rnase h  (New England Biolabs)


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    Name:
    RNase H
    Description:
    RNase H 1 250 units
    Catalog Number:
    M0297L
    Price:
    283
    Category:
    Ribonucleases RNase
    Size:
    1 250 units
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    New England Biolabs rnase h
    RNase H
    RNase H 1 250 units
    https://www.bioz.com/result/rnase h/product/New England Biolabs
    Average 99 stars, based on 1 article reviews
    Price from $9.99 to $1999.99
    rnase h - by Bioz Stars, 2021-05
    99/100 stars

    Images

    1) Product Images from "qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide"

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkaa500

    Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing RNase H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.
    Figure Legend Snippet: Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing RNase H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.

    Techniques Used: Synthesized, Amplification, In Vitro, Purification, Agarose Gel Electrophoresis, Hybridization, Staining, Real-time Polymerase Chain Reaction, Inhibition

    qDRIP provides strand-specific, high resolution RNA–DNA hybrid mapping. ( A ) Schematic of qDRIP experimental process. ( B ) Representative genome browser view of qDRIP-seq signal. From top to bottom: two qDRIP-seq biological replicates, RNase H digested sample pooled prior to IP, and input pooled from replicates. All tracks normalized by reads per million mapped. Negative strand signal in red, positive in blue. Bent arrows represent TSS, while large triangular arrows represent TES (transcription end site). ( C ) Read counts from template strand (TS) and non-template strand (NTS) of hybrids, as well as from ssDNA and dsDNA negative controls. ( D ) GC (green) and AT (red) skew across coding strand of qDRIP peaks, including 600 bp flanking 5’- and 3’-ends. ( E ) Fractions of qDRIP peaks overlapping noted genomic features ( P = 2.5e–2798, chi-square test). ( F ) Scaled metaplot of sense hybrids between TSS and first-intron exon boundary, as well as 1 kb upstream of TSS and 1 kb downstream of first intron-exon boundary. Tracks shown are mean IP (blue) and pooled input (grey). Bands represent 95% CI of mean read signal.
    Figure Legend Snippet: qDRIP provides strand-specific, high resolution RNA–DNA hybrid mapping. ( A ) Schematic of qDRIP experimental process. ( B ) Representative genome browser view of qDRIP-seq signal. From top to bottom: two qDRIP-seq biological replicates, RNase H digested sample pooled prior to IP, and input pooled from replicates. All tracks normalized by reads per million mapped. Negative strand signal in red, positive in blue. Bent arrows represent TSS, while large triangular arrows represent TES (transcription end site). ( C ) Read counts from template strand (TS) and non-template strand (NTS) of hybrids, as well as from ssDNA and dsDNA negative controls. ( D ) GC (green) and AT (red) skew across coding strand of qDRIP peaks, including 600 bp flanking 5’- and 3’-ends. ( E ) Fractions of qDRIP peaks overlapping noted genomic features ( P = 2.5e–2798, chi-square test). ( F ) Scaled metaplot of sense hybrids between TSS and first-intron exon boundary, as well as 1 kb upstream of TSS and 1 kb downstream of first intron-exon boundary. Tracks shown are mean IP (blue) and pooled input (grey). Bands represent 95% CI of mean read signal.

    Techniques Used:

    R-loop lifetimes. ( A ) Schematic of transcription with and without DRB. ( B ) Ratio of DRB to control signal in RNase H-sensitive peaks, compared to estimated time without transcription. Error bands are 95% CI of the mean. Horizontal dotted line indicates a 2-fold decrease in DRB signal. ( C ) qPCR measurements during a DRB timecourse at regions predicted to be unstable (top) or stable (bottom) by pseudo-timecourse obtained from sequencing data. Error bars represent 95% CI of the mean. In primer name, GB indicates gene body. ( D ) GC content across 500 bp regions with shorter, longer or close to average (NS) lifetimes ( P  = 2.5e–143, Kruskal–Wallis test). ( E ) Biochemically determined G-quadruplex counts (  37 ) over the same regions as (D) ( P  = 2.7e–7, ANOVA on Negative Binomial regression, likelihood ratio test). ( F ) Relative replication fork directionality (RFD) (  39 ) to transcription over the same regions as (D), where 1 represents fully co-directional and –1 represents fully head-on ( P  = 3.5e–12, Kruskal–Wallis test). ( G ) Distribution of half-lives assuming first-order decay.
    Figure Legend Snippet: R-loop lifetimes. ( A ) Schematic of transcription with and without DRB. ( B ) Ratio of DRB to control signal in RNase H-sensitive peaks, compared to estimated time without transcription. Error bands are 95% CI of the mean. Horizontal dotted line indicates a 2-fold decrease in DRB signal. ( C ) qPCR measurements during a DRB timecourse at regions predicted to be unstable (top) or stable (bottom) by pseudo-timecourse obtained from sequencing data. Error bars represent 95% CI of the mean. In primer name, GB indicates gene body. ( D ) GC content across 500 bp regions with shorter, longer or close to average (NS) lifetimes ( P = 2.5e–143, Kruskal–Wallis test). ( E ) Biochemically determined G-quadruplex counts ( 37 ) over the same regions as (D) ( P = 2.7e–7, ANOVA on Negative Binomial regression, likelihood ratio test). ( F ) Relative replication fork directionality (RFD) ( 39 ) to transcription over the same regions as (D), where 1 represents fully co-directional and –1 represents fully head-on ( P = 3.5e–12, Kruskal–Wallis test). ( G ) Distribution of half-lives assuming first-order decay.

    Techniques Used: Real-time Polymerase Chain Reaction, Sequencing

    RNase H resistant signal. ( A ) Aggregate plot of qDRIP-seq signal around the TSS of top 10,000 expressed genes as determined by GRO-seq ( 36 ). Tracks are IP (blue), RHR (red) and input (grey). Error bands represent 95% CI of mean. ( B ) Heatmaps of mean IP signal, RNase H-resistant signal and GC-skew around top 10,000 promoters ranked by GC-skew immediately (0–500 bp) downstream of the TSS. Correlation coefficient between IP signal and GC-skew was 0.06, whereas correlation coefficient for RHR signal was 0.22 (Spearman's rho). ( C ) GC-skew around RNase H resistant regions within the full (unfiltered) qDRIP-seq peak set. qDRIP peaks (red) compared to regions of equal lengths randomly selected from non-resistant qDRIP-peaks (blue). As before, bands represent 95% CI of mean read signal. ( D ) Same as (C), but showing biochemically determined G-quadruplex density ( 37 ) over these regions. ( E ) RNase H-resistant signal around RH-resistant peak calls. Peaks lying 5’ in genes (which DRB should affect) are in blue, while peaks lying 3’ in genes (which DRB should not affect) are in red. Left panel is RNase H treatment in control cells, while right panel is RNase H treatment in DRB treated cells. As before, error bands represent 95% CI of the mean.
    Figure Legend Snippet: RNase H resistant signal. ( A ) Aggregate plot of qDRIP-seq signal around the TSS of top 10,000 expressed genes as determined by GRO-seq ( 36 ). Tracks are IP (blue), RHR (red) and input (grey). Error bands represent 95% CI of mean. ( B ) Heatmaps of mean IP signal, RNase H-resistant signal and GC-skew around top 10,000 promoters ranked by GC-skew immediately (0–500 bp) downstream of the TSS. Correlation coefficient between IP signal and GC-skew was 0.06, whereas correlation coefficient for RHR signal was 0.22 (Spearman's rho). ( C ) GC-skew around RNase H resistant regions within the full (unfiltered) qDRIP-seq peak set. qDRIP peaks (red) compared to regions of equal lengths randomly selected from non-resistant qDRIP-peaks (blue). As before, bands represent 95% CI of mean read signal. ( D ) Same as (C), but showing biochemically determined G-quadruplex density ( 37 ) over these regions. ( E ) RNase H-resistant signal around RH-resistant peak calls. Peaks lying 5’ in genes (which DRB should affect) are in blue, while peaks lying 3’ in genes (which DRB should not affect) are in red. Left panel is RNase H treatment in control cells, while right panel is RNase H treatment in DRB treated cells. As before, error bands represent 95% CI of the mean.

