Data_Sheet_2_Genetic diversity and phylogeographic dynamics of avihepadnavirus: a comprehensive full-length genomic view.PDF

Avihepadnavirus is a genus of the Hepadnaviridae family. It primarily infects birds, including species of duck, geese, cranes, storks, and herons etc. To understand the genetic relatedness and evolutionary diversity among avihepadnavirus strains, a comprehensive analysis of the available 136 full-length viral genomes (n = 136) was conducted. The genomes were classified into two major genotypes, i.e., GI and GII. GI viruses were further classified into 8 sub-genotypes including DHBV-I (duck hepatitis B virus-I), DHBV-II (Snow goose Hepatitis B, SGHBV), DHBV-III, RGHBV (rossgoose hepatitis B virus), CHBV (crane hepatitis B virus), THBV (Tinamou hepatitis B virus), STHBV (stork hepatitis B virus), and HHBV (Heron hepatitis B virus). DHBV-I contains two sub-clades DHBV-Ia and DHBV-Ib. Parrot hepatitis B virus (PHBV) stains fall into GII which appeared as a separate phylogenetic branch/clade. All the subtypes of viruses in GI and GII seem to be genetically connected with viruses of DHBV-I by multiple mutational steps in phylogeographic analysis. Furthermore, 16 potential recombination events among different sub-genotypes in GI and one in GII were identified, but none of which is inter-genotypic between GI and GII. Overall, the results provide a whole picture of the genetic relatedness of avihepadnavirus strains, which may assist in the surveillance of virus spreading.</p

Data_Sheet_1_Genetic diversity and phylogeographic dynamics of avihepadnavirus: a comprehensive full-length genomic view.PDF

Avihepadnavirus is a genus of the Hepadnaviridae family. It primarily infects birds, including species of duck, geese, cranes, storks, and herons etc. To understand the genetic relatedness and evolutionary diversity among avihepadnavirus strains, a comprehensive analysis of the available 136 full-length viral genomes (n = 136) was conducted. The genomes were classified into two major genotypes, i.e., GI and GII. GI viruses were further classified into 8 sub-genotypes including DHBV-I (duck hepatitis B virus-I), DHBV-II (Snow goose Hepatitis B, SGHBV), DHBV-III, RGHBV (rossgoose hepatitis B virus), CHBV (crane hepatitis B virus), THBV (Tinamou hepatitis B virus), STHBV (stork hepatitis B virus), and HHBV (Heron hepatitis B virus). DHBV-I contains two sub-clades DHBV-Ia and DHBV-Ib. Parrot hepatitis B virus (PHBV) stains fall into GII which appeared as a separate phylogenetic branch/clade. All the subtypes of viruses in GI and GII seem to be genetically connected with viruses of DHBV-I by multiple mutational steps in phylogeographic analysis. Furthermore, 16 potential recombination events among different sub-genotypes in GI and one in GII were identified, but none of which is inter-genotypic between GI and GII. Overall, the results provide a whole picture of the genetic relatedness of avihepadnavirus strains, which may assist in the surveillance of virus spreading.</p

Graph of the ratio of fluorescent intensity to true normal by bead size

<p><b>Copyright information:</b></p><p>Taken from "Variance in multiplex suspension array assays: microsphere size variation impact"</p><p>http://www.tbiomed.com/content/4/1/31</p><p>Theoretical Biology & Medical Modelling 2007;4():31-31.</p><p>Published online 23 Aug 2007</p><p>PMCID:PMC2041949.</p><p></p> A ratio of 1 is for a microsphere sample equal to the specified mean microsphere diameter. This shows the relationship of microsphere diameter to brightness of signal, illustrating the correspondence of brightness as an rrelation

Histogram of a representative sample of events for one classifier from an event set

<p><b>Copyright information:</b></p><p>Taken from "Variance in multiplex suspension array assays: microsphere size variation impact"</p><p>http://www.tbiomed.com/content/4/1/31</p><p>Theoretical Biology & Medical Modelling 2007;4():31-31.</p><p>Published online 23 Aug 2007</p><p>PMCID:PMC2041949.</p><p></p> Classifier shown is region 97, N = 136. This histogram is based on fluorescent intensity of the reporter fluorophore. Compare with Figure 2

Ability of mutant RHAs to stimulate the synthesis of HIV-1 mRNAs.

<p>293T cells were cotransfected with SVC21.BH10 and either a plasmid expressing His-tagged wild-type or mutant RHA, or only the 6×His tag. 24 hours later, cell lysates and total cellular RNA were prepared and subjected to Western blotting and Northern blotting analysis respectively. (A) Western blots of cell lysates probed with anti-RHA, anti-His, or anti-β-actin. (B) Northern blotting. The total cellular RNA was resolved by electrophoresis on a denaturing 1% agarose gel, and blotted onto GeneScreen Plus membrane. The membrane was probed with the [32P]-labeled DNAs that are complimentary to HIV-1 5'-UTR. Ethidium bromide-stained rRNAs (18S and 28S) are included as an RNA loading control. Unspliced (US) ∼ 9.2 kb, singly spliced (SS) ∼ 4.0 kb, and multiply spliced (MS) ∼ 1.8 kb RNAs are indicated. (C) The intensity of RNA bands in panel B representing US, SS, or MS RNAs was quantitated using a PhosphorImager instrument, and are presented graphically. Shown is a representative of 3 independent experiments. (D) The ratio of US RNA to SS+MS RNA in panel B was determined. Shown are the mean values ± standard deviations of 3 independent experiments. *, <i>P</i> < 0.05 compared with values obtained with the 6×His tag alone (lane 1).</p

Ability of mutant RHAs to interact with HIV-1 RNA in the cell.

