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In this protocol, an RNA sample, fractionated by gel electrophoresis, is transferred from the gel onto a membrane by capillary transfer. Short-wave UV light is used to fix the transferred RNA to the membrane. The membrane is then pretreated to block nonspecific probe-binding sites, and hybridization of the immobilized RNA to a 32P-labeled DNA or RNA probe specific for the mRNA of interest is performed. Finally, the membrane is washed and subjected to autoradiography or phosphorimaging. Because exposure to UV cross-links the RNA to the membrane, the membrane can be stripped and hybridized with other probes. The procedure is suitable for detecting poly(A)+-selected mRNA or mRNA in total cellular RNA if the target transcript is relatively abundant. Using DNA or RNA probes labeled to 1 × 108–10 × 108 cpm/µg, it should be possible to detect ∼5 pg of a specific RNA.

Electrophoretic size fractionation can be used to denature and separate large mRNA molecules (0.5–10 kb) on formaldehyde-containing agarose gels. Formaldehyde contains a carbonyl group that reacts to form Schiff bases with the imino or amino groups of guanine, adenine, and cytosine. These covalent adducts prevent normal base pairing and maintain the RNA in a denatured state. Because these adducts are unstable, formaldehyde must be present in the gel to maintain the RNA in the denatured state. This protocol describes the preparation of an agarose gel with formaldehyde and its setup in a horizontal electrophoresis apparatus. RNA samples are prepared and denatured in a solution of formamide and formaldehyde and, with 0.5- to 10-kb size markers, subjected to electrophoresis through the gel. Following electrophoresis, the gel is stained to visualize RNA markers or rRNA using one of several different types of stains.

This protocol is used to denature and separate large mRNA molecules (0.5–10 kb) on agarose gels by electrophoretic size fractionation. Glyoxal (also called diformyl or ethanedial), the agent responsible for maintaining denaturation in this protocol, contains two carbonyl groups that react to form a cyclic ring structure with the imino and amino groups of guanine. It can also react with the amino groups of adenine and cytidine. When RNA is denatured in the presence of glyoxal, this covalent adduct prevents normal base pairing and maintains the RNA in a denatured state in agarose gels. Once formed, these adducts are stable at room temperature at pH

This protocol describes how to generate and analyze products and intermediates in a pre-mRNA splicing reaction. The reaction relies on the use of labeled, capped, synthetic pre-mRNAs, prepared by in vitro transcription, and Drosophila Kc cell culture nuclear extracts. The pre-mRNA substrate is incubated in the nuclear extract under splicing conditions for 1–2 h. The products of the reaction are purified by phenol:chloroform extraction and precipitation with ethanol, and then loaded directly onto a denaturing urea–acrylamide gel. Visualization of the splicing reactions will reveal the pre-mRNA, the spliced mRNA, and the intermediates generated by the first step of splicing. For inefficient reactions, a more sensitive detection method, such as RNase protection, primer extension, or RT-PCR (reverse transcription–polymerase chain reaction), may be required.

This protocol describes how to prepare nuclear extracts from cultured Drosophila cells. It uses the Kc cell line, which grows without fetal bovine serum and so is suitable for large-scale suspension cultures. The suspension cells are harvested by centrifugation, washed to remove media components, and resuspended in a low-salt (hypotonic) buffer that makes the cells swell in size and easy to homogenize. Usually, nuclear extract preparations are performed on 4–32 L of cells at ∼3.5 × 106 cells/mL.

This protocol describes the detection of small RNAs (∼10–200 nucleotides) by blot hybridization. The RNA samples, denatured in formamide, are separated by denaturing polyacrylamide gel electrophoresis. Because high-percentage polyacrylamide gels are required to separate RNAs in this size range, it is necessary to perform electrophoretic transfer to positively charged nylon membranes. After transfer, the immobilized RNAs are subjected to hybridization with a 32P-radiolabeled DNA or RNA probe and detected by phosphorimaging or autoradiography. This procedure is commonly used to detect small, U-rich spliceosomal small nuclear RNAs (snRNAs) and miRNAs. It should be possible also to detect most miRNAs using high-percentage (e.g., 15%) urea–polyacrylamide gel electrophoresis.