    Techniques Used:

    2) Product Images from "Structural basis for the tryptophan sensitivity of TnaC-mediated ribosome stalling"

    Article Title: Structural basis for the tryptophan sensitivity of TnaC-mediated ribosome stalling

    Journal: bioRxiv

    doi: 10.1101/2021.03.31.437805

    Complex purification and cryo-EM data processing workflow. Stalled ribosomal complexes were prepared using a bicistronic operon containing two identical copies of tnaC or tnaC(R23F) . A first sucrose gradient was performed to collect polysomes, followed by a second sucrose gradient after RNase H treatment to collect the monosomal fraction, which was the used to prepare the grids for cryo-EM data acquisition. The flowchart shows the workflow used to process and analyze cryo-EM data. Cross-sections of the final reconstructions are shown with the 70S ribosome in white, the mRNA in violet, the P-site tRNA in pink, the TnaC peptide in red and the L-Trp molecule in orange. Detailed maps of the TnaC peptide and L-Trp density are also shown for the TnaC–70S and TnaC(R23F)–70S complexes using the same color scheme, along with fitted atomic models. Fourier Shell Correlation (FSC) curves of the final reconstructions are shown as calculated by the RELION 3 . 1 (Ref. 51) post-processing algorithm. Cross-sections of maps displaying the local resolution calculated by RELION 3 . 1 (Ref. 51) are shown.
    Figure Legend Snippet: Complex purification and cryo-EM data processing workflow. Stalled ribosomal complexes were prepared using a bicistronic operon containing two identical copies of tnaC or tnaC(R23F) . A first sucrose gradient was performed to collect polysomes, followed by a second sucrose gradient after RNase H treatment to collect the monosomal fraction, which was the used to prepare the grids for cryo-EM data acquisition. The flowchart shows the workflow used to process and analyze cryo-EM data. Cross-sections of the final reconstructions are shown with the 70S ribosome in white, the mRNA in violet, the P-site tRNA in pink, the TnaC peptide in red and the L-Trp molecule in orange. Detailed maps of the TnaC peptide and L-Trp density are also shown for the TnaC–70S and TnaC(R23F)–70S complexes using the same color scheme, along with fitted atomic models. Fourier Shell Correlation (FSC) curves of the final reconstructions are shown as calculated by the RELION 3 . 1 (Ref. 51) post-processing algorithm. Cross-sections of maps displaying the local resolution calculated by RELION 3 . 1 (Ref. 51) are shown.

    Techniques Used: Purification, Cryo-EM Sample Prep

    3) Product Images from "Argonaute-based programmable RNase as a tool for cleavage of highly-structured RNA"

    Article Title: Argonaute-based programmable RNase as a tool for cleavage of highly-structured RNA

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gky496

    Comparing cleavage activity of DISC and RNase H. ( A ) Schematic of matched and mismatched guide and target pairs used to target four TRs across the HIV-1 ΔDIS 5′UTR RNA. For each pair, the HIV-1 ΔDIS 5′UTR sequence is shown on top and the perfectly matched gDNA strand is shown on the bottom. Circle indicates target position complementary to the first position of the guide that does not pair due to structural restrains by the protein. Black arrowheads indicate cleavage site. Mismatches between the guide and target strands are indicated by a black box around the bases of the guide that are mutated to the bases shown below the box. ( B ) Quantified cleavage products from the assay using matched and mismatched guide and target pairs described in (A) are plotted with solid bars representing the average of three replicates and circles representing individual replicates. Cleavage that was not detectable by the assay is indicated by ‘nd’. ( C and D ) Comparing DISC (circles) and RNase H (triangles) cleavage of the unstructured 60-nt target (C) or of a structured 352-nt RNA target (D). Bars indicate average cleavage of three replicates.
    Figure Legend Snippet: Comparing cleavage activity of DISC and RNase H. ( A ) Schematic of matched and mismatched guide and target pairs used to target four TRs across the HIV-1 ΔDIS 5′UTR RNA. For each pair, the HIV-1 ΔDIS 5′UTR sequence is shown on top and the perfectly matched gDNA strand is shown on the bottom. Circle indicates target position complementary to the first position of the guide that does not pair due to structural restrains by the protein. Black arrowheads indicate cleavage site. Mismatches between the guide and target strands are indicated by a black box around the bases of the guide that are mutated to the bases shown below the box. ( B ) Quantified cleavage products from the assay using matched and mismatched guide and target pairs described in (A) are plotted with solid bars representing the average of three replicates and circles representing individual replicates. Cleavage that was not detectable by the assay is indicated by ‘nd’. ( C and D ) Comparing DISC (circles) and RNase H (triangles) cleavage of the unstructured 60-nt target (C) or of a structured 352-nt RNA target (D). Bars indicate average cleavage of three replicates.

    Techniques Used: Activity Assay, Sequencing

    4) Product Images from "Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †"

    Article Title: Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †

    Journal:

    doi: 10.1128/MCB.01410-07

    RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)
    Figure Legend Snippet: RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)

    Techniques Used:

    5) Product Images from "The ApaH-like phosphatase TbALPH1 is the major mRNA decapping enzyme of trypanosomes"

    Article Title: The ApaH-like phosphatase TbALPH1 is the major mRNA decapping enzyme of trypanosomes

    Journal: PLoS Pathogens

    doi: 10.1371/journal.ppat.1006456

    mRNAs that accumulate after ALPH1 RNAi are deadenylated. Northern blots were probed for histone H4 , a very small mRNA that allows to visualise changes in poly(A) tail length by band shifts. (A) RNA was isolated over a time-course of ALPH1 RNAi depletion. Representative data from one of three RNAi clones are shown. (B) RNA isolated after 0 or 72 hours of ALPH1 RNAi depletion was treated with RNAse H in the absence or presence of oligo dT. Samples not treated with RNAse H served as control. The blot was re-probed for two small RNAs that have no poly(A) tail (5.8S rRNA and SL RNA) to demonstrate that the band-shift is specific to polyadenylated RNAs. (C) RNA was isolated over a time-course of CAF1 RNAi depletion. Representative data from one of three RNAi clones are shown. (D) RNA was isolated over a time-course of XRNA RNAi depletion. Representative data from one of two RNAi clones are shown.
    Figure Legend Snippet: mRNAs that accumulate after ALPH1 RNAi are deadenylated. Northern blots were probed for histone H4 , a very small mRNA that allows to visualise changes in poly(A) tail length by band shifts. (A) RNA was isolated over a time-course of ALPH1 RNAi depletion. Representative data from one of three RNAi clones are shown. (B) RNA isolated after 0 or 72 hours of ALPH1 RNAi depletion was treated with RNAse H in the absence or presence of oligo dT. Samples not treated with RNAse H served as control. The blot was re-probed for two small RNAs that have no poly(A) tail (5.8S rRNA and SL RNA) to demonstrate that the band-shift is specific to polyadenylated RNAs. (C) RNA was isolated over a time-course of CAF1 RNAi depletion. Representative data from one of three RNAi clones are shown. (D) RNA was isolated over a time-course of XRNA RNAi depletion. Representative data from one of two RNAi clones are shown.

    Techniques Used: Northern Blot, Isolation, Clone Assay, Electrophoretic Mobility Shift Assay

    6) Product Images from "Improvement of RNA secondary structure prediction using RNase H cleavage and randomized oligonucleotides"

    Article Title: Improvement of RNA secondary structure prediction using RNase H cleavage and randomized oligonucleotides

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkp587

    Phylogenetic secondary structure, the predicted lowest free energy secondary structure, and the four suboptimal structures of E. coli 5S rRNA. Loops A–E are labeled in the phylogenetic structure. Base-paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H cleavage are circled. The lowest free energy structure only has 27% of the base pairs present in the phylogenetic structure.
    Figure Legend Snippet: Phylogenetic secondary structure, the predicted lowest free energy secondary structure, and the four suboptimal structures of E. coli 5S rRNA. Loops A–E are labeled in the phylogenetic structure. Base-paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H cleavage are circled. The lowest free energy structure only has 27% of the base pairs present in the phylogenetic structure.

    Techniques Used: Labeling

    Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in yeast tRNA Phe with randomized 5-mer oligonucleotides.
    Figure Legend Snippet: Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in yeast tRNA Phe with randomized 5-mer oligonucleotides.

    Techniques Used:

    Schematic of the general method used in this study. Randomized DNA oligonucleotides are incubated with an RNA of interest. Only DNAs complementary to single-stranded regions bind, inducing RNase H cleavage of the RNA strand. Nucleotides which are subject to RNase H cleavage are used as single-stranded constraints in RNA secondary structure prediction.
    Figure Legend Snippet: Schematic of the general method used in this study. Randomized DNA oligonucleotides are incubated with an RNA of interest. Only DNAs complementary to single-stranded regions bind, inducing RNase H cleavage of the RNA strand. Nucleotides which are subject to RNase H cleavage are used as single-stranded constraints in RNA secondary structure prediction.

    Techniques Used: Incubation

    Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in E. coli 5S rRNA. The numbers above the lanes indicate to which nucleotides in the RNA the oligonucleotide probe is complementary.
    Figure Legend Snippet: Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in E. coli 5S rRNA. The numbers above the lanes indicate to which nucleotides in the RNA the oligonucleotide probe is complementary.

    Techniques Used:

    Phylogenetic secondary structure and three predicted suboptimal structures of yeast tRNA Phe . The lowest free energy structure has 95% of the base pairs predicted correctly. Base paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H after 1 or 2 h incubation are circled. D denotes dihydrouracil while Y denotes wybutosine.
    Figure Legend Snippet: Phylogenetic secondary structure and three predicted suboptimal structures of yeast tRNA Phe . The lowest free energy structure has 95% of the base pairs predicted correctly. Base paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H after 1 or 2 h incubation are circled. D denotes dihydrouracil while Y denotes wybutosine.