<p>293T cells were transfected with SVC21.BH10 and a plasmid expressing either His-tagged wild-type or mutant RHA, or only the 6×His tag. 24 hours later, cells were cross-linked, lysed, and sonicated. The cell lysates were incubated with Ni-NTA agarose to capture His-tagged protein. RNAs isolated from cell lysates (input) or from nucleoprotein bound to Ni-NTA agarose (precipitate) were subjected to RT-PCR analysis. (A) Western blot of cell lysates was probed with anti-His to detect expression of His-tagged RHA in transfected cells. The expressed 6×His tag peptide alone was not detectable in the Western blot. (B) Western blot of the precipitates was probed with anti-RHA. (C) The input RNA and RNA that was coprecipitated with His-tagged proteins were analyzed by RT-PCR, using primer pair P1-F/R <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0078596#pone.0078596-Xing2" target="_blank">[19]</a> specific to HIV-1 RNA. RT-PCR was performed in the presence (+) or absence (–) of reverse transcriptase.</p

Ability of mutant RHAs to promote the annealing of tRNA^Lys3 to viral RNA.

<p>293T cells were first treated with siRNA_Con or siRNA_RHA, and 16 hours later, were cotransfected with SVC21.BH10 and a plasmid expressing either 6×His tag, or His-tagged wild-type or mutant RHAs. 48 hours later, extracellular viruses were purified and cells were lysed. (A) Western blots of cell lysates probed with antibodies to RHA, His tag, or β-actin. (B) Western blots of viral lysates, containing equal amount of CAp24, probed with antibodies to RHA, His tag, CAp24, or RTp66/p51. (C) One nucleotide extension assay (+1 nt extension). Total viral RNA was isolated from purified HIV-1 particles, and tRNA^Lys3 annealed to viral RNA <i>in vivo</i> was extended by 1 nt ([³²P]-dCTP), using HIV-1 reverse transcriptase. The extended tRNA^Lys3 products are resolved by denaturing 1D PAGE, and visualized using a PhosphorImager. The control gel represents the +1 nt extension of a DNA primer annealed <i>in vitro</i> to viral RNA downstream of the tRNA^Lys3 binding site, and is used to show that approximately equal amounts of viral RNA were used in each extension reaction. (D) The values of the +1 nt extended tRNA^Lys3 products were quantitated using a PhosphorImager, normalized to the values obtained with virions produced from siRNA_Con-treated cells (lane 1), and are presented graphically as a percentage. Shown are the mean values ± standard deviations of 3 independent experiments. *, <i>P</i> < 0.05 compared with values obtained with virions produced from cells transfected with a plasmid expressing only His tag (lane 2).</p

Target the More Druggable Protein States in a Highly Dynamic Protein–Protein Interaction System

The proteins of the Bcl-2 family play key roles in the regulation of programmed cell death by controlling the integrity of the outer mitochondrial membrane and the initiation of the apoptosis process. We performed extensive molecular dynamics simulations to investigate the conformational flexibility of the Bcl-x_L protein in both the apo and holo (with Bad peptide and ABT-737) states. The accelerated molecular dynamics method implemented in Amber 14 was used to produce broader conformational sampling of 200 ns simulations. The pocket mining method based on the variational implicit-solvent model tracks the dynamic evolution of the ligand binding site with a druggability score characterizing the maximal affinity achievable by a drug-like molecule. Major movements were observed around the α3-helical domain and the loop region connecting the α1 and α2 helices, reshaping the ligand interaction in the BH3 binding groove. Starting with the apo crystal structure, which is recognized as “closed” and undruggable, the BH3 groove transitioned between the “open” and “closed” states during equilibrium simulation. Further analysis revealed a small percentage of the trajectory frames (∼10%) with a moderate degree of druggability that mimic the ligand-bound states. The ability to attain and detect by computer simulation the most suitable conformational states for ligand binding in advance of compound synthesis and crystal structure solution is of immense value to the application and success of structure-based drug design

Ability of purified mutant RHAs to bind and unwind dsRNA <i>in vitro</i>.

<p>(A) Isolation of mutant full-length RHAs. The mutant RHAs containing deletions in the linker region were purified from mammalian cells (HEK 293E), separated by 1D SDS-PAGE, and analyzed by either staining with Coomassie Brilliant Blue R250 (CBR) or probing Western blots with anti-His (WB). Lane M shows a protein size marker with indicated molecular weights in kDa. (B) Diagram of the duplex RNA substrate with one strand 3′-end-labeled with ³²pCp. (C) EMSA was carried out to examine the <i>in vitro</i> binding of purified proteins to [³²pCp]-labeled synthetic duplex RNA. GST was included as a negative control. Shown is a representative of 3 independent experiments. (D) Helicase activity assay. 10 nM of radioactive duplex RNAs and 150 nM of indicated proteins were incubated at 37°C for indicated time periods in the presence or absence of 1 mM ATP, and then the radioactive RNA strand in single or duplex form was resolved on a 15% native polyacrylamide gel, and visualized using a PhosphorImager. Lane C indicates the migration position of the ssRNA (boiled substrate), while lane N represents the migration position of untreated radioactive duplex RNA. Lanes at time point 0 represent the helicase reactions in the absence of 1 mM ATP. Shown is a representative of 3 independent experiments. (E) Duplex RNA bands in panel D were quantitated using a PhosphorImager, normalized to the results obtained at the time point 0, and presented graphically as percentage. Shown are the mean values ± standard deviations of 3 independent experiments. *, <i>P</i> < 0.05 compared with corresponding values obtained with wild-type RHA.</p

Effect of mutations in the linker region of RHA upon helicase activity <i>in vitro</i> and upon multiple steps in HIV-1 production in the cells<i>^a</i>.

a±<p>±, Partial reduction.</p