This protocol is used to label RNA molecules (in vitro–synthesized or in vivo–purified RNA molecules) that have free 3′-hydroxyl termini. The reaction is performed in 10 min using yeast poly(A) polymerase and 3′-deoxyadenosine 5′-[α-32P]triphosphate (cordycepin 5′-[α-32P]triphosphate), a chain-terminating nucleotide. At the end of the procedure, the reaction is desalted by gel filtration to remove any unincorporated nucleotides.

This protocol uses T4 polynucleotide kinase to catalyze the transfer of a radiolabeled, terminal (γ) phosphate of ATP to the 5′-hydroxyl terminus of a DNA or RNA molecule. The reaction is very efficient and hence is used as a general method for phosphorylating polynucleotides or oligonucleotides.

It is of interest to be able to define sets of cellular RNAs associated with specific RNA-binding proteins. This “guilt by association” can lead to new insights into how RNA-binding proteins modulate posttranscriptional gene expression of specific target RNAs. To identify these RNAs, antibodies against RNA-binding proteins can be used to immunopurify endogenous RNA–protein complexes from cells, and then the associated RNAs can be characterized. The method described here was developed to identify binding regions on nuclear transcripts for Drosophila heterogeneous nuclear ribonucleoproteins (hnRNPs). An antibody is added to an RNP extract and incubated to allow antigen–antibody complexes to form. The antibody–antigen complexes are then retrieved by binding of the antibody constant region to staphylococcal Protein A immobilized on Sepharose beads. The bead-immobilized complexes are then washed and RNA is prepared. The RNA is used to generate random-primed cDNA, cRNA, and biotinlyated cDNA probes for use on Affymetrix whole-genome Drosophila tiling arrays.

When a large amount (milligram quantities) of RNA is desired (e.g., for making RNAs for RNA interference studies or for structural analyses), it is advantageous to use optimized transcription conditions. The method described here uses high millimolar concentrations of rNTPs, pyrophosphatase, and a specially formulated high-yield transcription buffer. These conditions prevent product (pyrophosphate) inhibition of RNA polymerase, which permits the accumulation of large quantities of RNA.

One of the oldest and simplest (and still very useful) methods for detecting RNA–protein interactions is the filter-binding assay. If a mixture of RNA and protein is passed through a nitrocellulose filter, the protein will be retained and the RNA will pass through. But if the protein is capable of binding RNA, then RNA will be retained on the filter as well. This protocol requires a purified protein (or chromatographic fractions) of interest and 32P-labeled RNA. To perform the assay, the protein sample is serially diluted to several concentrations. It is then mixed with a fixed amount of radioactive RNA and allowed to bind under desired conditions for 30–60 min. The binding reactions are then applied to a 96-well dot-blot apparatus with low vacuum to trap the complexes on three membranes: The top membrane traps aggregates, the middle membrane (nitrocellulose) binds proteins and RNA–protein complexes, and the bottom membrane (which is charged) collects free RNA. After washing and drying, the membranes are exposed to phosphor-imaging screens for quantitation. Alternatively, single, larger filters (that can be counted in a scintillation counter) and a filter manifold can be used.

INTRODUCTIONMore than 95% of total RNA is composed of ribosomal RNAs (rRNAs) (28S, 18S, 5.8S, and 5S). Here, we present a method that is effective in removing rRNA before extraction and purification of total RNA. If you choose to prepare cytoplasmic RNA and wish to analyze any RNA other than rRNA, it is desirable to eliminate the rRNAs by taking advantage of the fact that the ribosomal subunits are very large (40S and 60S). Few, if any, other cellular RNAs are present in such large macromolecular complexes. The vast majority of rRNAs can be removed by sedimentation. Of course, steps must be taken to avoid co-sedimentation of desired RNAs. Co-sedimentation can be greatly reduced by first dissociating ribosomes into their respective subunits by EDTA treatment. The subunits are then “cleaned” by treatment with high salt and nonionic detergent. Ribosomal subunits remain intact under these conditions and can be sedimented free of other RNAs. Subsequently, the remaining RNAs (messenger RNAs [mRNAs] and all other RNAs) can be purified and analyzed by a variety of methods.

INTRODUCTIONAfter any RNA purification by any method, it is necessary to ascertain both the amount (yield) and the integrity (quality) of the RNA obtained. This is essential for any subsequent analysis and critical for any comparative analyses. This article discusses several methods for RNA quantitation and provides tips for each.