    Techniques Used: Incubation

    7) Product Images from "The final step of 40S ribosomal subunit maturation is controlled by a dual key lock"

    Article Title: The final step of 40S ribosomal subunit maturation is controlled by a dual key lock

    Journal: bioRxiv

    doi: 10.1101/2020.07.29.226936

    Assessment of the size of the ITS1 in purified pre-40S particles. Comparison of RNAse H digestion and alkaline hydrolysis assays shows nucleotide resolution between bands. Left panel, RNase H digestion of rRNAs from pre-40S particles purified using a HASt tagged version of LTV1 as bait (HASt-LTV1). Right panel, alkaline hydrolysis (OH - ladder) of an RNA molecule containing the 18S-ITS1 sequence recognized by the 3’18S probe at its 5’ end (see Suppl. File 2). The samples were fractionated ob a 12% polyacrylamide gel and northern blot was probed with the 3’18S probe.
    Figure Legend Snippet: Assessment of the size of the ITS1 in purified pre-40S particles. Comparison of RNAse H digestion and alkaline hydrolysis assays shows nucleotide resolution between bands. Left panel, RNase H digestion of rRNAs from pre-40S particles purified using a HASt tagged version of LTV1 as bait (HASt-LTV1). Right panel, alkaline hydrolysis (OH - ladder) of an RNA molecule containing the 18S-ITS1 sequence recognized by the 3’18S probe at its 5’ end (see Suppl. File 2). The samples were fractionated ob a 12% polyacrylamide gel and northern blot was probed with the 3’18S probe.

    Techniques Used: Purification, Sequencing, Northern Blot

    RPS26 is required for rRNA cleavage at site 3 as well as NOB1 and DIM2 release. HEK cell lines expressing tagged version of LTV1, the catalytically inactive RIO1-D324A (RIO1 (kd)) or wild-type RIO1 (RIO1(wt)) were treated with scramble or RPS26 siRNAs for 48 h. a , RNase H assays were conducted as in Figure 4 on rRNAs of pre-40S particles purified with the mentioned StHA tagged bait, either from RPS26-depleted cells or from control cells (scramble siRNA). b , Signals corresponding to the 18S-E and 18S rRNA detected in (a) were quantified and represented as the 18S/18S-E ratio for the different pre-40S particles. Error bars, s.d. (n=3) c , Cell extracts and purified particles were analysed by Western Blot using the indicated antibodies. d , Bands corresponding to DIM2 and NOB1 (in the eluates) were quantified, corrected for pre-40S particle loading (using RPS19) and normalized to the control condition (set to 1). Error bars, s.d. (n=3).
    Figure Legend Snippet: RPS26 is required for rRNA cleavage at site 3 as well as NOB1 and DIM2 release. HEK cell lines expressing tagged version of LTV1, the catalytically inactive RIO1-D324A (RIO1 (kd)) or wild-type RIO1 (RIO1(wt)) were treated with scramble or RPS26 siRNAs for 48 h. a , RNase H assays were conducted as in Figure 4 on rRNAs of pre-40S particles purified with the mentioned StHA tagged bait, either from RPS26-depleted cells or from control cells (scramble siRNA). b , Signals corresponding to the 18S-E and 18S rRNA detected in (a) were quantified and represented as the 18S/18S-E ratio for the different pre-40S particles. Error bars, s.d. (n=3) c , Cell extracts and purified particles were analysed by Western Blot using the indicated antibodies. d , Bands corresponding to DIM2 and NOB1 (in the eluates) were quantified, corrected for pre-40S particle loading (using RPS19) and normalized to the control condition (set to 1). Error bars, s.d. (n=3).

    Techniques Used: Expressing, Purification, Western Blot

    Late RIO1(kd)StHA pre-40S particles contain a high proportion of mature 18S rRNA. a , Diagram representing steps of the pre-40S rRNA digestion by RNase H. b , RNase H assays were performed on RNAs extracted from pre-40S particles purified with the mentioned StHA tagged bait, and separated on a 12% polyacrylamide gel. The 18S rRNA and its precursors were revealed by the 3’18S radiolabeled probe. Bands are separated with single nucleotide resolution, as shown in Figure 4 – figure supplement 1 . c , Signals corresponding to the 18S-E and 18S rRNAs were quantified by phosphorimaging and represented by the 18S/18S-E ratio for the different purified pre-40S particles. The average of three independent experiments is shown, with the standard deviation indicated on top of the histogram.
    Figure Legend Snippet: Late RIO1(kd)StHA pre-40S particles contain a high proportion of mature 18S rRNA. a , Diagram representing steps of the pre-40S rRNA digestion by RNase H. b , RNase H assays were performed on RNAs extracted from pre-40S particles purified with the mentioned StHA tagged bait, and separated on a 12% polyacrylamide gel. The 18S rRNA and its precursors were revealed by the 3’18S radiolabeled probe. Bands are separated with single nucleotide resolution, as shown in Figure 4 – figure supplement 1 . c , Signals corresponding to the 18S-E and 18S rRNAs were quantified by phosphorimaging and represented by the 18S/18S-E ratio for the different purified pre-40S particles. The average of three independent experiments is shown, with the standard deviation indicated on top of the histogram.

    Techniques Used: Purification, Standard Deviation

    In vitro cleavage of the 18S-E pre-rRNA within pre-40S particles is stimulated by ATP addition. HEK cell lines expressing tagged versions of wild-type RIO1 (RIO1(wt)) or of the catalytically-inactive RIO1 (kd) were treated with scramble or RPS26 siRNAs for 48h to enrich particles in state A. Pre-40S particles were purified and incubated in the presence of 1 mM ATP, 1mM AMP-PNP, or without nucleotide (mock condition). a , RNAse H assays were performed on the RNAs extracted from the particles. b , The variation of cleavage efficiency with the different nucleotides is indicated by the 18S/18S-E ratio and normalized to the mock-treated sample (set to 1). The data correspond to five independent experiments. Analysis of the results with a unilateral paired Wilcoxon test (“sample greater than mock”) indicates p-values of 0.031 for samples RIO1(wt)-ATP, RIO1(wt)-AMP-PNP, RIO1(kd)-ATP, and 0.063 for RIO1(kd)-AMP-PNP. c , Superimposition of atomic models of State A and B reveals overlapping distances (grey lines) between atoms of Proline 351 from RIO1 (green) and of Arginine 247 from DIM2 (orange). RPS5, which seems to be repositioned upon association of RIO1 / dissociation of DIM2 from the pre-40S particle, is shown in violet (State A) or white (State B).
    Figure Legend Snippet: In vitro cleavage of the 18S-E pre-rRNA within pre-40S particles is stimulated by ATP addition. HEK cell lines expressing tagged versions of wild-type RIO1 (RIO1(wt)) or of the catalytically-inactive RIO1 (kd) were treated with scramble or RPS26 siRNAs for 48h to enrich particles in state A. Pre-40S particles were purified and incubated in the presence of 1 mM ATP, 1mM AMP-PNP, or without nucleotide (mock condition). a , RNAse H assays were performed on the RNAs extracted from the particles. b , The variation of cleavage efficiency with the different nucleotides is indicated by the 18S/18S-E ratio and normalized to the mock-treated sample (set to 1). The data correspond to five independent experiments. Analysis of the results with a unilateral paired Wilcoxon test (“sample greater than mock”) indicates p-values of 0.031 for samples RIO1(wt)-ATP, RIO1(wt)-AMP-PNP, RIO1(kd)-ATP, and 0.063 for RIO1(kd)-AMP-PNP. c , Superimposition of atomic models of State A and B reveals overlapping distances (grey lines) between atoms of Proline 351 from RIO1 (green) and of Arginine 247 from DIM2 (orange). RPS5, which seems to be repositioned upon association of RIO1 / dissociation of DIM2 from the pre-40S particle, is shown in violet (State A) or white (State B).

    Techniques Used: In Vitro, Expressing, Purification, Incubation

    8) Product Images from "BRCA2 controls DNA:RNA hybrid level at DSBs by mediating RNase H2 recruitment"

    Article Title: BRCA2 controls DNA:RNA hybrid level at DSBs by mediating RNase H2 recruitment

    Journal: Nature Communications

    doi: 10.1038/s41467-018-07799-2

    DNA:RNA hybrids form at DSBs independently of the genomic context. a Schematic representation of DNA:RNA hybrids (in red) that can be generated upon the hybridization of mRNA (top) or dilncRNAs (bottom) with resected DNA ends at the I-PpoI cut site within DAB1 gene. b DRIP-qPCR analysis at the I-PpoI cut site within a genic ( DAB1 gene) or c nongenic locus in HeLa cells transfected with the I-PpoI nuclease. d DRIP-qPCR analysis at a nongenic AsiSI cut site in DIvA cells. Bar graphs in b , c and d show fold induction of DNA:RNA hybrid levels in cut samples relative to uncut. RNase H treatment was performed on cut samples to demonstrate specificity of the signal. Error bars represent s.e.m. ( n ≥ 3 independent experiments). * P
    Figure Legend Snippet: DNA:RNA hybrids form at DSBs independently of the genomic context. a Schematic representation of DNA:RNA hybrids (in red) that can be generated upon the hybridization of mRNA (top) or dilncRNAs (bottom) with resected DNA ends at the I-PpoI cut site within DAB1 gene. b DRIP-qPCR analysis at the I-PpoI cut site within a genic ( DAB1 gene) or c nongenic locus in HeLa cells transfected with the I-PpoI nuclease. d DRIP-qPCR analysis at a nongenic AsiSI cut site in DIvA cells. Bar graphs in b , c and d show fold induction of DNA:RNA hybrid levels in cut samples relative to uncut. RNase H treatment was performed on cut samples to demonstrate specificity of the signal. Error bars represent s.e.m. ( n ≥ 3 independent experiments). * P

