factor-DNA Complexes

Source: Hsieh, Y. W. et al., <a href="https://app.jove.com/v/54863">An Optimized Protocol for Electrophoretic Mobility Shift Assay Using Infrared Fluorescent Dye-labeled Oligonucleotides.</a> J. Vis. Exp. (2016)
In this video, we demonstrate the electrophoretic mobility shift assay to detect protein-DNA interactions using infrared fluorescent-labeled DNA probes. When the protein binds to the DNA probe, it forms a large complex that migrates slowly on and appears as a shifted band on the gel.

Source: Hsieh, Y. W. et al., An Optimized Protocol for Electrophoretic Mobility Shift Assay Using Infrared Fluorescent Dye-labeled Oligonucleotides. J. ...

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Biological Techniques

Biomolecular Interaction Detection Techniques

Protein-nucleic acid interactions

SELEX-Based In Vitro Binding Assay to Identify RNA-Protein Interactions

Source: Singh, R. <a href="https://app.jove.com/v/59635">Exploring Sequence Space to Identify Binding Sites for Regulatory RNA-Binding Proteins.</a> J. Vis. Exp. ( (2019).
In this video, we describe the systematic evolution of ligands by the exponential enrichment (SELEX) method to identify specific RNA-binding sequences for a target protein of interest. The protein is incubated with a large pool of randomized RNA sequences, and the protein-binding RNA sequences are isolated, PCR-amplified, and sequenced to identify their protein-binding sites.