Binding of MPA to the enzyme-GMP complex inhibits the interaction with RNA.

<p>(<b>A</b>) The enzyme-GMP complex was incubated with a radiolabeled RNA substrate (3 µM) of 81 nucleotides in the presence of increasing concentrations of MPA. UV-cross-linking assays were performed to monitor the binding of radiolabelled RNA to the enzyme-GMP complex and visualized by SDS-PAGE analysis and autoradiography. (<b>B</b>) The reaction products were quantified by phosphorimaging.</p

MPA inhibits the DNA ligase activity.

<p>(<b>A</b>) MPA inhibits the strand-joining activity of DNA ligase. Ligation reactions (50 mM Tris-HCl pH 7.5, 5 mM DTT, 10 mM MgCl₂, 1 mM ATP, 500 fmol 1-nt gap DNA substrate and 6 pmol of the <i>Chlorella</i> virus DNA ligase) were performed at 22°C for 15 min in the presence of increasing concentrations of MPA. The samples were analyzed by electrophoresis through a 17% polyacrylamide gel containing 7 M urea. An autoradiogram of the gel is shown. The positions of the input 5′-monophosphate 18-mer strand (pDNA) and the 36-mer ligation product are indicated. The radiolabeled ligated product was then quantified by phosphorimaging (<i>right side of the panel</i>). (<b>B</b>) High concentrations of MPA are required to inhibit the first step of the ligase reaction. The formation of the enzyme-AMP covalent intermediate was monitored by incubating the purified enzyme in the presence of [alpha-³²P]ATP and increasing concentrations of MPA. The radiolabeled covalent enzyme-AMP complex was then visualized by autoradiography following electrophoresis on a denaturant polyacrylamide gel. The radiolabeled enzyme-AMP complex was then quantified by phosphorimaging (right side of the panel). (<b>C</b>) The second step of the ligase reaction is inhibited by MPA. The transfer of the AMP moiety onto a radiolabeled 5′-monophosphate 18-mer strand (pDNA) was evaluated by pre-incubating the enzyme with ATP to ensure formation of the radiolabeled covalent enzyme-AMP complex, followed by the addition of a 1-nt gapped substrate in the presence of increasing concentrations of MPA. Conversion of the 5′-³²P-labeled 18-mer strand into an adenylated species (AppDNA) was monitored by electrophoresis on a denaturant polyacrylamide gel. Lane 1: reaction performed in the absence of protein (-). The formation of the radiolabeled ApppDNA was quantified by phosphorimaging (<i>right side of the panel</i>).</p

Mycophenolic acid inhibits the RNA guanylyltransferase activity.

<p>(<b>A</b>) Molecular structure of mycophenolic acid (MPA). (<b>B</b>) Increasing concentrations of MPA inhibit the complete RNA guanylyltransferase reaction. A standard GTase assay in which the purified enzyme (1 µM) was incubated with both [alpha-³²P]GTP and a 5′-diphosphate acceptor RNA was performed in the presence of increasing concentrations of MPA. The reaction products were analyzed on a denaturing polyacrylamide gel and quantified (<i>right side of the panel</i>). (<b>C</b>) MPA is not a potent inhibitor of the first step of the GTase reaction. The formation of the enzyme-GMP covalent intermediate was monitored by incubating the purified enzyme (1 µM) in the presence of [alpha-³²P]GTP and increasing concentrations of MPA. The radiolabeled covalent enzyme-GMP complex was then visualized by autoradiography following electrophoresis on a denaturing 12.5% polyacrylamide gel. The radiolabeled enzyme-GMP complex was quantified by phosphorimaging (<i>right side of the panel</i>). (<b>D</b>) The second step of the GTase reaction is inhibited by MPA. The transfer of the GMP moiety onto an acceptor RNA was evaluated by pre-incubating the enzyme (1 µM) with [alpha-³²P]GTP (10 mM) to ensure formation of the radiolabeled covalent enzyme-GMP complex, followed by the addition of the acceptor 5′-diphosphate RNA (3 µM) in the presence of MPA. Formation of the radiolabeled capped GpppRNA was monitored following electrophoresis on a denaturant polyacrylamide gel. The radiolabeled GpppRNA was then quantified by phosphorimaging (<i>right side of the panel</i>).</p

Structural and mechanistic pathway used by RNA guanylyltransferases.