    Techniques Used: Generated, Hybridization, Real-time Polymerase Chain Reaction, Transfection

    DNA:RNA hybrids are directly recognized by BRCA1 in vitro and in vivo. a Representative pictures of super-resolution imaging analysis of BRCA1 (cyan) and DNA:RNA hybrids (yellow) colocalization in S-phase synchronized NCS-treated U2OS cells. Scale bar: 5 μm. b Dot plot shows the normalized number of overlaps relative to random of BRCA1 and DNA:RNA hybrids signals in S-phase U2OS cells treated with DSMO or NCS. At least n = 40 events were counted from three independent experiments. Lines represent mean ± s.e.m. c Electrophoretic mobility shift assay (EMSA) of purified recombinant human BRCA1 or BRCA1-BARD1 with end-labeled (*) double-stranded DNA or DNA:RNA substrates. d Graph showing the percentage of protein-bound substrate at respective protein concentrations. Error bars represent s.e.m. ( n = 2 independent experiments). e Representative images of BRCA1 foci co-stained with cyclin A, as S/G2-phase marker, in irradiated (2 Gy) U2OS cells over-expressing GFP or GFP-RNase H1 (GFP-RH1). Scale bar: 5 μm. f Dot plot shows the number of foci in e . At least n = 80 cells were counted from at least three independent experiments. Lines represent mean ± s.e.m. g Representative images of BRCA1 foci co-stained with cyclin A, as S/G2-phase marker, in irradiated (2 Gy) U2OS cells treated with RNase H prior to fixation. Scale bar: 10 μm. h Dot plot shows the number of foci in g . At least n = 80 cells were counted from three independent experiments. Lines represent mean ± s.e.m. * P
    Figure Legend Snippet: DNA:RNA hybrids are directly recognized by BRCA1 in vitro and in vivo. a Representative pictures of super-resolution imaging analysis of BRCA1 (cyan) and DNA:RNA hybrids (yellow) colocalization in S-phase synchronized NCS-treated U2OS cells. Scale bar: 5 μm. b Dot plot shows the normalized number of overlaps relative to random of BRCA1 and DNA:RNA hybrids signals in S-phase U2OS cells treated with DSMO or NCS. At least n = 40 events were counted from three independent experiments. Lines represent mean ± s.e.m. c Electrophoretic mobility shift assay (EMSA) of purified recombinant human BRCA1 or BRCA1-BARD1 with end-labeled (*) double-stranded DNA or DNA:RNA substrates. d Graph showing the percentage of protein-bound substrate at respective protein concentrations. Error bars represent s.e.m. ( n = 2 independent experiments). e Representative images of BRCA1 foci co-stained with cyclin A, as S/G2-phase marker, in irradiated (2 Gy) U2OS cells over-expressing GFP or GFP-RNase H1 (GFP-RH1). Scale bar: 5 μm. f Dot plot shows the number of foci in e . At least n = 80 cells were counted from at least three independent experiments. Lines represent mean ± s.e.m. g Representative images of BRCA1 foci co-stained with cyclin A, as S/G2-phase marker, in irradiated (2 Gy) U2OS cells treated with RNase H prior to fixation. Scale bar: 10 μm. h Dot plot shows the number of foci in g . At least n = 80 cells were counted from three independent experiments. Lines represent mean ± s.e.m. * P

    Techniques Used: In Vitro, In Vivo, Imaging, Electrophoretic Mobility Shift Assay, Purification, Recombinant, Labeling, Staining, Marker, Irradiation, Expressing

    9) Product Images from "Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter"

    Article Title: Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkx403

    Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.
    Figure Legend Snippet: Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.

    Techniques Used: Sequencing, Quantitation Assay

    Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).
    Figure Legend Snippet: Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).

    Techniques Used: Sequencing

    10) Product Images from "qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide"

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkaa500

    Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing RNase H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.
    Figure Legend Snippet: Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing RNase H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.

    Techniques Used: Synthesized, Amplification, In Vitro, Purification, Agarose Gel Electrophoresis, Hybridization, Staining, Real-time Polymerase Chain Reaction, Inhibition

    qDRIP provides strand-specific, high resolution RNA–DNA hybrid mapping. ( A ) Schematic of qDRIP experimental process. ( B ) Representative genome browser view of qDRIP-seq signal. From top to bottom: two qDRIP-seq biological replicates, RNase H digested sample pooled prior to IP, and input pooled from replicates. All tracks normalized by reads per million mapped. Negative strand signal in red, positive in blue. Bent arrows represent TSS, while large triangular arrows represent TES (transcription end site). ( C ) Read counts from template strand (TS) and non-template strand (NTS) of hybrids, as well as from ssDNA and dsDNA negative controls. ( D ) GC (green) and AT (red) skew across coding strand of qDRIP peaks, including 600 bp flanking 5’- and 3’-ends. ( E ) Fractions of qDRIP peaks overlapping noted genomic features ( P = 2.5e–2798, chi-square test). ( F ) Scaled metaplot of sense hybrids between TSS and first-intron exon boundary, as well as 1 kb upstream of TSS and 1 kb downstream of first intron-exon boundary. Tracks shown are mean IP (blue) and pooled input (grey). Bands represent 95% CI of mean read signal.
    Figure Legend Snippet: qDRIP provides strand-specific, high resolution RNA–DNA hybrid mapping. ( A ) Schematic of qDRIP experimental process. ( B ) Representative genome browser view of qDRIP-seq signal. From top to bottom: two qDRIP-seq biological replicates, RNase H digested sample pooled prior to IP, and input pooled from replicates. All tracks normalized by reads per million mapped. Negative strand signal in red, positive in blue. Bent arrows represent TSS, while large triangular arrows represent TES (transcription end site). ( C ) Read counts from template strand (TS) and non-template strand (NTS) of hybrids, as well as from ssDNA and dsDNA negative controls. ( D ) GC (green) and AT (red) skew across coding strand of qDRIP peaks, including 600 bp flanking 5’- and 3’-ends. ( E ) Fractions of qDRIP peaks overlapping noted genomic features ( P = 2.5e–2798, chi-square test). ( F ) Scaled metaplot of sense hybrids between TSS and first-intron exon boundary, as well as 1 kb upstream of TSS and 1 kb downstream of first intron-exon boundary. Tracks shown are mean IP (blue) and pooled input (grey). Bands represent 95% CI of mean read signal.

    Techniques Used:

    R-loop lifetimes. ( A ) Schematic of transcription with and without DRB. ( B ) Ratio of DRB to control signal in RNase H-sensitive peaks, compared to estimated time without transcription. Error bands are 95% CI of the mean. Horizontal dotted line indicates a 2-fold decrease in DRB signal. ( C ) qPCR measurements during a DRB timecourse at regions predicted to be unstable (top) or stable (bottom) by pseudo-timecourse obtained from sequencing data. Error bars represent 95% CI of the mean. In primer name, GB indicates gene body. ( D ) GC content across 500 bp regions with shorter, longer or close to average (NS) lifetimes ( P  = 2.5e–143, Kruskal–Wallis test). ( E ) Biochemically determined G-quadruplex counts (  37 ) over the same regions as (D) ( P  = 2.7e–7, ANOVA on Negative Binomial regression, likelihood ratio test). ( F ) Relative replication fork directionality (RFD) (  39 ) to transcription over the same regions as (D), where 1 represents fully co-directional and –1 represents fully head-on ( P  = 3.5e–12, Kruskal–Wallis test). ( G ) Distribution of half-lives assuming first-order decay.
    Figure Legend Snippet: R-loop lifetimes. ( A ) Schematic of transcription with and without DRB. ( B ) Ratio of DRB to control signal in RNase H-sensitive peaks, compared to estimated time without transcription. Error bands are 95% CI of the mean. Horizontal dotted line indicates a 2-fold decrease in DRB signal. ( C ) qPCR measurements during a DRB timecourse at regions predicted to be unstable (top) or stable (bottom) by pseudo-timecourse obtained from sequencing data. Error bars represent 95% CI of the mean. In primer name, GB indicates gene body. ( D ) GC content across 500 bp regions with shorter, longer or close to average (NS) lifetimes ( P = 2.5e–143, Kruskal–Wallis test). ( E ) Biochemically determined G-quadruplex counts ( 37 ) over the same regions as (D) ( P = 2.7e–7, ANOVA on Negative Binomial regression, likelihood ratio test). ( F ) Relative replication fork directionality (RFD) ( 39 ) to transcription over the same regions as (D), where 1 represents fully co-directional and –1 represents fully head-on ( P = 3.5e–12, Kruskal–Wallis test). ( G ) Distribution of half-lives assuming first-order decay.

