Kaposi's Sarcoma-Associated Herpesvirus Utilizes and Manipulates RNA N6-Adenosine Methylation To Promote Lytic Replication.

N6-adenosine methylation (m6A) is the most common posttranscriptional RNA modification in mammalian cells. We found that most transcripts encoded by the Kaposi's sarcoma-associated herpesvirus (KSHV) genome undergo m6A modification. The levels of m6A-modified mRNAs increased substantially upon stimulation for lytic replication. The blockage of m6A inhibited splicing of the pre-mRNA encoding the replication transcription activator (RTA), a key KSHV lytic switch protein, and halted viral lytic replication. We identified several m6A sites in RTA pre-mRNA crucial for splicing through interactions with YTH domain containing 1 (YTHDC1), an m6A nuclear reader protein, in conjunction with serine/arginine-rich splicing factor 3 (SRSF3) and SRSF10. Interestingly, RTA induced m6A and enhanced its own pre-mRNA splicing. Our results not only demonstrate an essential role of m6A in regulating RTA pre-mRNA splicing but also suggest that KSHV has evolved a mechanism to manipulate the host m6A machinery to its advantage in promoting lytic replication.IMPORTANCE KSHV productive lytic replication plays a pivotal role in the initiation and progression of Kaposi's sarcoma tumors. Previous studies suggested that the KSHV switch from latency to lytic replication is primarily controlled at the chromatin level through histone and DNA modifications. The present work reports for the first time that KSHV genome-encoded mRNAs undergo m6A modification, which represents a new mechanism at the posttranscriptional level in the control of viral replication.

Cotranslational microRNA mediated messenger RNA destabilization.

MicroRNAs are small (22 nucleotide) regulatory molecules that play important roles in a wide variety of biological processes. These RNAs, which bind to targeted mRNAs via limited base pairing interactions, act to reduce protein production from those mRNAs. Considerable evidence indicates that miRNAs destabilize targeted mRNAs by recruiting enzymes that function in normal mRNA decay and mRNA degradation is widely thought to occur when mRNAs are in a ribosome free state. Nevertheless, when examined, miRNA targeted mRNAs are invariably found to be polysome associated; observations that appear to be at face value incompatible with a simple decay model. Here, we provide evidence that turnover of miRNA-targeted mRNAs occurs while they are being translated. Cotranslational mRNA degradation is initiated by decapping and proceeds 5' to 3' behind the last translating ribosome. These results provide an explanation for a long standing mystery in the miRNA field.

Preparing Cellular DNA from Nuclei or Whole Cells.

It is often desirable to have cellular DNA on hand. DNA is stable and can be maintained intact for many years. This protocol describes the preparation of DNA from nuclei after the cytoplasmic extract has been removed. The resulting DNA is suitable for polymerase chain reactions and Southern blots to determine copy number and sites of integration of plasmids in stable cell lines. Quantitation of DNA may not be exact because RNA is not completely removed. The method can also be used on whole cells, but there will be more RNA contamination.

Removal of rRNA from Deproteinized, Phenol-Extracted Total RNA by Hybrid Selection.

In this protocol, rRNAs are selectively removed from a total RNA sample by hybridizing the rRNAs to complementary biotinylated oligodeoxynucleotides that can be affinity-purified using streptavidin beads, leaving all other RNAs behind. Although commercially available kits can be used to perform this procedure, they are expensive. We recommend that investigators order species-specific oligodeoxynucleotides for their own applications. There are well-established secondary structure predictions for all rRNAs.

Boundary Analysis to Determine the Minimal RNA Sequence Required for Protein Binding.

Boundary analysis is a powerful and often overlooked method for determining the sequences within an RNA molecule that are required for a specific RNA-protein interaction. In this approach, 5'- and 3'-end-labeled RNAs are fragmented randomly (usually by limited alkaline hydrolysis, but nuclease fragmentation can also be used) such that a ladder of fragments covering the whole molecule of interest is produced. This mixture of fragments is then allowed to bind to the protein (or other molecule) of interest. Bound fragments are selected by affinity or antibody binding and then displayed on a gel. The point at which banding is lost for 5'-end-labeled RNAs defines the 3' boundary of the binding site, and the point at which banding is lost for 3'-end-labeled RNAs defines the 5' boundary of the binding site.

Chemical modification interference.