<p>The mechanism for phosphoryltransfer involves conformational changes between an open and closed form of the enzyme <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0024806#pone.0024806-Doherty1" target="_blank">[22]</a>, <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0024806#pone.0024806-Hakansson1" target="_blank">[23]</a>. GTP (grey sphere) initially binds to the apo-enzyme (structure 1) which promotes closure of the N-terminal nucleotidyl transferase (NT) domain and the C-terminal oligomer-binding (OB) fold domain (structure 3). This is followed by hydrolysis of the GTP substrate to produce the enzyme-GMP covalent intermediate (structure 4). Hydrolysis of GTP disrupts the interactions between the bound guanylate and the C-terminal OB fold domain, thus destabilizing the closed form of the enzyme, which opens up with the concomitant release of pyrophosphate (structure 5). This exposes the RNA-binding site of the enzyme (exact location unknown), thereby allowing the subsequent transfer of the GMP moiety onto the acceptor RNA (structure 7). The capped RNA is then released and the enzyme can reinitiate the pathway.</p

MPA inhibits RNA capping in <i>S. cerevisiae</i> cells.

<p>Primer extension assays were performed with a 5′ P³²-labeled 18-mer oligonucleotide complementary to the 5′ region of the <i>SSA1</i> mRNA. The oligonucleotide was annealed to total mRNAs extracted from cells that were grown in the absence (<i>−MPA</i>) or presence (<i>+MPA</i>) of 500 µg/ml MPA for 3 h, and extended with reverse transcriptase. The primer extension reactions were analyzed by electrophoresis through a 8% polyacrylamide gel containing 7 M urea in TBE and visualized by autoradiography. Control P³²-labeled RNA transcripts of 74 (<i>C1</i>) and 73 (<i>C2</i>) nucleotides were run in parallel. The positions and sizes (in nt) of the size markers are indicated on the <i>left</i>.</p

MPA inhibits the GTases of various origins.

<p>(<b>A</b>) Sequence conservation in members of the RNA/DNA nucleotidyltransferase superfamily. Members of this family share six conserved motifs (I, III, IIIa, IV, V, VI). An amino acid alignment of the GTases from <i>S. cerevisiae</i>, <i>C. albicans</i>, <i>Chlorella</i> virus, vaccinia virus, human, and the DNA ligase of <i>Chlorella</i> virus is presented. (B) Members of the RNA/DNA nucleotidyltransferase superfamily harbor a similar three-dimensional architecture consisting of an N-terminal NT domain and a C-terminal OB fold domain. The structures of the <i>S. cerevisiae</i> GTase (PDB: 3KYH), <i>C. albicans</i> GTase (PDB: 1P16), <i>Chlorella</i> virus GTase (PDB: 1CKN), and <i>Chlorella</i> virus DNA ligase (PDB: 1P8L) are shown. (C) The effect of MPA on the first and second step of the GTase reaction was monitored on the GTases from <i>S. cerevisiae</i>, <i>Chlorella</i> virus, vaccinia virus, and human. Both reactions were performed in the presence of increasing concentrations of MPA as described in the legend of <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0024806#pone-0024806-g001" target="_blank">figure 1</a>.</p

MPA is not a substrate for the RNA guanylyltransferase.