    Techniques Used: Real-time Polymerase Chain Reaction, Sequencing

    RNase H resistant signal. ( A ) Aggregate plot of qDRIP-seq signal around the TSS of top 10,000 expressed genes as determined by GRO-seq ( 36 ). Tracks are IP (blue), RHR (red) and input (grey). Error bands represent 95% CI of mean. ( B ) Heatmaps of mean IP signal, RNase H-resistant signal and GC-skew around top 10,000 promoters ranked by GC-skew immediately (0–500 bp) downstream of the TSS. Correlation coefficient between IP signal and GC-skew was 0.06, whereas correlation coefficient for RHR signal was 0.22 (Spearman's rho). ( C ) GC-skew around RNase H resistant regions within the full (unfiltered) qDRIP-seq peak set. qDRIP peaks (red) compared to regions of equal lengths randomly selected from non-resistant qDRIP-peaks (blue). As before, bands represent 95% CI of mean read signal. ( D ) Same as (C), but showing biochemically determined G-quadruplex density ( 37 ) over these regions. ( E ) RNase H-resistant signal around RH-resistant peak calls. Peaks lying 5’ in genes (which DRB should affect) are in blue, while peaks lying 3’ in genes (which DRB should not affect) are in red. Left panel is RNase H treatment in control cells, while right panel is RNase H treatment in DRB treated cells. As before, error bands represent 95% CI of the mean.
    Figure Legend Snippet: RNase H resistant signal. ( A ) Aggregate plot of qDRIP-seq signal around the TSS of top 10,000 expressed genes as determined by GRO-seq ( 36 ). Tracks are IP (blue), RHR (red) and input (grey). Error bands represent 95% CI of mean. ( B ) Heatmaps of mean IP signal, RNase H-resistant signal and GC-skew around top 10,000 promoters ranked by GC-skew immediately (0–500 bp) downstream of the TSS. Correlation coefficient between IP signal and GC-skew was 0.06, whereas correlation coefficient for RHR signal was 0.22 (Spearman's rho). ( C ) GC-skew around RNase H resistant regions within the full (unfiltered) qDRIP-seq peak set. qDRIP peaks (red) compared to regions of equal lengths randomly selected from non-resistant qDRIP-peaks (blue). As before, bands represent 95% CI of mean read signal. ( D ) Same as (C), but showing biochemically determined G-quadruplex density ( 37 ) over these regions. ( E ) RNase H-resistant signal around RH-resistant peak calls. Peaks lying 5’ in genes (which DRB should affect) are in blue, while peaks lying 3’ in genes (which DRB should not affect) are in red. Left panel is RNase H treatment in control cells, while right panel is RNase H treatment in DRB treated cells. As before, error bands represent 95% CI of the mean.

    Techniques Used:

    11) Product Images from "Harmful DNA:RNA hybrids are formed in cis and in a Rad51-independent manner"

    Article Title: Harmful DNA:RNA hybrids are formed in cis and in a Rad51-independent manner

    Journal: eLife

    doi: 10.7554/eLife.56674

    Detection of co-transcriptional DNA:RNA hybrids in hpr1Δ and rnh1Δ rnh201Δ mutants at the LacZ -containing constructs under the GAL1 or tet promoters. DNA:RNA Immuno-Precipitation (DRIP) with the S9.6 antibody in WT (W303), hpr1Δ (U678.4C) and rnh1Δ rnh201Δ (HRN2.10C) strains in asynchronous cultures treated or not in vitro with RNase H in the GL- LacZ , tetp:LacZ and GL- LacZi constructs turned transcriptionally off (2% glucose or 5 μg/mL doxycycline) or on (2% galactose and in the absence of doxycycline). Average and SEM of three independent experiments are shown *, p≤0.05; **, p≤0.01; ***, p≤0.001 (unpaired Student’s t-test). Detection of co-transcriptional DNA:RNA hybrids.
    Figure Legend Snippet: Detection of co-transcriptional DNA:RNA hybrids in hpr1Δ and rnh1Δ rnh201Δ mutants at the LacZ -containing constructs under the GAL1 or tet promoters. DNA:RNA Immuno-Precipitation (DRIP) with the S9.6 antibody in WT (W303), hpr1Δ (U678.4C) and rnh1Δ rnh201Δ (HRN2.10C) strains in asynchronous cultures treated or not in vitro with RNase H in the GL- LacZ , tetp:LacZ and GL- LacZi constructs turned transcriptionally off (2% glucose or 5 μg/mL doxycycline) or on (2% galactose and in the absence of doxycycline). Average and SEM of three independent experiments are shown *, p≤0.05; **, p≤0.01; ***, p≤0.001 (unpaired Student’s t-test). Detection of co-transcriptional DNA:RNA hybrids.

    Techniques Used: Construct, Immunoprecipitation, In Vitro

    12) Product Images from "Scalable and cost-effective ribonuclease-based rRNA depletion for transcriptomics"

    Article Title: Scalable and cost-effective ribonuclease-based rRNA depletion for transcriptomics

    Journal: bioRxiv

    doi: 10.1101/645895

    Oligo probes can be applied to closely related species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples using an oligo probe library designed for B. dorei across closely related species ( B. uniformis and B. vulgatus ) and distantly related species ( C. aerofaciens and D. longicatena ). b) Scatter plot between probe-to-target sequence similarity and fold enrichment of non-rRNA reads for five species. Probe-to-target sequence similarity was calculated as the average of percentages of base with mismatches in 16S and 23S rRNA alignments.
    Figure Legend Snippet: Oligo probes can be applied to closely related species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples using an oligo probe library designed for B. dorei across closely related species ( B. uniformis and B. vulgatus ) and distantly related species ( C. aerofaciens and D. longicatena ). b) Scatter plot between probe-to-target sequence similarity and fold enrichment of non-rRNA reads for five species. Probe-to-target sequence similarity was calculated as the average of percentages of base with mismatches in 16S and 23S rRNA alignments.

    Techniques Used: Sequencing

    Workflow for bacterial RNase H based rRNA depletion. a) Probes used for depletion can be either designed and chemically synthesized from known rRNA sequences (oligo-based) or generated by PCR from genomic DNA with 5’-phosphorylated forward primers and subsequent lambda exonuclease digestion (amplicon-based). b) Probes are then hybridized to total RNA and the rRNA bound by the ssDNA probes is degraded by RNase H. Finally, all remaining probes are degraded by DNase I or removed by SPRI beads-based size selection, resulting in enriched mRNAs.
    Figure Legend Snippet: Workflow for bacterial RNase H based rRNA depletion. a) Probes used for depletion can be either designed and chemically synthesized from known rRNA sequences (oligo-based) or generated by PCR from genomic DNA with 5’-phosphorylated forward primers and subsequent lambda exonuclease digestion (amplicon-based). b) Probes are then hybridized to total RNA and the rRNA bound by the ssDNA probes is degraded by RNase H. Finally, all remaining probes are degraded by DNase I or removed by SPRI beads-based size selection, resulting in enriched mRNAs.

    Techniques Used: Synthesized, Generated, Polymerase Chain Reaction, Amplification, Selection

    RNase H based rRNA depletion with amplicons. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples with chemically synthesized oligonucleotides (oligo) or amplicon-based ssDNA (amplicon) probes. b, c) Scatter plot (b) and Quantile-Quantile (Q-Q) plot (c) for depleted sample using amplicon probes with 5:1 probe-to-RNA ratio and un-depleted sample.
    Figure Legend Snippet: RNase H based rRNA depletion with amplicons. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples with chemically synthesized oligonucleotides (oligo) or amplicon-based ssDNA (amplicon) probes. b, c) Scatter plot (b) and Quantile-Quantile (Q-Q) plot (c) for depleted sample using amplicon probes with 5:1 probe-to-RNA ratio and un-depleted sample.

    Techniques Used: Synthesized, Amplification

    High-throughput microbial RNA-seq screening of B. dorei on different substrates. a) The 5 short-chain fatty acids (SCFAs) and 14 carbohydrates tested. B. dorei cultures were treated with each substrate at 5 mg/mL final concentration during exponential growth phase. b) The RNase H based method showed efficient and consistent rRNA depletion on pooled RNA samples from different conditions. Three biological replicates were performed for each condition. c) Multidimensional scaling ordination of 20 conditions based on overall transcriptomes. Pairwise Spearman correlations between conditions were calculated using expression profiles and multidimensional scaling was then performed on normalized Spearman correlations to visualize the impact of substrates on bacterial transcriptome.
    Figure Legend Snippet: High-throughput microbial RNA-seq screening of B. dorei on different substrates. a) The 5 short-chain fatty acids (SCFAs) and 14 carbohydrates tested. B. dorei cultures were treated with each substrate at 5 mg/mL final concentration during exponential growth phase. b) The RNase H based method showed efficient and consistent rRNA depletion on pooled RNA samples from different conditions. Three biological replicates were performed for each condition. c) Multidimensional scaling ordination of 20 conditions based on overall transcriptomes. Pairwise Spearman correlations between conditions were calculated using expression profiles and multidimensional scaling was then performed on normalized Spearman correlations to visualize the impact of substrates on bacterial transcriptome.

