This project received funding from the I-CORE Program of the Planning and Budgeting Committee and the Israel Science Foundation (Grant No. 152/11), Marie Curie Reintegration Grant No. PCIG11-GA- 2012-321675, and from the European Union&#39;s Horizon 2020 Research and Innovation Program under grant agreement no. 664918 - MRG-Grammar.

1. System Preparation
<ol>
	<li>Design of binding-site plasmids 
	<ol>
		<li>Design the binding site cassette as depicted in Figure 1. Each minigene contains the following parts (5&#39; to 3&#39;): Eagl restriction site, &#8764;40 bases of the 5&#39; end of the kanamycin (Kan) resistance gene, pLac-Ara promoter, ribosome binding site (RBS), AUG of the mCherry gene, a spacer (&#948;), an RBP binding site, 80 bases of the 5&#39; end of the mCherry gene, and an ApaLI restriction site. 
		NOTE: To increase the success rate of the assay, design three binding-site cassettes for each binding site, with spacers consisting of at least one, two, and three bases. See Representative Results section for further guidelines.</li>
	</ol>
	</li>
	<li>Cloning of binding site plasmids
	<ol>
		<li>Order the binding-site cassettes as double-stranded DNA (dsDNA) minigenes. Each minigene is &#8764;500&#8201;bp long and contains an Eagl restriction site and an ApaLI restriction site at the 5&#39; and 3&#39; ends, respectively (see step 1.1.1). 
		NOTE: In this experiment, mini-genes with half of the kanamycin gene were ordered to facilitate screening for positive colonies. However, Gibson assembly17 is also suitable here, in which case the binding site can be ordered as two shorter complementary single-stranded DNA oligos.</li>
		<li>Double-digest both the mini-genes and the target vector with Eagl-HF and ApaLI by the restriction protocol18, and column purify19.</li>
		<li>Ligate the digested minigenes to the binding-site backbone containing the rest of the mCherry reporter gene, terminator, and a kanamycin resistance gene20.</li>
		<li>Transform the ligation solution into Escherichia coli TOP10 cells21.</li>
		<li>Identify positive transformants via Sanger sequencing.
		<ol>
			<li>Design a primer 100 bases upstream to the region of interest (see Table 1 for primer sequences).</li>
			<li>Miniprep a few bacterial colonies22.</li>
			<li>Prepare 5 &#181;L of a 5 mM solution of the primer and 10 &#181;L of the DNA at 80 ng/&#181;L concentration.</li>
			<li>Send the two solution to a convenient facility for Sanger sequencing23.</li>
		</ol>
		</li>
		<li>Store purified plasmids at -20 &#176;C, and bacterial strains as glycerol stocks24, both in the 96-well format. DNA will then be used for transformation into E. coli TOP10 cells containing one of four fusion-RBP plasmids (see step 1.3.5).</li>
	</ol>
	</li>
	<li>Design and construction of the RBP plasmid 
	NOTE: Amino acid and nucleotide sequences of the coat proteins used in this study are listed in Table 2.
	<ol>
		<li>Order the required RBP sequence lacking a stop codon as a custom-ordered dsDNA minigene lacking a stop codon with restriction sites at the ends (Figure 1).</li>
		<li>Clone the tested RBP lacking a stop codon immediately downstream of an inducible promoter and upstream of a fluorescent protein lacking a start codon (Figure 1), similar to steps 1.2.2-1.2.4. Make sure that the RBP plasmid contains a different antibiotic resistance gene than the binding-site plasmid.</li>
		<li>Identify positive transformants via Sanger sequencing, similar to step 1.2.5 (see Table 1 for primer sequences).</li>
		<li>Choose one positive transformant and make it chemically-competent25. Store as glycerol purified plasmids at -20 &#176;C and glycerol stocks of bacterial strains24 at -80 &#176;C in 96-well plates.</li>