INTRODUCTIONDNA contamination of RNA preparations can present significant problems, especially for polymerase chain reaction (PCR)-based applications. No RNA extraction procedure excludes DNA entirely. Cytoplasmic RNA can be contaminated with DNA because of nuclei breakage during preparation. Moreover, TRIzol preparations do not exclude plasmids or other small DNA fragments. This article describes the reliable and effective method of eliminating DNA from RNA preparations via DNase digestion. DNase I specifically digests DNA into small oligonucleotides, leaving RNA intact.

Perhaps the most important and certainly the most often used technique in RNA analysis is gel electrophoresis. This technique is generally applicable for RNA detection, quantification, purification by size, and quality assessment. Because RNAs are negatively charged, they migrate toward the anode in the presence of electric current. The gel acts as a sieve to selectively impede the migration of the RNA in proportion to its mass, given that its mass is generally proportional to its charge. Because mass is approximately related to chain length, the length of an RNA is more generally determined by its migration. In addition, topology (i.e., circularity) can affect migration, making RNAs appear longer on the gel than they actually are. Gels are used in a wide variety of techniques, including Northern blotting, primer extension, footprinting, and analyzing processing reactions. They are invaluable as preparative and fractionating tools. There are two common types of gel: polyacrylamide and agarose. For most applications, denaturing acrylamide gels are most appropriate. These gels are extremely versatile and can resolve RNAs from ~600 to/=20 nucleotides (nt). In certain circumstances, e.g., resolving different conformers of RNAs or RNA-protein complexes, native gels are appropriate. The only disadvantage to acrylamide gels is that they are not suitable for analyzing large RNAs (or =600 nt); for such applications, agarose gels are preferred. This protocol describes how to prepare, load, and run polyacrylamide gels for RNA analysis.

INTRODUCTIONRecovery of deproteinized RNA almost always is performed via precipitation with ethanol. If the RNA is free of contaminants (i.e., it is well extracted) as determined by spectrophotometry, it will resuspend easily and be ready for any subsequent manipulation. Ethanol precipitation is quite useful because it provides quantitative recovery of any-sized RNA, from several kilobases to 20 nucleotides (nt). Major considerations in this protocol are RNA concentration, pellet identification, and resuspension.

Reverse transcription coupled to the polymerase chain reaction (RT–PCR) is commonly used to detect the presence of mRNAs, pre-mRNAs, or other types of RNA such as noncoding RNAs. The method involves using a primer annealed to the RNA of interest. For mRNA, the primer is usually a synthetic oligo(dT)15–18, a random hexamer mixture (dN)6, or a synthetic DNA oligonucleotide that is complementary to a specific transcript (a gene-specific primer). This DNA:RNA hybrid serves as a template during reverse transcription, in which the enzyme reverse transcriptase (RT) generates a single-stranded cDNA copy of a portion of the target RNA molecule. Using random hexamer priming, it is possible to obtain representative cDNA copies of sequences from the entire length of the mRNAs and pre-mRNAs in a population. This cDNA can then be used as a template for PCR. On addition of gene-specific primers, a specific DNA fragment corresponding to a portion of the RNA of interest is generated.

The electrophoretic mobility shift assay (EMSA), or gel mobility shift assay, is a popular and powerful technique for the detection of RNA–protein interactions. It relies on the fact that naked RNA has certain mobility on nondenaturing gels, but if the RNA is bound by protein, the mobility of the RNA is reduced. Therefore, the binding of protein results in a characteristic upward shift of the RNA on a gel, as monitored using radiolabeled RNA. For reasons that are not completely understood, most RNA–protein complexes—particularly those that result from high-affinity interactions—do not dissociate during the prolonged times required for electrophoretic separation. Because high-affinity interactions are more stable, it is often possible to identify specific interactions over a “background” of weak interactions. Accordingly, EMSAs can be performed using complex mixtures of proteins such as cell extracts. They can be used to investigate a wide range of RNA–protein interactions—from single protein-binding events to assembly of large complexes such as the spliceosome. EMSAs can also be useful for determining kinetic parameters (such as affinity constants) for RNA–protein interactions.

Here we describe how to prepare nucleoplasmic extracts from suspension cultures of Drosophila S2 (Schneider Line 2) cells. Harvested cells are washed in phosphate-buffered saline supplemented with MgCl2, resuspended in hypotonic buffer, and homogenized. Nuclei are isolated by centrifugation and then sonicated. The nuclear sonicate is placed on a 30% sucrose cushion and sedimented. The soluble nuclear ribonucleoprotein (RNP) complexes remain in the supernatant and the nuclear membrane fragments and chromatin pellet through the sucrose cushion.