    Techniques Used: High Throughput Screening Assay, RNA Sequencing Assay, Concentration Assay, Expressing

    Application of RNase H based rRNA depletion with oligos to three diverse gut microbiota species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples under optimized reaction condition across three microbiota species from different phyla. The same RNA samples used for Ribo-Zero depletion were pooled together to yield the RNA used for RNase H reaction. b, c) Consistency in transcriptome between rRNA depleted samples and un-depleted samples in terms of expression correlation (b) and expression distribution (c) . TPM indicates transcripts per million for each CDS. c) Points lie on a straight line in Quantile-Quantile (Q-Q) plots if there is no global shift in the distribution of expression profile between depleted samples and un-depleted samples.
    Figure Legend Snippet: Application of RNase H based rRNA depletion with oligos to three diverse gut microbiota species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples under optimized reaction condition across three microbiota species from different phyla. The same RNA samples used for Ribo-Zero depletion were pooled together to yield the RNA used for RNase H reaction. b, c) Consistency in transcriptome between rRNA depleted samples and un-depleted samples in terms of expression correlation (b) and expression distribution (c) . TPM indicates transcripts per million for each CDS. c) Points lie on a straight line in Quantile-Quantile (Q-Q) plots if there is no global shift in the distribution of expression profile between depleted samples and un-depleted samples.

    Techniques Used: Expressing

    Optimization of RNase H reaction conditions. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples for two RNase H enzymes and various reaction times with probe-to-RNA fixed to 1:1, or b) various probe-to-RNA ratios with reaction time fixed to 30 minutes. The same RNA sample isolated from Bacteroides dorei was split for rRNA depletion across different reaction conditions.
    Figure Legend Snippet: Optimization of RNase H reaction conditions. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples for two RNase H enzymes and various reaction times with probe-to-RNA fixed to 1:1, or b) various probe-to-RNA ratios with reaction time fixed to 30 minutes. The same RNA sample isolated from Bacteroides dorei was split for rRNA depletion across different reaction conditions.

    Techniques Used: Isolation

    13) Product Images from "Modulation of yeast telomerase activity by Cdc13 and Est1 in vitro"

    Article Title: Modulation of yeast telomerase activity by Cdc13 and Est1 in vitro

    Journal: Scientific Reports

    doi: 10.1038/srep34104

    Single-molecule TPM experiments for telomerase inhibition by Cdc13. ( A ) Effect of Cdc13 on telomerase activity of Est2-TAP/Tlc1 RNP. Thirty nM of T21/C20 DNA was incubated with 2.5, 10, 40, or 160 nM of purified Cdc13 at room temperature for 5 min. Est2-TAP/Tlc1 RNP was then added to the reaction mixtures and the telomerase activity was analyzed by primer extension assay. An image of the gel taken by PhosphorImager (top) and the quantification of telomease activity (bottom) are presented. M shows the position of primer T21 DNA as a size maker. In RNase lane, the Est2-TAP/Tlc1 RNP was pretreated with RNase A at 30 °C for 10 min. ( B ) Single-molecule TPM method for monitoring telomerase activity inhibited by Cdc13. Digoxigenin (Dig)-labeled DNA substrates were incubated with 4 nM Est2-TAP/Tlc1 RNP at 30 °C. The reaction products were stopped by Proteinase K and RNase H and then immobilized in glass surface by anti-digoxigenin (anti-Dig) antibody. The extended DNA were then annealed with biotin-labeled oligonucleotide probes and then with streptavidin-coated polystyrene beads for TPM analysis. BM histograms of DNA substrates without telomerase incubation, extended by Est2-TAP/Tlc1 RNP, and incubated with 10, 40, or 160 nM Cdc13 are presented. ( C ) The percentage of BM peaks with values exceeding 15 nm decreases with increasing Cdc13 concentration.
    Figure Legend Snippet: Single-molecule TPM experiments for telomerase inhibition by Cdc13. ( A ) Effect of Cdc13 on telomerase activity of Est2-TAP/Tlc1 RNP. Thirty nM of T21/C20 DNA was incubated with 2.5, 10, 40, or 160 nM of purified Cdc13 at room temperature for 5 min. Est2-TAP/Tlc1 RNP was then added to the reaction mixtures and the telomerase activity was analyzed by primer extension assay. An image of the gel taken by PhosphorImager (top) and the quantification of telomease activity (bottom) are presented. M shows the position of primer T21 DNA as a size maker. In RNase lane, the Est2-TAP/Tlc1 RNP was pretreated with RNase A at 30 °C for 10 min. ( B ) Single-molecule TPM method for monitoring telomerase activity inhibited by Cdc13. Digoxigenin (Dig)-labeled DNA substrates were incubated with 4 nM Est2-TAP/Tlc1 RNP at 30 °C. The reaction products were stopped by Proteinase K and RNase H and then immobilized in glass surface by anti-digoxigenin (anti-Dig) antibody. The extended DNA were then annealed with biotin-labeled oligonucleotide probes and then with streptavidin-coated polystyrene beads for TPM analysis. BM histograms of DNA substrates without telomerase incubation, extended by Est2-TAP/Tlc1 RNP, and incubated with 10, 40, or 160 nM Cdc13 are presented. ( C ) The percentage of BM peaks with values exceeding 15 nm decreases with increasing Cdc13 concentration.

    Techniques Used: Inhibition, Activity Assay, Incubation, Purification, Primer Extension Assay, Labeling, Concentration Assay

    14) Product Images from "R-loops Associated with Triplet Repeat Expansions Promote Gene Silencing in Friedreich Ataxia and Fragile X Syndrome"

    Article Title: R-loops Associated with Triplet Repeat Expansions Promote Gene Silencing in Friedreich Ataxia and Fragile X Syndrome

    Journal: PLoS Genetics

    doi: 10.1371/journal.pgen.1004318

    R-loops are formed over (CGG) n expanded repeats of FMR1 gene. A. Diagram of FMR1 gene. Black boxes are exons, white box is 5′ UTR and lines are introns. Red triangle is (CGG) n expansion. qPCR amplicons are shown below the diagram. TSS is the transcriptional start site. Numbers indicate the distances to TSS in kilobases. B. RT-qPCR analysis of FMR1 mRNA in control and FXS cells, treated with 1 µM 5-azadC for 7 days, normalized to GAPDH. C. DIP analysis on endogenous FMR1 gene in control and FXS cells, treated with 1 µM 5-azadC for 7 days. Values are relative to ex1 region in control untreated cells. D. FMR1 R-loops are sensitive to RNase H digestion, following the treatment with 25 U of RNase H for 6 hours at 37°C prior to immuno-precipitation. Values are relative to in15 region in control untreated cells. E. R-loop kinetics on exon 1 of FMR1 gene in control and FXS cells during the process of transcriptional re-activation with 1 µM 5-azadC (7 days) followed by 5-azadC wash out with drug-free media (28 days). Values are relative to ex1 region in control untreated cells on day 7. F. RT-qPCR analysis of FMR1 mRNA in control and FXS cells, treated with 1 µM 5-azadC (7 days) followed by 5-azadC wash out with drug-free media (28 days). The level of FMR1 mRNA in control cells is taken as 1. Bars in B–D are average values +/− SEM (n > 3).
    Figure Legend Snippet: R-loops are formed over (CGG) n expanded repeats of FMR1 gene. A. Diagram of FMR1 gene. Black boxes are exons, white box is 5′ UTR and lines are introns. Red triangle is (CGG) n expansion. qPCR amplicons are shown below the diagram. TSS is the transcriptional start site. Numbers indicate the distances to TSS in kilobases. B. RT-qPCR analysis of FMR1 mRNA in control and FXS cells, treated with 1 µM 5-azadC for 7 days, normalized to GAPDH. C. DIP analysis on endogenous FMR1 gene in control and FXS cells, treated with 1 µM 5-azadC for 7 days. Values are relative to ex1 region in control untreated cells. D. FMR1 R-loops are sensitive to RNase H digestion, following the treatment with 25 U of RNase H for 6 hours at 37°C prior to immuno-precipitation. Values are relative to in15 region in control untreated cells. E. R-loop kinetics on exon 1 of FMR1 gene in control and FXS cells during the process of transcriptional re-activation with 1 µM 5-azadC (7 days) followed by 5-azadC wash out with drug-free media (28 days). Values are relative to ex1 region in control untreated cells on day 7. F. RT-qPCR analysis of FMR1 mRNA in control and FXS cells, treated with 1 µM 5-azadC (7 days) followed by 5-azadC wash out with drug-free media (28 days). The level of FMR1 mRNA in control cells is taken as 1. Bars in B–D are average values +/− SEM (n > 3).