		<li>Transform the binding-site plasmids (from step 1.2.6) stored in 96-well plates into chemically-competent bacterial cells already containing an RBP-mCerulean plasmid21. To save time, instead of plating the cells on Petri dishes, plate them using an 8-channel pipettor on 8-lane plates containing Luria-Bertani (LB)26 agar with relevant antibiotics (Kan and Amp). Colonies should appear in 16 h.</li>
		<li>Select a single colony for each double transformant and grow overnight in LB medium with the relevant antibiotics (Kan and Amp) and store as glycerol stocks24 at -80 &#176;C in 96-well plates.</li>
	</ol>
	</li>
</ol>
2. Experiment Setup
NOTE: The protocol presented here was performed using a liquid-handling robotic system in combination with an incubator and a plate reader. Each measurement was carried out for 24 inducer concentrations, with two duplicates for each strain + inducer combination. Using this robotic system, data for 16 strains per day with 24 inducer concentrations was collected. However, if such a device is unavailable, or if fewer experiments are necessary, these can easily be done by hand using an 8-channel multi-pipette and adapting the protocol accordingly. For example, preliminary results for four strains per day with 12 inducer concentrations and four time-points were acquired in this manner.
<ol>
	<li>Prepare, in advance, 1 L of bioassay buffer (BA) by mixing 0.5 g of tryptone, 0.3 mL of glycerol, 5.8 g of NaCl, 50 mL of 1 M MgSO4, 1 mL of 10x phosphate-buffered saline (PBS) buffer pH 7.4, and 950 mL of double distilled water (DDW). Autoclave or sterile filter the BA buffer.</li>
	<li>Grow the double-transformant strains at 37 &#176;C and 250 rpm shaking in 1.5 mL LB with appropriate antibiotics (kanamycin at a final concentration of 25 &#956;g/mL and ampicillin at a final concentration of 100 &#956;g/mL), in 48-well plates, over a period of 18 h (overnight).</li>
	<li>In the morning, make the following preparations.
	<ol>
		<li>Inducer plate. In a clean 96-well plate, prepare wells with semi-poor medium (SPM) consisting of 95% BA and 5% LB26 in the incubator at 37 &#176;C. The number of wells corresponds to the desired number of inducer concentrations. Add C4-HSL to the wells in the inducer plate that will contain the highest inducer concentration (218 nM).</li>
		<li>Program the robot to serially dilute medium from each of the highest-concentration wells into 23 lower concentrations ranging from 0 to 218 nM. The volume of each inducer dilution should be sufficient for all strains (including duplicates).</li>
		<li>While the inducer dilutions are being prepared, warm 180 &#956;L of SPM in the incubator at 37 &#176;C, in 96-well plates.</li>
		<li>Dilute the overnight strains from step 2.2 by a factor of 100 by serial dilutions: first dilute by a factor of 10 by mixing 100 &#956;L of bacteria with 900 &#956;L of SPM in 48-well plates, and then dilute again by a factor of 10 by taking 20 &#956;L from the diluted solution into 180 &#956;L of pre-warmed SPM, in 96-well plates suitable for fluorescent measurements.</li>
		<li>Add the diluted inducer from the inducer plate to the 96-well plates with the diluted strains according to the final concentrations.</li>
	</ol>
	</li>
	<li>Shake the 96-well plates at 37 &#176;C for 6 h, while taking measurements of optical density at 595 nm (OD595), mCherry (560 nm/612 nm) and mCerulean (460 nm/510 nm) fluorescence via a plate reader every 30 min. For normalization purposes, measure growth of SMP with no cells added.</li>
</ol>
3. Preliminary Results Analysis
<ol>