RNA is photoreactive on exposure to ultraviolet (UV) light in the 250- to 270-nm range. On UV treatment, RNA bases absorb energy to generate free radicals that can covalently attach to nearby amino acid residues in RNA-bound proteins. UV cross-linking experiments have been extensively used to identify and characterize RNA-binding proteins. The method described here involves the use of 32P-labeled RNA and crude extracts or purified proteins.

For large-scale transcription reactions or for cost savings, a laboratory may want to prepare its own recombinant T7-, SP6-, or T3-phage RNA polymerases. It is convenient to perform this preparation every 2–3 years and have a consistent and reliable source of phage RNA polymerase for many in vitro transcription reactions. In the protocol presented here, the recombinant plasmid expressing T7 RNA polymerase (RNAP) as a his6-tagged molecule is under an isopropyl β-d-1-thio-galactopyranoside (IPTG)-inducible promoter. The bacteria are lysed by sonication, the his6-tagged protein in the bacterial lysate is purified by binding to Ni-NTA agarose, and the resin is then extensively washed and eluted with imidazole. The purified enzyme is dialyzed against a glycerol-containing storage buffer and can then be stored for months or years at −20°C.

All in vitro RNA transcription reactions involve the use of common components: a template, phage RNA polymerase, and ribonucleoside triphosphates (rNTPs). The concentration of rNTPs is a crucial variable that is manipulated to determine the specific activities of labeled RNAs. This protocol describes methods used to synthesize RNAs of low specific activity (trace labeled), medium specific activity, and high specific activity. Also described is how the same principles that determine the level of incorporation of labeled rNTPs are applicable to the synthesis of transcripts containing modified nucleotides, including 5′ cap and internal (body) modifications such as biotinylated uridine, 4-thiouridine, and phosphorothiotated nucleotides. Careful attention to detail in setting up the transcription reaction will permit the synthesis of any number of “designer RNAs.”

INTRODUCTIONWith the proliferation of researchers investigating various aspects of RNA biology has come a corresponding proliferation of commercially available kits for RNA purification. Some of these kits have specialized uses and applications, and it is not always possible to simply mix and match the different protocols. Although the instructions are often written with a minimum of specifics concerning the chemical basis for the separation characteristics that the kits use, the competent researcher will want to understand these important features. Unfortunately, commercial sources work hard to hide the specifics of their methods. Fortunately, patent documents are publicly available, and the government mandates material safety data disclosures of the composition of at least those components that represent a threat to environmental health and safety. Thus, it is possible to infer many aspects of such “proprietary” materials, which in most cases are only slightly and unimportantly modified from published techniques. We do not describe in detail the specific steps involved in the use of particular kits; they are detailed in the literature that comes with each kit and their components (e.g., proprietary buffers). Instead, we detail some advantages and disadvantages of using RNA purification kits.

INTRODUCTIONThis protocol describes a method for enrichment of poly(A)+ messenger RNA (mRNA) using immobilized oligo(dT). It takes advantage of the fact that the vast majority of eukaryotic mRNAs contain a 3′ tail of polyadenylic acid added post-transcriptionally. It relies on the ability of these tails to form stable hybrids with oligo(dT) under high-salt conditions. The hybrids are destabilized when salt is absent, allowing mRNAs to be recovered. This protocol is effective in removing the bulk of non-mRNA RNAs. A drawback of the technique is that it does not enrich for mRNAs that lack poly(A) tails (e.g., normal histone mRNAs or deadenylated mRNAs whose poly[A] tails have been removed during mRNA turnover). Several commercially available kits can be used for selection of poly(A)+ RNAs. However, they are expensive and do not perform any better than “homemade” reagents using (dT) cellulose. One obvious advantage of this method is that mRNAs are enriched ~50-fold. Accordingly, detection by Northern blotting or reverse transcriptase polymerase chain reaction (RT-PCR) is considerably more sensitive.