    Techniques Used: Real-time Polymerase Chain Reaction, Quantitative RT-PCR, Immunoprecipitation, Activation Assay

    R-loops are formed over expanded repeats of FXN gene in FRDA cells. A. Diagram of FXN gene. Black boxes are exons, white boxes are 5′ and 3′UTRs, lines are introns, red triangle is (GAA) n expansion. TSS2 is the major transcriptional start site in lymphoblastoid cells. qPCR amplicons are shown below the diagram. Numbers indicate the distances to TSS2 in kilobases. B. Cell lines used in the study. The repeat sizes are indicated. C. RT-qPCR analysis of γ-actin, β-actin, GAPDH and FXN mRNAs in control (GM15851) and FRDA (GM15850) cells. Values are normalised to 5S rRNA and relative to control cells. D. RNA Pol II ChIP in control (GM15851) and FRDA (GM15850) cells. E. RT-qPCR analysis of FXN nascent RNA in control (GM15851) and FRDA (GM15850) cells, normalised to 5S rRNA and relative to ex1 RNA in control cells. F. DIP on endogenous FXN gene in control (GM15851) and FRDA (GM15851) cells. γ-actin is positive control. G. R-loops are sensitive to RNase H digestion. DIP samples were treated with 25 U of recombinant E.coli RNase H (NEB, M0297S) for 6 hours at 37°C. γ-actin is positive control. Bars in C–G are average values +/− SEM (n > 3).
    Figure Legend Snippet: R-loops are formed over expanded repeats of FXN gene in FRDA cells. A. Diagram of FXN gene. Black boxes are exons, white boxes are 5′ and 3′UTRs, lines are introns, red triangle is (GAA) n expansion. TSS2 is the major transcriptional start site in lymphoblastoid cells. qPCR amplicons are shown below the diagram. Numbers indicate the distances to TSS2 in kilobases. B. Cell lines used in the study. The repeat sizes are indicated. C. RT-qPCR analysis of γ-actin, β-actin, GAPDH and FXN mRNAs in control (GM15851) and FRDA (GM15850) cells. Values are normalised to 5S rRNA and relative to control cells. D. RNA Pol II ChIP in control (GM15851) and FRDA (GM15850) cells. E. RT-qPCR analysis of FXN nascent RNA in control (GM15851) and FRDA (GM15850) cells, normalised to 5S rRNA and relative to ex1 RNA in control cells. F. DIP on endogenous FXN gene in control (GM15851) and FRDA (GM15851) cells. γ-actin is positive control. G. R-loops are sensitive to RNase H digestion. DIP samples were treated with 25 U of recombinant E.coli RNase H (NEB, M0297S) for 6 hours at 37°C. γ-actin is positive control. Bars in C–G are average values +/− SEM (n > 3).

    Techniques Used: Real-time Polymerase Chain Reaction, Quantitative RT-PCR, Chromatin Immunoprecipitation, Positive Control, Recombinant

    15) Product Images from "Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †"

    Article Title: Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †

    Journal:

    doi: 10.1128/MCB.01410-07

    RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)
    Figure Legend Snippet: RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)

    Techniques Used:

    16) Product Images from "Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †"

    Article Title: Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †

    Journal:

    doi: 10.1128/MCB.01410-07

    RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)
    Figure Legend Snippet: RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)

    Techniques Used:

    17) Product Images from "Isolation and genome-wide characterization of cellular DNA:RNA triplex structures"

    Article Title: Isolation and genome-wide characterization of cellular DNA:RNA triplex structures

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gky1305

    NEAT1 forms triplexes at numerous genomic sites. ( A ) NEAT1 profiles in TriplexRNA-seq (DNA-IP) (red) and nuclear RNA (blue) from HeLa S3 and U2OS cells with shaded TFR1 and TFR2. Minus (-) and plus (+) strands are shown. The position and sequence of NEAT1-TFR1 and -TFR2 are shown below. ( B ) EMSAs using 10 or 100 pmol of synthetic NEAT1 versions comprising TFR1 (40 or 52 nt) or TFR2 incubated with 0.25 pmol of double–stranded  32 P-labeled oligonucleotides which harbor sequences of NEAT1 target genes predicted from CHART-seq (  Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control, RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( C ) Schematic depiction of the TFR-based capture assay. Biotinylated RNA oligos covering NEAT1-TFR1 and NEAT1-TFR2 were used to capture genomic DNA. ( D ) MEME motif analysis identifying consensus motifs in DNA captured by NEAT1-TFR1 (399 of top 500 peaks) and by NEAT1-TFR2 (500 of top 500 peaks ranked by peak  P -value). Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( E ) TDF analysis of the triplex-forming potential of NEAT1-TFR1 and NEAT1-TFR2 RNAs with top 500 TFR-associated and control DNA peaks (ranked by peak  P -value) compared to 500 randomized regions ( N  = 1000, colored grey).  P -values were obtained from one-tailed Mann–Whitney test. ( F ) Scheme presenting antisense oligo (ASO)-based capture of NEAT1-associated DNA. ( G ) Consensus motif in NEAT1-associated DNA sites (314 of top 500 peaks ranked by peak  P -value). ( H ) TDF analysis predicting the triplex-forming potential of NEAT1 on ASO-captured DNA regions. Significant TFRs along NEAT1 are shown in orange, the number of target sites (DBS) for each TFR in purple. For TFR- and ASO-based capture assays nucleic acids isolated from HeLa S3 chromatin were used.
    Figure Legend Snippet: NEAT1 forms triplexes at numerous genomic sites. ( A ) NEAT1 profiles in TriplexRNA-seq (DNA-IP) (red) and nuclear RNA (blue) from HeLa S3 and U2OS cells with shaded TFR1 and TFR2. Minus (-) and plus (+) strands are shown. The position and sequence of NEAT1-TFR1 and -TFR2 are shown below. ( B ) EMSAs using 10 or 100 pmol of synthetic NEAT1 versions comprising TFR1 (40 or 52 nt) or TFR2 incubated with 0.25 pmol of double–stranded 32 P-labeled oligonucleotides which harbor sequences of NEAT1 target genes predicted from CHART-seq ( Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control, RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( C ) Schematic depiction of the TFR-based capture assay. Biotinylated RNA oligos covering NEAT1-TFR1 and NEAT1-TFR2 were used to capture genomic DNA. ( D ) MEME motif analysis identifying consensus motifs in DNA captured by NEAT1-TFR1 (399 of top 500 peaks) and by NEAT1-TFR2 (500 of top 500 peaks ranked by peak P -value). Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( E ) TDF analysis of the triplex-forming potential of NEAT1-TFR1 and NEAT1-TFR2 RNAs with top 500 TFR-associated and control DNA peaks (ranked by peak P -value) compared to 500 randomized regions ( N = 1000, colored grey). P -values were obtained from one-tailed Mann–Whitney test. ( F ) Scheme presenting antisense oligo (ASO)-based capture of NEAT1-associated DNA. ( G ) Consensus motif in NEAT1-associated DNA sites (314 of top 500 peaks ranked by peak P -value). ( H ) TDF analysis predicting the triplex-forming potential of NEAT1 on ASO-captured DNA regions. Significant TFRs along NEAT1 are shown in orange, the number of target sites (DBS) for each TFR in purple. For TFR- and ASO-based capture assays nucleic acids isolated from HeLa S3 chromatin were used.

    Techniques Used: Sequencing, Incubation, Labeling, One-tailed Test, MANN-WHITNEY, Allele-specific Oligonucleotide, Isolation

    Validation of triplex-forming RNA and DNAs. ( A ) TDF analysis predicting the potential of top 1000 enriched TriplexRNA (DNA-IP) regions (ranked by peak  P -value) to bind to active promoters defined by ChromHMM. Number of TFRs in RNA (per kilobase of RNA, left) and the number of putative DBSs at promoters (per kilobase of RNA, right) are shown. Boxplot borders are defined by the 1st and 3rd quantiles of the distributions, the middle line corresponds to the median value. The top whisker denotes the maximum value within the third quartile plus 1.5 times the interquartile range (bottom whisker is defined analogously). Dark gray dots represent outliers with values higher or lower than whiskers. Further box plots are based on the same definitions. ( B ) Motif analysis of triplexes formed between TriplexRNA (DNA-IP) and active promoters. The diagram depicts the fraction of antiparallel and parallel triplexes with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( C ) TDF analysis comparing the triplex-forming potential of top 2000 TriplexDNA-seq regions with top 1000 TriplexRNA (DNA-IP) (ranked by peak  P -value). The number of putative DBSs (per kilobase of RNA) is shown. ( D ) Motif analysis of predicted triplexes formed between TriplexRNAs (DNA-IP) and TriplexDNA. The diagram depicts the fraction of antiparallel and parallel triplexes, with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( E ) Box plot classifying triplex interactions between TriplexRNAs (DNA-IP) and TriplexDNA-seq regions as  cis  ( > 10 kb in the same chromosome) and  trans  (at different chromosomes) interactions, excluding underrepresented local interactions (within 10 kb distance). ( F ) EMSAs using 10 or 100 pmol of synthetic TriplexRNAs and 0.25 pmol of double–stranded  32 P-labeled oligonucleotides comprising target regions from TriplexDNA (  Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control (C), RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). TriplexRNA-seq and TriplexDNA-seq data are from HeLa S3 cells. Adjusted  P -values
    Figure Legend Snippet: Validation of triplex-forming RNA and DNAs. ( A ) TDF analysis predicting the potential of top 1000 enriched TriplexRNA (DNA-IP) regions (ranked by peak P -value) to bind to active promoters defined by ChromHMM. Number of TFRs in RNA (per kilobase of RNA, left) and the number of putative DBSs at promoters (per kilobase of RNA, right) are shown. Boxplot borders are defined by the 1st and 3rd quantiles of the distributions, the middle line corresponds to the median value. The top whisker denotes the maximum value within the third quartile plus 1.5 times the interquartile range (bottom whisker is defined analogously). Dark gray dots represent outliers with values higher or lower than whiskers. Further box plots are based on the same definitions. ( B ) Motif analysis of triplexes formed between TriplexRNA (DNA-IP) and active promoters. The diagram depicts the fraction of antiparallel and parallel triplexes with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( C ) TDF analysis comparing the triplex-forming potential of top 2000 TriplexDNA-seq regions with top 1000 TriplexRNA (DNA-IP) (ranked by peak P -value). The number of putative DBSs (per kilobase of RNA) is shown. ( D ) Motif analysis of predicted triplexes formed between TriplexRNAs (DNA-IP) and TriplexDNA. The diagram depicts the fraction of antiparallel and parallel triplexes, with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( E ) Box plot classifying triplex interactions between TriplexRNAs (DNA-IP) and TriplexDNA-seq regions as cis ( > 10 kb in the same chromosome) and trans (at different chromosomes) interactions, excluding underrepresented local interactions (within 10 kb distance). ( F ) EMSAs using 10 or 100 pmol of synthetic TriplexRNAs and 0.25 pmol of double–stranded 32 P-labeled oligonucleotides comprising target regions from TriplexDNA ( Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control (C), RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). TriplexRNA-seq and TriplexDNA-seq data are from HeLa S3 cells. Adjusted P -values