	<li>For each day of experiment, choose a time interval of logarithmic growth according to the measured growth curves, between the linear growth phase and the stationary (T&#173;0, Tfinal). Take approximately 6&#8722;8 time points, while discarding the first and last measurements to avoid error derived from inaccuracy of exponential growth detection (see Figure 2A, top panel). 
	NOTE: Discard strains that show abnormal growth curves or strains where logarithmic growth phase could not be detected and repeat the experiment.</li>
	<li>Calculate the average normalized fluorescence of mCerulean and rate of production of mCherry, from the raw data of both mCerulean and mCherry fluorescence for each inducer concentration (Figure 2A).
	<ol>
		<li>Calculate normalized mCerulean as follows: 
		<img alt="Equation 1" src="/files/ftp_upload/59611/59611eq1.jpg" /> 
		&#8203;where blank(mCerulean) is the mCerulean level [a.u.] for medium only, blank(OD) is the optical density for medium only, and mCerulean and OD are the mCerulean fluorescence and optical density values, respectively.</li>
		<li>Average mCerulean over the different time points (Figure 2B, top two panels) as follows: 
		<img alt="Equation 2" src="/files/ftp_upload/59611/59611eq2.jpg" /> 
		&#8203;where #Time points is the number of data timepoints taken into account, T0 is the time at which the exponential growth phase begins, and Tfinal is the time at which the exponential growth phase ends.</li>
		<li>Calculate mCherry rate of production (Figure 2B, bottom two panels) as follows: 
		<img alt="Equation 3" src="/files/ftp_upload/59611/59611eq3.jpg" /> 
		where mCherry(t) is the mCherry level [a.u.] at time t, OD is the optical density value, T0 is the time at which the exponential growth phase begins, and Tfinal is the time at which the exponential growth phase ends.</li>
	</ol>
	</li>
	<li>Finally, plot the mCherry rate of production as a function of mCerulean, creating dose response curves as a function of RBP-mCerulean fusion fluorescence (Figure 2C). Such plots represent production of the reporter gene as a function of RBP presence in the cell.</li>
</ol>
4. Dose Response Function Fitting Routine and KRBP Extraction
<ol>
	<li>Under the assumption that the ribosome rate of translation with the RBP bound is constant, model the mCherry production rate as follows (see Figure 2D, green line): 
	<img alt="Equation 4" src="/files/ftp_upload/59611/59611eq4.jpg" /> 
	where [x] is the normalized average mCerulean fluorescence calculated according to Eq. 2, mCherry production rate is the value calculated according to Eq. 3, KRBP is the relative binding affinity [a.u.], Kunbound is the ribosome rate of translation with the RBP unbound, n is the cooperativity factor, and C is the base fluorescence [a.u.]. C, n, Kunbound, and KRBP are found by fitting the mCherry production rate data to the model (Eq. 4).</li>
	<li>Using data analysis software, conduct a fitting procedure on plots depicting mCherry production rate as a function of averaged mCerulean (step 3.3), and extract the fit parameters according to the formula in Eq. 4. 
	NOTE: Only fitting results with R2 &#62; 0.6 are taken into account. For those fits, KRBP error is mostly in the range of 0.5% to 20% of KRBP values, for a 0.67 confidence interval, while those with higher KRBP error can be also verified by eye.</li>
	<li>Normalize KRBP values by the respective maximal value of averaged mCerulean for each dose-response function. 
	<img alt="Equation 5" src="/files/ftp_upload/59611/59611eq5.jpg" /> 
	where KRBP in [a.u.] is the value extracted from the fitting procedure in Eq. 4, and max (averaged mCerulean) is the maximal averaged mCerulean signal [a.u] observed for the current strain. 
	NOTE: The normalization facilitates correct comparison of the regulatory effect across strains by eliminating the dependence on the particular maximal RBP expression levels.</li>
</ol>