Chemical modification interference is a powerful method for surveying an entire RNA molecule to identify functionally important chemical groups. The basic idea is to generate a pool of end-labeled RNAs wherein each RNA molecule is chemically modified (e.g., by diethyl pyrocarbonate [DEPC], hydrazine, dimethyl sulfate, CMCT, or kethoxal) at a different position. The pool of RNAs is then allowed to participate in the reaction of interest. The functionally important RNA molecules (e.g., those bound by protein or that successfully participate in a processing reaction) are then separated from the nonfunctional RNA molecules (e.g., those not bound by protein or unable to participate in a processing reaction). This is often achieved by straightforward gel electrophoretic analysis. In the case of protein binding, it is necessary to be able to separate bound RNA from unbound RNA, which can be accomplished using electrophoretic mobility shift assays, filter binding, or affinity approaches (e.g., by immunoprecipitation or the use of tagged proteins). None of these techniques requires that a large fraction of RNA be bound or reacted, and, as a result, they are quite sensitive. Here we describe one example of a chemical modification interference assay in which RNA is modified with DEPC or hydrazine before binding to a protein. This technique can be readily adapted for use with other chemicals.

Nucleotide analog interference mapping.

Methods collectively known as modification interference are exceptionally powerful approaches used to identify functionally important chemical groups in the phosphodiester backbone or nucleobases of an RNA. In a modification interference assay, end-labeled RNAs that have been modified at different positions are allowed to participate in a reaction of interest, and then functional RNA molecules (e.g., those bound by protein or that successfully participate in a processing reaction) are separated from nonfunctional RNA molecules (e.g., those not bound by protein or unable to participate in a processing reaction). Nucleotide analog interference mapping (NAIM) involves the incorporation of α-thionucleotides containing a modified base into the RNA molecule of interest. The sites containing the modified base are identified by cleavage with iodoethanol. NAIM is useful whenever the thiophosphate substitution on its own does not prevent or inhibit a specific reaction. To perform NAIM, it is first necessary to perform a thiophosphate interference analysis. Any positions that are not affected by thiophosphate substitution can then be analyzed by NAIM.

Measuring the length of poly(A) tails.

Adenylation status has an important role in the regulation of mRNA metabolism: mRNAs are deadenylated before degradation, microRNAs (miRNAs) can cause deadenylation, and the poly(A) length of certain mRNAs is regulated during development. This protocol describes methods that can be used to measure the poly(A) tail length of specific mRNAs. These include, in the order of increasing sensitivity, (1) northern blotting of intact and experimentally deadenylated mRNAs and (2) northern blotting of intact and experimentally deadenylated mRNA fragments that have been cleaved near the 3' end with RNase H. Highly sensitive polymerase chain reaction (PCR)-based approaches are also discussed.

An alternative method for processing northern blots after capillary transfer.

Different laboratories use different methods for the prehybridization, hybridization, and washing steps of the northern blotting procedure. In this protocol, a northern blot is pretreated with Church and Gilbert hybridization buffer to block nonspecific probe-binding sites. The immobilized RNA is then hybridized to a DNA probe specific for the RNA of interest. Finally, the membrane is washed and subjected to autoradiography or phosphorimaging. The solutions and conditions described here may be ideal for those who prefer to use fewer ingredients in their solutions. This protocol is designed to achieve the same goals as other northern blotting approaches. It minimizes background (nonspecific adherence of probe to membrane and nonspecific hybridization) and maximizes specific hybridization to RNAs immobilized on a membrane.

An intimate view of a spliceosome component.

A high-resolution structure reveals how the ribonucleoprotein particle called U1 snRNP engages with 5' splice sites.

Direct chemical sequencing of end-labeled RNA.

Chemical sequencing of RNA relies on the fact that each of the four bases in RNA is susceptible to chemical modification in a different way. In this protocol, end-labeled RNAs are subjected to base-specific chemical modification reactions that make the RNA strand adjacent to the modified base susceptible to cleavage. The chemical modification reaction is base-specific but limited so that not every base in every strand is modified. After cleavage, the resulting set of radioactive fragments is resolved via polyacrylamide gel electrophoresis.

Poisoned primer extension.

Poisoned primer extension is primarily used to distinguish between RNAs that are nearly identical in sequence but cannot be distinguished by standard primer extension because they are the same size (e.g., edited vs. nonedited transcripts). It is conceptually identical to the standard primer extension reaction but involves the use of a chain-terminating dideoxynucleotide (the "poison") in the presence of the other three nucleotides. A radioactively labeled primer that hybridizes a short-distance downstream from the "changed" region of interest is extended by reverse transcription into this region of sequence variation. The reactions contain three of the four substrates for extension (e.g., dATP, dGTP, and dTTP) and a chain-terminating dideoxynucleotide (e.g., ddCTP). The extension reaction stops when reverse transcriptase adds a chain-terminating dideoxynucleotide to the template (e.g., it will add ddCTP when it encounters a G in the template sequence). RNAs that differ in sequence at that position will yield different-sized extension products that can be resolved on a denaturing gel.

