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The recent development of prime editing (PE) genome engineering technologies has the potential to significantly simplify the generation of human pluripotent stem cell (hPSC)-based disease models. PE is a multicomponent editing system that uses a Cas9-nickase fused to a reverse transcriptase (nCas9-RT) and an extended PE guide RNA (pegRNA). Once reverse transcribed, the pegRNA extension functions as a repair template to introduce precise designer mutations at the target site. Here, we systematically compared the editing efficiencies of PE to conventional gene editing methods in hPSCs. This analysis revealed that PE is overall more efficient and precise than homology-directed repair of site-specific nuclease-induced double-strand breaks. Specifically, PE is more effective in generating heterozygous editing events to create autosomal dominant disease-associated mutations. By stably integrating the nCas9-RT into hPSCs we achieved editing efficiencies equal to those reported for cancer cells, suggesting that the expression of the PE components, rather than cell-intrinsic features, limit PE in hPSCs. To improve the efficiency of PE in hPSCs, we optimized the delivery modalities for the PE components. Delivery of the nCas9-RT as mRNA combined with synthetically generated, chemically-modified pegRNAs and nicking guide RNAs improved editing efficiencies up to 13-fold compared with transfecting the PE components as plasmids or ribonucleoprotein particles. Finally, we demonstrated that this mRNA-based delivery approach can be used repeatedly to yield editing efficiencies exceeding 60% and to correct or introduce familial mutations causing Parkinson's disease in hPSCs.From muscles to nerves, our body is formed of many kinds of cells which can each respond slightly differently to the same harmful genetic changes. Understanding the exact relationship between mutations and cell-type specific function is essential to better grasp how conditions such as Parkinson’s disease or amyotrophic lateral sclerosis progress and can be treated. Stem cells could be an important tool in that effort, as they can be directed to mature into many cell types in the laboratory. Yet it remains difficult to precisely introduce disease-relevant mutations in these cells. To remove this obstacle, Li et al. focused on prime editing, a cutting-edge ‘search and replace’ approach which can introduce new genetic information into a specific DNA sequence. However, it was unclear whether this technique could be used to efficiently create stem cell models of human diseases. A first set of experiments showed that prime editing is superior to conventional approaches when generating mutated genes in stem cells. Li et al. then further improved the efficiency and precision of the method by tweaking how prime editing components are delivered into the cells. The refined approach could be harnessed to quickly generate large numbers of stem cells carrying mutations associated with Parkinson’s disease; crucially, prime editing could then also be used to revert a mutated gene back to its healthy form. The improved prime editing approach developed by Li et al. removes a major hurdle for scientists hoping to use stem cells to study genetic diseases. This could potentially help to unlock progress in how we understand and ultimately treat these conditions.

Peer reviewed: True, P elements were first discovered in the fruit fly Drosophila melanogaster as the causative agents of a syndrome of aberrant genetic traits called hybrid dysgenesis. This occurs when P element-carrying -males mate with females that lack P elements and results in progeny displaying sterility, mutations and chromosomal rearrangements. Since then numerous genetic, developmental, biochemical and structural studies have culminated in a deep understanding of P element transposition: from the cellular regulation and repression of transposition to the mechanistic details of the transposase nucleoprotein complex. Recent studies have revealed how piwi-interacting small RNA pathways can act to control splicing of the P element pre-mRNA to modulate transposase production in the germline. A recent cryo-electron microscopy structure of the P element transpososome reveals an unusual DNA architecture at the transposon termini and shows that the bound GTP cofactor functions to position the transposon ends within the transposase active site. Genome sequencing efforts have shown that there are P element transposase-homologous genes (called THAP9) in other animal genomes, including humans). This review highlights recent and previous studies, which together have led to new insights, and surveys our current understanding of the biology, biochemistry, mechanism and regulation of P element transposition.

