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 pre...

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 &#38; 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 ab...

Watch this Scientific Journal Video about Exploring Sequence Space to Identify Binding Sites for Regulatory RNA-Binding Proteins at JoVE.com

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

Universidad de Cartagena UDEC

Research

JoVE Journal

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

Methyl-binding DNA capture Sequencing for Patient Tissues

Methylation is one of the essential epigenetic modifications to the DNA, which is responsible for the precise regulation of genes required for stable development and differentiation of different tissue types. Dysregulation of this process is often the hallmark of various diseases like cancer. Here, we outline one of the recent sequencing techniques, Methyl-Binding DNA Capture sequencing (MBDCap-seq), used to quantify methylation in various normal and disease tissues for large patient cohorts. We describe a detailed protocol of this affinity enrichment approach along with a bioinformatics pipeline to achieve optimal quantification. This technique has been used to sequence hundreds of patients across various cancer types as a part of the 1,000 methylome project (Cancer Methylome System).

Here we present a protocol to investigate genome wide DNA methylation in large scale clinical patient screening studies using the Methyl-Binding DNA Capture sequencing (MBDCap-seq or MBD-seq) technology and the subsequent bioinformatics analysis pipeline.

Methylation is one of the essential epigenetic modifications to the DNA, which is responsible for the precise regulation of genes required for stable ...

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 ...

Conjugative Mating Assays for Sequence-specific Analysis of Transfer Proteins Involved in Bacterial Conjugation

The transfer of genetic material by bacterial conjugation is a process that takes place via complexes formed by specific transfer proteins. In Escherichia coli, these transfer proteins make up a DNA transfer machinery known as the mating pair formation, or DNA transfer complex, which facilitates conjugative plasmid transfer. The objective of this paper is to provide a method that can be used to determine the role of a specific transfer protein that is involved in conjugation using a series of deletions and/or point mutations in combination with mating assays. The target gene is knocked out on the conjugative plasmid and is then provided in trans through the use of a small recovery plasmid harboring the target gene. Mutations affecting the target gene on the recovery plasmid can reveal information about functional aspects of the target protein that result in the alteration of mating efficiency of donor cells harboring the mutated gene. Alterations in mating efficiency provide insight into the role and importance of the particular transfer protein, or a region therein, in facilitating conjugative DNA transfer. Coupling this mating assay with detailed three-dimensional structural studies will provide a comprehensive understanding of the function of the conjugative transfer protein as well as provide a means for identifying and characterizing regions of protein-protein interaction.

Here, we present a protocol to knockout a gene of interest involved in plasmid conjugation and subsequently analyze the impact of its absence using mating assays. The function of the gene is further explored to a specific region of its sequence using deletion or point mutations.

The transfer of genetic material by bacterial conjugation is a process that takes place via complexes formed by specific transfer proteins. In Escherichia ...

Analysis of Cap-binding Proteins in Human Cells Exposed to Physiological Oxygen Conditions

Translational control is a focal point of gene regulation, especially during periods of cellular stress. Cap-dependent translation via the eIF4F complex is by far the most common pathway to initiate protein synthesis in eukaryotic cells, but stress-specific variations of this complex are now emerging. Purifying cap-binding proteins with an affinity resin composed of Agarose-linked m7GTP (a 5' mRNA cap analog) is a useful tool to identify factors involved in the regulation of translation initiation. Hypoxia (low oxygen) is a cellular stress encountered during fetal development and tumor progression, and is highly dependent on translation regulation. Furthermore, it was recently reported that human adult organs have a lower oxygen content (physioxia 1-9% oxygen) that is closer to hypoxia than the ambient air where cells are routinely cultured. With the ongoing characterization of a hypoxic eIF4F complex (eIF4FH), there is increasing interest in understanding oxygen-dependent translation initiation through the 5' mRNA cap. We have recently developed a human cell culture method to analyze cap-binding proteins that are regulated by oxygen availability. This protocol emphasizes that cell culture and lysis be performed in a hypoxia workstation to eliminate exposure to oxygen. Cells must be incubated for at least 24 hr for the liquid media to equilibrate with the atmosphere within the workstation. To avoid this limitation, pre-conditioned media (de-oxygenated) can be added to cells if shorter time points are required. Certain cap-binding proteins require interactions with a second base or can hydrolyze the m7GTP, therefore some cap interactors may be missed in the purification process. Agarose-linked to enzymatically resistant cap analogs may be substituted in this protocol. This method allows the user to identify novel oxygen-regulated translation factors involved in cap-dependent translation.