<p>(<b>A</b>) Capillary electrophoresis analysis of the RNA capping reaction. The GTase reactions were performed in the presence of the purified enzyme (1 µM) and GTP (1 mM) or MPA (1 mM), and the reaction products were analyzed by capillary electrophoresis. An untreated protein was also used as a control (-). The positions and sizes (in kDa) of the size markers (M) are indicated on the left. Masses are shown above the corresponding bands. (<b>B</b>) RNA capping reaction. The reaction mixtures contained 1 µg of purified enzyme, 23 pmol of radiolabelled 5′ diphosphate-terminated RNA (5′ p<b>p</b>G-RNA 3′, where the boldface indicates the radiolabelled moiety), and either 1 mM GTP or 1 mM MPA. An untreated control was also used in these assays (Ø). The reactions were incubated at 30°C for 30 min, and unincorporated nucleotides were removed by multiple rounds of ethanol precipitation. The RNAs were extracted with phenol/chloroform and recovered by ethanol precipitation. Aliquots of the RNA samples were adjusted to 50 mm NaOAc, pH 5.2, and digested with nuclease P1 (5 µg) for 60 min at 37°C. The reaction was then adjusted to 50 mm Tris-HCl, pH 8.0, and digested with alkaline phosphatase (1 unit) for 60 min at 37°C (<i>P1</i>+<i>AP</i>). The reaction products were analyzed by thin layer chromatography on a polyethyleneimine-cellulose plate developed with 0.5 M LiCl and 1 M formic acid. An autoradiogram of the plate is shown. The positions of the chromatographic origin (<i>ori</i>), inorganic phosphate (<i>Pi</i>), and GpppG are indicated.</p

Transcriptome-wide analysis of alternative RNA splicing events in Epstein-Barr virus-associated gastric carcinomas

<div><p>Multiple human diseases including cancer have been associated with a dysregulation in RNA splicing patterns. In the current study, modifications to the global RNA splicing landscape of cellular genes were investigated in the context of Epstein-Barr virus-associated gastric cancer. Global alterations to the RNA splicing landscape of cellular genes was examined in a large-scale screen from 295 primary gastric adenocarcinomas using high-throughput RNA sequencing data. RT-PCR analysis, mass spectrometry, and co-immunoprecipitation studies were also used to experimentally validate and investigate the differential alternative splicing (AS) events that were observed through RNA-seq studies. Our study identifies alterations in the AS patterns of approximately 900 genes such as tumor suppressor genes, transcription factors, splicing factors, and kinases. These findings allowed the identification of unique gene signatures for which AS is misregulated in both Epstein-Barr virus-associated gastric cancer and EBV-negative gastric cancer. Moreover, we show that the expression of Epstein–Barr nuclear antigen 1 (EBNA1) leads to modifications in the AS profile of cellular genes and that the EBNA1 protein interacts with cellular splicing factors. These findings provide insights into the molecular differences between various types of gastric cancer and suggest a role for the EBNA1 protein in the dysregulation of cellular AS.</p></div

DXO hydrolyzes only capped RNAs without a 2’-<i>O</i>-methylation.

<p>(A) The RNA 5’ cap structure is composed of a guanosine (blue) linked to the RNA (black) through a 5’-5’ triphosphate bridge. The subsequent N7-methylation of the guanosine (magenta) confers a positive charge to the cap structure. Additional 2’-<i>O</i>-methylations (orange) can be found on the first few nucleotides. (B) Nomenclature of the different cap structures. (C) Aliquots (2μg) of the purified preparations of DXO and mutant DXO protein (D236A/E253A) were analyzed by electrophoresis through a 12.5% polyacrylamide gel containing 0.1% SDS and visualized with Coomassie Blue Dye. The positions and sizes (in kDa) of the size markers are indicated on the left. (D) RNAs harbouring different cap structures were transcribed and capped (incorporation of [α-³²P]GTP) <i>in vitro</i>. They were then subjected to degradation by different enzymes, and reaction products were separated by thin layer chromatography. Lanes 1–4 show reaction products after treatment of differently capped RNAs with Nuclease P1. Degradation products after incubation of differently capped RNAs with purified DXO are shown in lanes 5–12. The origin of spotting and dinucleotide identities are listed on the left. NOTE: During the preparation of differently capped RNAs, only approximately 30% of GpppN-RNA was methylated to form GpppN_m-RNA (lanes 2,7–8), resulting in a mixture of GpppN-RNA and GpppN_m-RNA. Degradation products observed in lane 8 are due to the degradation of GpppN-RNA.</p