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</ol>

The authors have nothing to disclose.

The method described in this article facilitates quantitative in vivo measurement of RBP-RNA binding affinity in E. coli cells. The protocol is relatively easy and can be conducted without the use of sophisticated machinery, and data analysis is straightforward. Moreover, the results are produced immediately, without the relatively long wait-time associated with next generation sequencing (NGS) results.
One limitation to this method is that it works only in bacterial cells. However, a...

RNA binding protein (RBP)-based post-transcriptional regulation, specifically characterization of the interaction between RBPs and RNA, has been studied extensively in recent decades. There are multiple examples of translational down-regulation in bacteria originating from RBPs inhibiting, or directly competing with, ribosome binding1,2,3. In the field of synthetic biology, RBP-RNA interactions are emerging as a significant tool for the design of transcription-based genetic circuits4,5. Therefore, there is an increase in demand for characterization of such RBP-RNA interactions in a cellular context.
The most common methods for studying protein-RNA interactions are the electrophoretic mobility shift assay (EMSA)6, which is limited to in vitro settings, and various pull-down assays7, including the CLIP method8,9. While such methods enable the discovery of de novo RNA binding sites, they suffer from drawbacks such as labor-intensive protocols and expensive deep sequencing reactions and may require a specific antibody for RBP pull-down. Due to the susceptible nature of RNA to its environment, many factors can affect RBP-RNA interactions, emphasizing the importance of interrogating RBP-RNA binding in the cellular context. For example, we and others have demonstrated significant differences between RNA structures in vivo and in vitro10,11.
Based on the approach of a previous study12, we recently demonstrated10 that when placing pre-designed binding sites for the capsid RBPs from the bacteriophages GA13, MS214, PP715, and Q&#946;16 in the translation initiation region of a reporter mRNA, reporter expression is strongly repressed. We present a relatively simple and quantitative method, based on this repression phenomenon, to measure the affinity between RBPs and their corresponding RNA binding sites in vivo.

<table><tbody><tr><td>Ampicillin sodium salt</td><td>SIGMA</td><td>A9518</td><td></td></tr><tr><td>Magnesium sulfate (MgSO&lt;sub&gt;4&lt;/sub&gt;)</td><td>ALFA AESAR</td><td>33337</td><td></td></tr><tr><td>48 plates</td><td>Axygen</td><td>P-5ML-48-C-S</td><td></td></tr><tr><td>8-lane plates</td><td>Axygen</td><td>RESMW8I</td><td></td></tr><tr><td>96-well plates</td><td>Axygen</td><td>P-DW-20-C</td><td></td></tr><tr><td>96-well plates for plate reader</td><td>Perkin Elmer</td><td>6005029</td><td></td></tr><tr><td>ApaLI</td><td>NEB</td><td>R0507</td><td></td></tr><tr><td>Binding site sequences</td><td>Gen9 Inc. and Twist Bioscience</td><td></td><td>see Table 1</td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; TOP10 cells</td><td>Invitrogen</td><td>C404006</td><td></td></tr><tr><td>Eagl-HF</td><td>NEB</td><td>R3505</td><td></td></tr><tr><td>Glycerol</td><td>BIO LAB</td><td>071205</td><td></td></tr><tr><td>Incubator</td><td>TECAN</td><td>liconic incubator</td><td></td></tr><tr><td>Kanamycin solfate</td><td>SIGMA</td><td>K4000</td><td></td></tr><tr><td>KpnI- HF</td><td>NEB</td><td>R0142</td><td></td></tr><tr><td>Ligase</td><td>NEB</td><td>B0202S</td><td></td></tr><tr><td>Liquid-handling robotic system</td><td>TECAN</td><td>EVO 100, MCA 96-channel</td><td></td></tr><tr><td>Matlab analysis software</td><td>Mathworks</td><td></td><td></td></tr><tr><td>Multi- pipette 8 lanes</td><td>Axygen</td><td>BR703710</td><td></td></tr><tr><td>N-butanoyl-L-homoserine lactone (C&lt;sub&gt;4&lt;/sub&gt;-HSL)</td><td>cayman</td><td>K40982552 019</td><td></td></tr><tr><td>PBS buffer</td><td>Biological Industries</td><td>020235A</td><td></td></tr><tr><td>Platereader</td><td>TECAN</td><td>Infinite F200 PRO</td><td></td></tr><tr><td>Q5 HotStart Polymerase</td><td>NEB</td><td>M0493</td><td></td></tr><tr><td>RBP seqeunces</td><td>Addgene</td><td>27121 &amp;amp; 40650</td><td>see Table 2</td></tr><tr><td>Sodium Chloride (NaCL)</td><td>BIO LAB</td><td>190305</td><td></td></tr><tr><td>SV Gel and PCR Clean-Up System</td><td>Promega</td><td>A9281</td><td></td></tr><tr><td>Tryptone</td><td>BD</td><td>211705</td><td></td></tr></tbody></table>

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.

The presented method utilizes the competition between an RBP and the ribosome for binding to the mRNA molecule (Figure 1). This competition is reflected by decreasing mCherry levels as a function of increased production of RBP-mCerulean, due to increasing concentrations of inducer. In the case of increasing mCerulean fluorescence, with no significant changes in mCherry, a lack of RBP binding is deduced. Representative results for both a positive and a negativ...