INTRODUCTIONTRIzol solubilization and extraction is a relatively recently developed general method for deproteinizing RNA. This method is particularly advantageous in situations where cells or tissues are enriched for endogenous RNases or when separation of cytoplasmic RNA from nuclear RNA is impractical. TRIzol (or TRI Reagent) is a monophasic solution of phenol and guanidinium isothiocyanate that simultaneously solubilizes biological material and denatures protein. After solubilization, the addition of chloroform causes phase separation (much like extraction with phenol:chloroform:isoamyl alcohol), where protein is extracted to the organic phase, DNA resolves at the interface, and RNA remains in the aqueous phase. Therefore, RNA, DNA, and protein can be purified from a single sample (hence, the name TRIzol). TRIzol extraction is also an effective method for isolating small RNAs, such as microRNAs, piwi-associated RNAs, or endogeneous, small interfering RNAs. However, TRIzol is expensive and RNA pellets can be difficult to resuspend. Thus, the use of TRIzol is not recommend when regular phenol extraction is practical.

INTRODUCTIONPerhaps the most important and certainly the most often used technique in RNA analysis is gel electrophoresis. Because RNAs are negatively charged, they migrate toward the anode in the presence of electric current. The gel acts as a sieve to selectively impede the migration of the RNA in proportion to its mass, given that its mass is generally proportional to its charge. Because mass is approximately related to chain length, the length of an RNA is more generally determined by its migration. In addition, topology (i.e., circularity) can affect migration, making RNAs appear longer on the gel than they actually are. There are two common types of gel: polyacrylamide and agarose. For most applications involving RNAs of ≤600 nucleotides, denaturing acrylamide gels are most appropriate. In contrast, agarose gels are generally used to analyze RNAs of ≥600 nucleotides, and are especially useful for analysis of mRNAs (e.g., by Northern blotting). RNA analysis on agarose gels is essentially identical to DNA analysis (except that the gel boxes used must be dedicated to RNA work or to other ribonuclease-free work). Here we describe the use of straightforward Tris borate, EDTA (TBE) gels for routine analysis. These gels are appropriate for determining the quantity and integrity of RNA before using it for other applications. This procedure should not be used to determine size with accuracy, because the RNA will not remain in its extended state throughout the run.

INTRODUCTIONThis protocol describes a method for RNA purification by sodium dodecyl sulfate (SDS) solubilization and phenol extraction. It is of wide utility and is used routinely to deproteinize RNAs in biological material that has been solubilized in SDS, an ionic detergent that dissolves membranes, disrupts protein-nucleic acid interactions, and inactivates ribonucleases. Once solubilized, addition of phenol or phenol:chloroform:isoamyl alcohol (PCA) completely denatures the protein, and it becomes insoluble in aqueous solution. PCA extraction is the method of choice for preparing cytoplasmic RNA from tissue culture cells or in any other situation (e.g., enzyme reactions) where solubilization in SDS is easily achievable.

INTRODUCTIONPerhaps the most important and certainly the most often used technique in RNA analysis is gel electrophoresis. This technique is generally applicable for RNA detection, quantification, purification by size, and quality assessment. Because RNAs are negatively charged, they migrate toward the anode in the presence of electric current. The gel acts as a sieve to selectively impede the migration of the RNA in proportion to its mass, given that its mass is generally proportional to its charge. Because mass is approximately related to chain length, the length of an RNA is more generally determined by its migration. In addition, topology (i.e., circularity) can affect migration, making RNAs appear longer on the gel than they actually are. Gels are used in a wide variety of techniques, including Northern blotting, primer extension, footprinting, and analyzing processing reactions. They are invaluable as preparative and fractionating tools. There are two common types of gel: polyacrylamide and agarose. For most applications, denaturing acrylamide gels are most appropriate. These gels are extremely versatile and can resolve RNAs from ~600 to ≤20 nucleotides (nt). In certain circumstances, e.g., resolving different conformers of RNAs or RNA-protein complexes, native gels are appropriate. The only disadvantage to acrylamide gels is that they are not suitable for analyzing large RNAs (≥600 nt); for such applications, agarose gels are preferred. This protocol describes how to prepare, load, and run polyacrylamide gels for RNA analysis.

INTRODUCTIONIt often is desirable to “prefractionate” RNA before analysis. Ordinarily, this can only be done with tissue culture cells, although it is possible to isolate nuclei and cytoplasm from certain “soft” tissues such as liver and white blood cells. This protocol describes a method for separating nuclei from the cytoplasm that can be used for many tissue culture types. This procedure also is useful for cells grown in suspension or for adherent cells. The procedure relies on swelling in hypotonic buffer, subsequent gentle homogenization, and centrifugation. This method is not appropriate for material (e.g., bacteria, yeast) that has high intrinsic RNase activity, or tissues that are difficult to solubilize, such as muscle tissue or plant material.