Although alternative pre-mRNA splicing (AS) significantly diversifies the neuronal proteome, the extent of AS is still unknown due in part to the large number of diverse cell types in the brain. To address this complexity issue, we used an annotation-free computational method to analyze and compare the AS profiles between small specific groups of Drosophila circadian neurons. The method, the Junction Usage Model (JUM), allows the comprehensive profiling of both known and novel AS events from specific RNA-seq libraries. The results show that many diverse and novel pre-mRNA isoforms are preferentially expressed in one class of clock neuron and also absent from the more standard Drosophila head RNA preparation. These AS events are enriched in potassium channels important for neuronal firing, and there are also cycling isoforms with no detectable underlying transcriptional oscillations. The results suggest massive AS regulation in the brain that is also likely important for circadian regulation., eLife digest The life of nearly all creatures on Earth follows the rhythm of day and night. For example, in fruit flies, darkness and light dictate when the insects feed, rest, move or mate. This is possible thanks to the circadian clock, an internal program which is synchronized with the environment to tell cells in the body when to perform certain roles. In fruit flies, the structure that keeps the body clock ticking is formed of about 150 ‘circadian neurons’, which are divided into several subgroups. In these cells, a complex genetic programis at work, with networks of genes being ‘switched on’ in a cyclical way. To understand how this program works, scientists need to know which genes are turned on and when, as well as which proteins are created based on the information contained in these genes. This can be difficult because one gene does not necessarily code for only one protein. Indeed, when a gene is turned on, it gets copied into a pre-messenger RNA (pre-mRNA), which is formed of several modules. The pre-mRNA can then go through a process called alternative splicing that shuffles or removes the different modules. This means that one gene can give rise to different pre-mRNA molecules that will each serve as a template to build a distinct protein. Until now, there have been few studies that examine the different types of pre-mRNAs found in circadian neurons, and how these change with the time of day. Here, Wang, Abruzzi et al. extract three subgroups of circadian neurons, and one subgroup of non-circadian neurons, from the brain of fruit flies. The pre-mRNAs are isolated, and then a new computational method, called JUM, identifies, counts and categorizes the pre-mRNA molecules in the different groups of neurons. This analysis reveals hundreds of previously unknown pre-mRNA molecules, many of which differed extensively between the types of brain cells. When comparing circadian and non-circadian neurons, Wang, Abruzzi et al. show that the circadian cells had more pre-mRNAs that code for proteins that help the cell communicate with other neurons. Finally, many genes in the circadian neurons use alternative splicing to turn on different types of pre-mRNA molecules at different times of the day in a cyclical way; this suggests that these pre-mRNAs might be participating in the genetic circadian program. Many human disorders, such as certain forms of insomnia, emerge when the circadian clock is thrown off balance. The results reported by Wang, Abruzzi et al. show that alternative splicing may be an overlooked mechanism that shapes this complex program.

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.

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.

We have developed an in vitro reaction system for Drosophila P element transposition. Transposition products were recovered by selection in E. coli, and contained simple P element insertions flanked by 8 bp target site duplications as observed in vivo. Transposition required Mg +2 and partially purified P element transposase. Unlike other DNA rearrangement reactions, P element transposition in vitro used GTP as a cofactor; deoxyGTP, dideoxyGTP, or the nonhydrolyzable GTP analogs GMP-PNP or GMP-PCP were also used. Transposon DNA molecules cleaved at the P element termini were able to transpose, but those lacking 3′-hydroxyl groups were inactive. These biochemical data are consistent with genetic data suggesting that P element transposition occurs via a "cut-and-paste" mechanism.

Structure of RNA Prediction and experimental investigation of RNA secondary and tertiary foldings Aminoacyl-tRNA synthetase-tRNA recognition RNA-protein interactions in ribosomes RNA-protein interactions in RNase P Structure and function of hnRNP proteins RNA-protein interactions in the splicing snRNPs Interaction of 5S with TFIIIA Control of human immunodeficiency virus gene expression by the RNA-binding proteins tat and rev Using peptides to study RNA-protein recognition Study of RNA-protein recognition by in vitro selection.

Drosophila P transposable elements encode two proteins, an 87 kd transposase protein and a 66 kd protein that has been hypothesized to repress transposition. We have made germline transformants carrying modified P element derivatives that encode only the 66 kd protein and shown that these elements repress transposase activity in both the germline and the soma. The position of these elements in the genome quantitatively affected their ability to negatively regulate transposase and to express the 66 kd protein. Single 66 kd element-containing strains did not exhibit the maternal inheritance of P cytotype characteristic of P strains. However, we demonstrated that a true P strain produced higher levels of the 66 kd protein during oogenesis than single 66 kd P elements. Thus, the expression of the 66 kd repressor during oogenesis may be a major determinant of the maternal effect of P cytotype.

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.

P elements are a family of mobile DNA elements found in Drosophila . P‐element transposition is tightly regulated, and P‐element‐encoded repressor proteins are responsible for inhibiting transposition in vivo . To investigate the molecular mechanisms by which one of these repressors, the KP protein, inhibits transposition, a variety of mutant KP proteins were prepared and tested for their biochemical activities. The repressor activities of the wild‐type and mutant KP proteins were tested in vitro using several different assays for P‐element transposase activity. These studies indicate that the site‐specific DNA‐binding activity of the KP protein is essential for repressing transposase activity. The DNA‐binding domain of the KP repressor protein is also shared with the transposase protein and resides in the N‐terminal 88 amino acids. Within this region, there is a C 2 HC putative metal‐binding motif that is required for site‐specific DNA binding. In vitro the KP protein inhibits transposition by competing with the transposase enzyme for DNA‐binding sites near the P‐element termini.

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.

Sex determination and X chromosome dosage compensation in Drosophila melanogaster are directed by the Sex-lethal (Sxl) protein. In part, Sxl functions by regulating the splicing of the transformer pre-mRNA by binding to a 3' splice site polypyrimidine tract. Polypyrimidine tracts are essential for splicing of metazoan pre-mRNAs. To unravel the mechanism of splicing regulation at polypyrimidine tracts we analyzed the interaction of Sxl with RNA. The RNA binding activity of Sxl was mapped to the two ribonucleoprotein consensus sequence domains of the protein. Quantitation of binding showed that both RNA binding domains (RBDs) were required in cis for site-specific RNA binding. Individual RBDs interacted with RNA more weakly and had lost the ability to discriminate between wild-type and mutant transformer polypyrimidine tracts. Structural elements in one of the RBDs that are likely to interact with a polypyrimidine tract were identified using nuclear magnetic resonance techniques. In addition, our data suggest that multiple imino protons of the transformer polypyrimidine tract were involved in hydrogen bonding. Interestingly, in vitro Sxl bound with equal affinity to polypyrimidine tracts of pre-mRNAs that it does not regulate in vivo. We discuss the implications of this finding for the mechanism through which Sxl may gain selectivity for particular polypyrimidine tracts in vivo.