Watch this Scientific Journal Video about An Assay for Quantifying Protein-RNA Binding in Bacteria at JoVE.com

An Assay for Quantifying Protein-RNA Binding in Bacteria

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 ...

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Research

JoVE Journal

Genetics

6.6K Views.  Technion-Israel Institute of Technology. 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.

Video: An Assay for Quantifying Protein-RNA Binding in Bacteria

Rapid Colorimetric Assays to Qualitatively Distinguish RNA and DNA in Biomolecular Samples

Biochemical experimentation generally requires accurate knowledge, at an early stage, of the nucleic acid, protein, and other biomolecular components in potentially heterogeneous specimens. Nucleic acids can be detected via several established approaches, including analytical methods that are spectrophotometric (e.g., A260), fluorometric (e.g., binding of fluorescent dyes), or colorimetric (nucleoside-specific chromogenic chemical reactions).1 Though it cannot readily distinguish RNA from DNA, the A260/A280 ratio is commonly employed, as it offers a simple and rapid2 assessment of the relative content of nucleic acid, which absorbs predominantly near 260 nm and protein, which absorbs primarily near 280 nm. Ratios &lt; 0.8 are taken as indicative of 'pure' protein specimens, while pure nucleic acid (NA) is characterized by ratios &gt; 1.53.
However, there are scenarios in which the protein/NA content cannot be as clearly or reliably inferred from simple uv-vis spectrophotometric measurements. For instance, (i) samples may contain one or more proteins which are relatively devoid of the aromatic amino acids responsible for absorption at &asymp;280 nm (Trp, Tyr, Phe), as is the case with some small RNA-binding proteins, and (ii) samples can exhibit intermediate A260/A280 ratios (~0.8 &lt; ~1.5), where the protein/NA content is far less clear and may even reflect some high-affinity association between the protein and NA components. For such scenarios, we describe herein a suite of colorimetric assays to rapidly distinguish RNA, DNA, and reducing sugars in a potentially mixed sample of biomolecules. The methods rely on the differential sensitivity of pentoses and other carbohydrates to Benedict's, Bial's (orcinol), and Dische's (diphenylamine) reagents; the streamlined protocols can be completed in a matter of minutes, without any additional steps of having to isolate the components. The assays can be performed in parallel to differentiate between RNA and DNA, as well as indicate the presence of free reducing sugars such as glucose, fructose, and ribose (Figure 1).

A suite of colorimetric assays is described for rapidly distinguishing protein, RNA, DNA, and reducing sugars in potentially heterogeneous biomolecular samples.

Biochemical experimentation generally requires accurate knowledge, at an early stage, of the nucleic acid, protein, and other biomolecular components in ...

Chemistry

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 ...

Site-Directed Mutagenesis for In Vitro and In Vivo Experiments Exemplified with RNA Interactions in Escherichia Coli

Site-directed mutagenesis is a technique used to introduce specific mutations in DNA to investigate the interaction between small non-coding ribonucleic acid (sRNA) molecules and target messenger RNAs (mRNAs). In addition, site-directed mutagenesis is used to map specific protein binding sites to RNA. A 2-step and 3-step PCR based introduction of mutations is described. The approach is relevant to all protein-RNA and RNA-RNA interaction studies. In short, the technique relies on designing primers with the desired mutation(s), and through 2 or 3 steps of PCR synthesizing a PCR product with the mutation. The PCR product is then used for cloning. Here, we describe how to perform site-directed mutagenesis with both the 2- and 3-step approach to introduce mutations to the sRNA, McaS, and the mRNA, csgD, to investigate RNA-RNA and RNA-protein interactions. We apply this technique to investigate RNA interactions; however, the technique is applicable to all mutagenesis studies (e.g., DNA-protein interactions, amino-acid substitution/deletion/addition). It is possible to introduce any kind of mutation except for non-natural bases but the technique is only applicable if a PCR product can be used for downstream application (e.g., cloning and template for further PCR).