Here, we present human cell culture protocols to analyze translation initiation factors that bind the 5' cap of mRNA during physiological oxygen conditions. This method utilizes an Agarose-linked m7GTP cap analog and is suitable to investigate cap-binding factors and their interacting partners.

Translational control is a focal point of gene regulation, especially during periods of cellular stress. Cap-dependent translation via the eIF4F complex ...

Retroviral Scanning: Mapping MLV Integration Sites to Define Cell-specific Regulatory Regions

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV integration sites can be used as functional markers of active regulatory elements. Here, we present a retroviral scanning tool, which allows the genome-wide identification of cell-specific enhancers and promoters. Briefly, the target cell population is transduced with an mLV-derived vector and genomic DNA is digested with a frequently cutting restriction enzyme. After ligation of genomic fragments with a compatible DNA linker, linker-mediated polymerase chain reaction (LM-PCR) allows the amplification of the virus-host genome junctions. Massive sequencing of the amplicons is used to define the mLV integration profile genome-wide. Finally, clusters of recurrent integrations are defined to identify cell-specific regulatory regions, responsible for the activation of cell-type specific transcriptional programs.
The retroviral scanning tool allows the genome-wide identification of cell-specific promoters and enhancers in prospectively isolated target cell populations. Notably, retroviral scanning represents an instrumental technique for the retrospective identification of rare populations (e.g. somatic stem cells) that lack robust markers for prospective isolation.

Here, we describe a protocol for genome-wide mapping of the integration sites of Moloney murine leukemia virus-based retroviral vectors in human cells.

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV ...

Real-time Analysis of Transcription Factor Binding, Transcription, Translation, and Turnover to Display Global Events During Cellular Activation

Upon activation, cells rapidly change their functional programs and, thereby, their gene expression profile. Massive changes in gene expression occur, for example, during cellular differentiation, morphogenesis, and functional stimulation (such as activation of immune cells), or after exposure to drugs and other factors from the local environment. Depending on the stimulus and cell type, these changes occur rapidly and at any possible level of gene regulation. Displaying all molecular processes of a responding cell to a certain type of stimulus/drug is one of the hardest tasks in molecular biology. Here, we describe a protocol that enables the simultaneous analysis of multiple layers of gene regulation. We compare, in particular, transcription factor binding (Chromatin-immunoprecipitation-sequencing (ChIP-seq)), de novo transcription (4-thiouridine-sequencing (4sU-seq)), mRNA processing, and turnover as well as translation (ribosome profiling). By combining these methods, it is possible to display a detailed and genome-wide course of action.
Sequencing newly transcribed RNA is especially recommended when analyzing rapidly adapting or changing systems, since this depicts the transcriptional activity of all genes during the time of 4sU exposure (irrespective of whether they are up- or downregulated). The combinatorial use of total RNA-seq and ribosome profiling additionally allows the calculation of RNA turnover and translation rates. Bioinformatic analysis of high-throughput sequencing results allows for many means for analysis and interpretation of the data. The generated data also enables tracking co-transcriptional and alternative splicing, just to mention a few possible outcomes.
The combined approach described here can be applied for different model organisms or cell types, including primary cells. Furthermore, we provide detailed protocols for each method used, including quality controls, and discuss potential problems and pitfalls.

This protocol describes the combinatorial use of ChIP-seq, 4sU-seq, total RNA-seq, and ribosome profiling for cell lines and primary cells. It enables tracking changes in transcription-factor binding, de novo transcription, RNA processing, turnover and translation over time, and displaying the overall course of events in activated and/or rapidly changing cells.

Upon activation, cells rapidly change their functional programs and, thereby, their gene expression profile. Massive changes in gene expression occur, for ...

Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly defined states of chromatin organization. The dynamic architecture of the nucleus is controlled by multiple mechanisms and shapes the transcriptional output programs. It is, therefore, important to determine locus-specific chromatin accessibility in a reliable fashion that is preferably independent from antibodies, which can be a potentially confounding source of experimental variability. Chromatin accessibility can be measured by various methods, including the Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) assay, that allow the determination of general chromatin accessibility in a relatively low number of cells. Here we describe a FAIRE protocol that allows simple, reliable, and fast identification of genomic regions with a low protein occupancy. In this method, the DNA is covalently bound to the chromatin proteins using formaldehyde as a crosslinking agent and sheared to small pieces. The free DNA is afterwards enriched using phenol:chloroform extraction. The ratio of free DNA is determined by quantitative polymerase chain reaction (qPCR) or DNA sequencing (DNA-seq) compared to a control sample representing total DNA. The regions with a looser chromatin structure are enriched in the free DNA sample, thus allowing the identification of genomic regions with lower chromatin compaction.

The plasticity of nuclear architecture is controlled by dynamic epigenetic mechanisms including non-coding RNAs, DNA methylation, nucleosome repositioning as well as histone composition and modification. Here we describe a Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) protocol which allows the determination of chromatin accessibility in a reproducible, inexpensive, and straightforward manner.

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly ...

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 ...

Exploring the Effects of Spaceflight on Mouse Physiology using the Open Access NASA GeneLab Platform

Performing biological experiments in space requires special accommodations and procedures to ensure that these investigations are performed effectively and efficiently. Moreover, given the infrequency of these experiments it is imperative that their impacts be maximized. The rapid advancement of omics technologies offers an opportunity to dramatically increase the volume of data produced from precious spaceflight specimens. To capitalize on this, NASA has developed the GeneLab platform to provide unrestricted access to spaceflight omics data and encourage its widespread analysis. Rodents (both rats and mice) are common model organisms used by scientists to investigate space-related biological impacts. The enclosure that house rodents during spaceflight are called Rodent Habitats (formerly Animal Enclosure Modules), and are substantially different from standard vivarium cages in their dimensions, air flow, and access to water and food. In addition, due to environmental and atmospheric conditions on the International Space Station (ISS), animals are exposed to a higher CO2 concentration. We recently reported that mice in the Rodent Habitats experience large changes in their transcriptome irrespective of whether animals were on the ground or in space. Furthermore, these changes were consistent with a hypoxic response, potentially driven by higher CO2 concentrations. Here we describe how a typical rodent experiment is performed in space, how omics data from these experiments can be accessed through the GeneLab platform, and how to identify key factors in this data. Using this process, any individual can make critical discoveries that could change the design of future space missions and activities.

The NASA GeneLab platform provides unfettered access to precious omics data from biological spaceflight experiments. We describe how a typical mouse experiment is conducted in space and how data from such experiments can be accessed and analyzed.

Performing biological experiments in space requires special accommodations and procedures to ensure that these investigations are performed effectively ...

Novel Sequence Discovery by Subtractive Genomics

Subtractive genomics can be used in any research where the goal is to identify the sequence of a gene, protein, or general region that is embedded in a larger genomic context. Subtractive genomics enables a researcher to isolate a target sequence of interest (T) by comprehensive sequencing and subtracting out known genetic elements (reference, R). The method can be used to identify novel sequences such as mitochondria, chloroplasts, viruses, or germline restricted chromosomes, and is particularly useful when T cannot be easily isolated from R. Beginning with the comprehensive genomic data (R + T), the method uses Basic Local Alignment Search Tool (BLAST) against a reference sequence, or sequences, to remove the matching known sequences (R), leaving behind the target (T). For subtraction to work best, R should be a relatively complete draft that is missing T. Since sequences remaining after subtraction are tested through quantitative Polymerase Chain Reaction (qPCR), R does not need to be complete for the method to work. Here we link computational steps with experimental steps into a cycle that can be iterated as needed, sequentially removing multiple reference sequences and refining the search for T. The advantage of subtractive genomics is that a completely novel target sequence can be identified even in cases in which physical purification is difficult, impossible, or expensive. A drawback of the method is finding a suitable reference for subtraction and obtaining T-positive and negative samples for qPCR testing. We describe our implementation of the method in the identification of the first gene from the germline-restricted chromosome of zebra finch. In that case computational filtering involved three references (R), sequentially removed over three cycles: an incomplete genomic assembly, raw genomic data, and transcriptomic data.

The purpose of this protocol is to use a combination of computational and bench research to find novel sequences that cannot be easily separated from a co-purifying sequence, which may be only partially known.

Subtractive genomics can be used in any research where the goal is to identify the sequence of a gene, protein, or general region that is embedded in a ...