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.”

An intron is an extended genomic feature whose function requires multiple constrained positions—donor and acceptor splice sites, a branch point, a polypyrimidine tract and suitable splicing enhancers—that may be distributed over hundreds or thousands of nucleotides. New introns are therefore unlikely to emerge by incremental accumulation of functional sub-elements. Here we demonstrate that a functional intron can be created de novo in a single step by a segmental genomic duplication. This experiment recapitulates in vivo the birth of an intron that arose in the ancestral jawed vertebrate lineage nearly half-a-billion years ago., The appearance of a new intron that splits an exon without disrupting the corresponding peptide sequence is a rare event in vertebrate genomes. Hellsten et al. demonstrate that, under certain circumstances, a functional intron can be produced in a single step by segmental genomic duplication.

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.

We analyzed the DNA binding properties of wild-type simian virus 40 large T antigen and found that the protein binds cooperatively to three tandem sites at a regulatory region of SV40 DNA. One consequence of this T antigen:DNA interaction is the specific repression of SV40 early RNA synthesis in vitro. We mapped a region of 85 base pairs that is necessary and sufficient to initiate early SV40 transcription in vitro. This promoter region lies directly adjacent to the third T antigen binding site but does not include the "TATA" sequence. To determine how T antigen interacts with its binding sites to repress RNA synthesis, we analyzed transcription directed by a variety of wild-type, mutant and hybrid template DNAs. Our findings suggest that the cooperative binding of T antigen to its sites is directly responsible for inhibiting the initiation, rather than blocking elongation, of early RNA synthesis. A model is presented to explain the role of T antigen binding in the regulation of viral transcription and DNA replication during SV40 lytic infection.

Deletion and substitution mutations in the control region of simian virus 40 (SV40) were used to study regulation of early transcription by T antigen. A mutant, pIN4, containing substitutions within T antigen binding site II, is transcribed in a cell-free extract at wild-type efficiency but is unable to be repressed in vitro by purified T antigen under conditions that fully repress wild-type transcription. These results suggest a functional role for T antigen binding site II in the repression of early SV40 transcription. To investigate autoregulation in vivo, transcription from the SV40 early promoter was quantitated in COS7 monkey cells transfected with plasmid vectors carrying the mouse dihydrofolate reductase gene (SV-dhfr vectors). Mutant SV-dhfr vectors lacking T antigen site I or site II sequences overproduce dhfr RNA from the SV40 early promoter three to four fold, whereas deletion of both sites I and II or the presence of a temperature-sensitive T antigen (tsA209) results in an eight to ten fold increase in dhfr RNA. Our results indicate that binding of T antigen to both sites I and II plays a role in autoregulation.

Drosophila P transposable elements encode an 87 kd trans-acting protein, transposase, that is required to catalyze P element transposition and excision. We show here that purified transposase is a site-specific DNA binding protein. P element transposase does not interact with the terminal 31 bp inverted repeats but instead interacts specifically with an internal 10 bp consensus sequence present at both the 5' and 3' ends of P element DNA. These binding sites lie within sequences known to be important for transposition in vivo. Transposase also displays an unusually high nonspecific affinity for DNA. The transposase binding site at the 5' and overlaps sequences we show to be essential for transcription from the P element promoter in vitro, which raises the possibility that either transposase or the related 66 kd P element protein may affect P element transcription. From these and other observations, we suggest that the P element transposition reaction probably requires the binding of additional Drosophila protein factors to the terminal DNA sequences.

We have identified proteins encoded by P transposable elements expressed in transformed Drosophila tissue culture cells. Two proteins have been identified by immunochemical techniques. One, an 87,000 dalton polypeptide, is encoded by a P element mRNA lacking the third (ORF2-ORF3) intervening sequence. The other protein, a 66,000 dalton polypeptide, is encoded by an mRNA that retains the third intron and is found in somatic tissues. Furthermore, tissue culture cell lines expressing the 87,000 dalton polypeptide are able to catalyze both the precise and imprecise excision of a nonautonomous P element. The 87,000 dalton protein is encoded by sequences from all four P element open reading frames. Taken together, these data strongly suggest that the 87,000 dalton polypeptide is the P element transposase.

We show that the germline specificity of P element transposition is controlled at the level of mRNA splicing and not at the level of transcription. In the major P element RNA transcript, isolated from somatic cells, the first three open reading frames are joined by the removal of two introns. Using in vitro mutagenesis and genetic analysis we demonstrate the existence of a third intron whose removal is required for transposase production. We propose that this intron is only removed in the germline and that its removal is the sole basis for the germline restriction of P element transposition.