Site-Directed Mutagenesis for In Vitro and In Vivo Experiments Exemplified with RNA Interactions in Escherichia Coli

Site-directed mutagenesis is a technique used to introduce specific mutations in deoxyribonucleic acid (DNA). This protocol describes how to do site-directed mutagenesis with a 2-step and 3-step polymerase chain reaction (PCR) based approach, which is applicable to any DNA fragment of interest.

Site-directed mutagenesis is a technique used to introduce specific mutations in DNA to investigate the interaction between small non-coding ribonucleic ...

Biochemistry

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 ...

Biology

Qualitative and Quantitative Assays for Detection and Characterization of Protein Antimicrobials

Initial evaluations of large microbial libraries for potential producers of novel antimicrobial proteins require both qualitative and quantitative methods to screen for target enzymes prior to investing greater research effort and resources. The goal of this protocol is to demonstrate two complementary assays for conducting these initial evaluations. The microslide diffusion assay provides an initial or simple detection screen to enable the qualitative and rapid assessment of proteolytic activity against an array of both viable and heat-killed bacterial target substrates. As a counterpart, the increased sensitivity and reproducibility of the dye-release assay provides a quantitative platform for evaluating and comparing environmental influences affecting the hydrolytic activity of protein antimicrobials. The ability to label specific heat-killed cell culture substrates with Remazol brilliant blue R dye expands this capability to tailor the dye-release assay to characterize enzymatic activity of interest.

Here, we present protocols to rapidly screen and characterize enzymes for antimicrobial activity. The microslide diffusion assay and the dye-release assay utilize target bacterial substrates for qualitative and quantitative enzymatic activity evaluation.

Initial evaluations of large microbial libraries for potential producers of novel antimicrobial proteins require both qualitative and quantitative methods ...

Immunology and Infection

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 ...

A Quantitative Assay to Study Protein:DNA Interactions, Discover Transcriptional Regulators of Gene Expression, and Identify Novel Anti-tumor Agents

Many DNA-binding assays such as electrophoretic mobility shift assays (EMSA), chemiluminescent assays, chromatin immunoprecipitation (ChIP)-based assays, and multiwell-based assays are used to measure transcription factor activity. However, these assays are nonquantitative, lack specificity, may involve the use of radiolabeled oligonucleotides, and may not be adaptable for the screening of inhibitors of DNA binding. On the other hand, using a quantitative DNA-binding enzyme-linked immunosorbent assay (D-ELISA) assay, we demonstrate nuclear protein interactions with DNA using the RUNX2 transcription factor that depend on specific association with consensus DNA-binding sequences present on biotin-labeled oligonucleotides. Preparation of cells, extraction of nuclear protein, and design of double stranded oligonucleotides are described. Avidin-coated 96-well plates are fixed with alkaline buffer and incubated with nuclear proteins in nucleotide blocking buffer. Following extensive washing of the plates, specific primary antibody and secondary antibody incubations are followed by the addition of horseradish peroxidase substrate and development of the colorimetric reaction. Stop reaction mode or continuous kinetic monitoring were used to quantitatively measure protein interaction with DNA. We discuss appropriate specificity controls, including treatment with non-specific IgG or without protein or primary antibody. Applications of the assay are described including its utility in drug screening and representative positive and negative results are discussed.

We developed a quantitative DNA-binding, ELISA-based assay to measure transcription factor interactions with DNA. High specificity for the RUNX2 protein was achieved with a consensus DNA-recognition oligonucleotide and specific monoclonal antibody. Colorimetric detection with an enzyme-coupled antibody substrate reaction was monitored in real time.

Many DNA-binding assays such as electrophoretic mobility shift assays (EMSA), chemiluminescent assays, chromatin immunoprecipitation (ChIP)-based assays, ...

Reporter Gene Repression Assay to Study Translational Regulation of a Target Gene

Source: Katz, N., et al. <a href="https://app.jove.com/v/59611">An Assay for Quantifying Protein-RNA Binding in Bacteria.</a> J. Vis. Exp. (2019)
This video demonstrates an in vivo assay in bacteria to study the interaction of RNA-binding proteins (RBPs) with RNA. The bacterial cells are transformed with two plasmid constructs &#8212; a binding-site plasmid encoding an mRNA containing a fluorescent reporter gene downstream of an RBP-binding site &#8212; while an RBP plasmid expresses the RBP proteins under the control of an inducer. Upon inducer-mediated increase in RBP production, the RBPs bind to the mRNA, resulting in translational repression of the reporter via inhibition of ribosome binding.