Mapping RNA-protein interactions using hydroxyl-radical footprinting.

The binding of a protein to an RNA sequence protects the region of the RNA from cleavage by chemicals or RNases; this protected region is known as the protein's "footprint." In the footprinting protocol presented here, end-labeled RNAs with and without bound protein are cleaved using chemical methods. Fe(II)-EDTA is used to generate hydroxyl radicals in the presence of a reducing agent. These hydroxyl radicals indiscriminately cleave ribose groups in regions of the ribose-phosphate backbone that are exposed to solvent. After termination of cleavage, the resulting RNA fragments are analyzed by gel electrophoresis on denaturing polyacrylamide gels. Because hydroxyl radicals are smaller and cleave less specifically than RNases, this approach, if feasible, is often the method of choice for monitoring sites of RNA-protein interactions.

Mapping RNA-protein interactions using iodine footprinting.

Footprinting methods are used to determine the binding site of a protein on an RNA. They are based on the fact that a protein bound to an RNA protects the RNA from cleavage by chemicals or nucleases. The footprinting method described here relies on the ability of iodoethanol to cleave the backbone of RNA when a phosphodiester bond contains sulfur in place of a nonbridging oxygen. A potential advantage of using iodoethanol for cleavage is that one can prepare RNAs that contain selective thiol substitutions such that the resulting cleavage patterns contain fewer bands, making quantification "easier" and the results cleaner. For example, a population of RNAs that only contains nonbridging thiol substitutions 5' to each adenine can be prepared by including αS ATP in the transcription reaction. In this protocol, all positions on an RNA are surveyed. First, a series of RNAs is synthesized by transcription in the presence of αS ATP, αS CTP, αS UTP, or αS GTP. Each of the selectively substituted RNAs is probed with the binding protein of interest. The portion of the RNA that is not bound by protein is accessible and vulnerable to cleavage by iodoethanol. Finally, the cleavage products are analyzed by gel electrophoresis on denaturing polyacrylamide gels.

Detecting RNA-RNA interactions using psoralen derivatives.

Psoralens are tricyclic compounds that intercalate into double-stranded DNA or RNA and, on irradiation with long-wavelength (365-nm) UV light, covalently link pyrimidines on adjacent strands. More rarely, psoralen cross-links can be observed at the ends of helices (i.e., double-stranded-single-stranded boundaries). Although psoralens can, in some instances, cross-link protein to RNA, their primary application is to detect RNA-RNA base-pairing interactions. The most useful psoralen derivative is 4'-aminomethyl trioxsalen (AMT), which is soluble in H2O. This protocol describes the use of AMT to detect RNA-RNA interactions in tissue culture cells or in extracts. Cross-linked RNAs are detectable by their reduced mobility in polyacrylamide gels. Cross-links can be reversed by exposure to short-wavelength (254 nm) UV light.

Preparation of cross-linked cellular extracts with formaldehyde.

Formaldehyde is a powerful cross-linking agent that elicits protein-protein and protein-nucleic acid cross-links. This protocol describes the formaldehyde cross-linking of intact cells followed by either the preparation of a whole-cell extract by sonication or the preparation of nuclear and cytoplasmic extracts by fractionation. Cross-linked extracts are treated with high salt or sodium dodecyl sulfate (SDS) to disrupt non-cross-linked aggregates. Protein-protein and protein-nucleic acid interactions can then be studied by purifying the components of interest from the extracts (e.g., using immunoprecipitation). Formaldehyde cross-links are reversible by heat.

Detection of RNAs by 3'-end labeling and RNase H digestion.

This protocol describes an extremely sensitive procedure for detecting the presence of known or unknown RNAs in a complex mixture. A selectively enriched population of RNAs is subjected to 3'-end labeling with [(32)P]pCp, and labeled products are separated from unincorporated label. The labeled RNAs are hybridized to sequence-specific complementary oligodeoxynucleotides, treated with RNase H (which cleaves RNA in an RNA-DNA hybrid) and the products analyzed by electrophoresis through denaturing polyacrylamide gels with the appropriate controls. If the RNA of interest was present and hybridized to its complementary oligonucleotide, its digestion with RNase H will result in a shift in its mobility through the gel or, if the RNA was fully degraded, its band will not appear. If the RNA of interest is not cleaved in the presence of any known complementary oligodeoxynucleotides, then its position in the gel will remain unaltered. This result may suggest the presence of a new or unknown RNA that may be identified using a variety of cloning techniques or by direct chemical sequencing methods.