Source: Katz, N., et al. An Assay for Quantifying Protein-RNA Binding in Bacteria. J. Vis. Exp. (2019)
This video demonstrates an in vivo assay in ...

JoVE Encyclopedia of Experiments

Biological Techniques

Biomolecular Interaction Detection Techniques

Protein-nucleic acid interactions

Differential Radial Capillary Action of Ligand Assay: A High-Throughput Technique to Identify Bacterial Nucleotide Second Messenger-Binding Intracellular Proteins

Source: Schicketanz, M. L., et al. <a href="https://app.jove.com/v/62331">Identifying the Binding Proteins of Small Ligands with the Differential Radial Capillary Action of Ligand Assay (DRaCALA).</a> J. Vis. Exp. (2021)
This video demonstrates the differential radial capillary action of ligand assay &#8212; a technique for systematic high-throughput screening of intracellular nucleotide second messenger-binding proteins. This technique allows for direct binding of the messengers to the overexpressed target protein in the cell lysate &#8212; bypassing the need for protein purification.

Source: Schicketanz, M. L., et al. Identifying the Binding Proteins of Small Ligands with the Differential Radial Capillary Action of Ligand Assay ...

Assay Techniques

Biochemical assays

RNA-Protein Pull-Down Assay to Isolate RNA-Binding Proteins via Affinity Extraction

Pre-wash 50 microliters of streptavidin-magnetic beads per 50 picomoles of RNA. Vortex the tube with streptavidin-magnetic beads for 15 seconds. After vortexing quickly, remove 200 microliters into a clean 1.5-milliliter safe lock tube using cut pipette tips. Next, place the tube on a magnetic stand so that the beads collect at the side of the tube, and wait one minute. Remove the resuspension liquid.
To wash the beads, remove the tube from the magnetic stand at 400 microliters of 0.1 molar sodium hydroxide, 0.05 molar sodium chloride solution, and gently pipette up and down several times. Place the tube back on the magnetic stand and wait one minute, then, collect the supernatant.
Wash the beads with 200 microliters of 100-millimolar sodium chloride, and remove the supernatant. Add 200 microliters of 20 millimolar Tris. Resuspend the beads by pipetting and place the tube on the magnetic stand. After one minute, remove the supernatant. Then, remove the tube from the magnetic stand at 200 microliters of 1x RNA capture buffer and resuspend streptavidin-magnetic beads by briefly vortexing.
Remove 50 microliters of streptavidin-magnetic beads and add it to each labeled-RNA tube using a cut pipette tip. Incubate the tubes for 30 minutes at room temperature on a roller. After the tube has been on the magnetic stand for one minute, remove the supernatant. Add 50 microliters of 20 millimolar Tris to the beads and resuspend them. Then, place the tubes into the magnetic stand. After one minute, remove the supernatant.
Add 100 microliters of 1x protein-RNA binding buffer to the beads and resuspend them. Then, place the tubes back into the magnetic stand. Once the tubes have been in the stand for 1 minute, collect the supernatant. Add 100 microliter of master mix B. Resuspend by gently pipetting up and down and avoid creating bubbles. Incubate reaction tubes for 60 minutes at 4 degrees Celsius on a roller.
Move on to the wash and elution steps. Place the tubes into a magnetic stand. Collect the flow-through and transfer it into a new tube. Pipette 100 microliters of 1X wash buffer on the beads. Resuspend gently. Place them back into the stand. Wait one minute and discard the supernatant. Repeat the step two times.
Add 40 microliters of elution buffer to the magnetic beads. Mix by pipetting up and down and incubate on a tube shaker. Place the tubes into a magnetic stand, wait one minute, and collect eluate sample. Place on ice and use for downstream analysis.

An Assay for Quantifying Protein-RNA Binding in Bacteria

Summary

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