    Techniques Used: Whisker Assay, Labeling

    18) Product Images from "Improvement of RNA secondary structure prediction using RNase H cleavage and randomized oligonucleotides"

    Article Title: Improvement of RNA secondary structure prediction using RNase H cleavage and randomized oligonucleotides

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkp587

    Phylogenetic secondary structure, the predicted lowest free energy secondary structure, and the four suboptimal structures of E. coli 5S rRNA. Loops A–E are labeled in the phylogenetic structure. Base-paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H cleavage are circled. The lowest free energy structure only has 27% of the base pairs present in the phylogenetic structure.
    Figure Legend Snippet: Phylogenetic secondary structure, the predicted lowest free energy secondary structure, and the four suboptimal structures of E. coli 5S rRNA. Loops A–E are labeled in the phylogenetic structure. Base-paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H cleavage are circled. The lowest free energy structure only has 27% of the base pairs present in the phylogenetic structure.

    Techniques Used: Labeling

    Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in yeast tRNA Phe with randomized 5-mer oligonucleotides.
    Figure Legend Snippet: Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in yeast tRNA Phe with randomized 5-mer oligonucleotides.

    Techniques Used:

    Schematic of the general method used in this study. Randomized DNA oligonucleotides are incubated with an RNA of interest. Only DNAs complementary to single-stranded regions bind, inducing RNase H cleavage of the RNA strand. Nucleotides which are subject to RNase H cleavage are used as single-stranded constraints in RNA secondary structure prediction.
    Figure Legend Snippet: Schematic of the general method used in this study. Randomized DNA oligonucleotides are incubated with an RNA of interest. Only DNAs complementary to single-stranded regions bind, inducing RNase H cleavage of the RNA strand. Nucleotides which are subject to RNase H cleavage are used as single-stranded constraints in RNA secondary structure prediction.

    Techniques Used: Incubation

    Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in E. coli 5S rRNA. The numbers above the lanes indicate to which nucleotides in the RNA the oligonucleotide probe is complementary.
    Figure Legend Snippet: Representative gel autoradiogram of RNase H cleavage experiments to identify single-stranded regions in E. coli 5S rRNA. The numbers above the lanes indicate to which nucleotides in the RNA the oligonucleotide probe is complementary.

    Techniques Used:

    Phylogenetic secondary structure and three predicted suboptimal structures of yeast tRNA Phe . The lowest free energy structure has 95% of the base pairs predicted correctly. Base paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H after 1 or 2 h incubation are circled. D denotes dihydrouracil while Y denotes wybutosine.
    Figure Legend Snippet: Phylogenetic secondary structure and three predicted suboptimal structures of yeast tRNA Phe . The lowest free energy structure has 95% of the base pairs predicted correctly. Base paired regions that are predicted correctly in the suboptimal structures are shaded. Nucleotides cleaved by RNase H after 1 or 2 h incubation are circled. D denotes dihydrouracil while Y denotes wybutosine.

    Techniques Used: Incubation

    19) Product Images from "Formation and Repair of Mismatches Containing Ribonucleotides and Oxidized Bases at Repeated DNA Sequences *"

    Article Title: Formation and Repair of Mismatches Containing Ribonucleotides and Oxidized Bases at Repeated DNA Sequences *

    Journal: The Journal of Biological Chemistry

    doi: 10.1074/jbc.M115.679209

    BER and RER activity on complex mispairs containing oxidized bases and ribonucleotides. When POL β incorporates rCMP opposite 8-oxodG (dG*), RNase H2 is going to efficiently remove rC from the resulting 8-oxodG:rC mispair, whereas OGG1 repair of dG* is slightly reduced. Should 8-oxodG:rA arise after rAMP incorporation, RER will process the rA containing strand, whereas MUTYH-mediated BER will be inhibited. Possible interference on RER activity might occur by concurrent recognition of the lesion. In the likelihood of limiting MTH1 hydrolytic activity, 8-oxorGTP (rG*TP) might be used by POL β to produce rG*:dA mispairs. These substrates will be efficiently processed by MUTYH and RNase H2. Simultaneous BER and RER activities might lead to the formation of double strand breaks ( DSB ) or intermediate repair products of unknown reparability.
    Figure Legend Snippet: BER and RER activity on complex mispairs containing oxidized bases and ribonucleotides. When POL β incorporates rCMP opposite 8-oxodG (dG*), RNase H2 is going to efficiently remove rC from the resulting 8-oxodG:rC mispair, whereas OGG1 repair of dG* is slightly reduced. Should 8-oxodG:rA arise after rAMP incorporation, RER will process the rA containing strand, whereas MUTYH-mediated BER will be inhibited. Possible interference on RER activity might occur by concurrent recognition of the lesion. In the likelihood of limiting MTH1 hydrolytic activity, 8-oxorGTP (rG*TP) might be used by POL β to produce rG*:dA mispairs. These substrates will be efficiently processed by MUTYH and RNase H2. Simultaneous BER and RER activities might lead to the formation of double strand breaks ( DSB ) or intermediate repair products of unknown reparability.

    Techniques Used: Activity Assay

    20) Product Images from "Formation and Repair of Mismatches Containing Ribonucleotides and Oxidized Bases at Repeated DNA Sequences *"

    Article Title: Formation and Repair of Mismatches Containing Ribonucleotides and Oxidized Bases at Repeated DNA Sequences *

    Journal: The Journal of Biological Chemistry

    doi: 10.1074/jbc.M115.679209

    BER and RER activity on complex mispairs containing oxidized bases and ribonucleotides. When POL β incorporates rCMP opposite 8-oxodG (dG*), RNase H2 is going to efficiently remove rC from the resulting 8-oxodG:rC mispair, whereas OGG1 repair of dG* is slightly reduced. Should 8-oxodG:rA arise after rAMP incorporation, RER will process the rA containing strand, whereas MUTYH-mediated BER will be inhibited. Possible interference on RER activity might occur by concurrent recognition of the lesion. In the likelihood of limiting MTH1 hydrolytic activity, 8-oxorGTP (rG*TP) might be used by POL β to produce rG*:dA mispairs. These substrates will be efficiently processed by MUTYH and RNase H2. Simultaneous BER and RER activities might lead to the formation of double strand breaks ( DSB ) or intermediate repair products of unknown reparability.
    Figure Legend Snippet: BER and RER activity on complex mispairs containing oxidized bases and ribonucleotides. When POL β incorporates rCMP opposite 8-oxodG (dG*), RNase H2 is going to efficiently remove rC from the resulting 8-oxodG:rC mispair, whereas OGG1 repair of dG* is slightly reduced. Should 8-oxodG:rA arise after rAMP incorporation, RER will process the rA containing strand, whereas MUTYH-mediated BER will be inhibited. Possible interference on RER activity might occur by concurrent recognition of the lesion. In the likelihood of limiting MTH1 hydrolytic activity, 8-oxorGTP (rG*TP) might be used by POL β to produce rG*:dA mispairs. These substrates will be efficiently processed by MUTYH and RNase H2. Simultaneous BER and RER activities might lead to the formation of double strand breaks ( DSB ) or intermediate repair products of unknown reparability.

    Techniques Used: Activity Assay

    21) Product Images from "Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter"

    Article Title: Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkx403

    Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.
    Figure Legend Snippet: Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.

    Techniques Used: Sequencing, Quantitation Assay

    Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).
    Figure Legend Snippet: Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).

    Techniques Used: Sequencing

    22) Product Images from "Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter"

    Article Title: Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkx403

    Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.
    Figure Legend Snippet: Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.

    Techniques Used: Sequencing, Quantitation Assay

    Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).
    Figure Legend Snippet: Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).

    Techniques Used: Sequencing

    23) Product Images from "SAC3B, a central component of the mRNA export complex TREX-2, is required for prevention of epigenetic gene silencing in Arabidopsis"

    Article Title: SAC3B, a central component of the mRNA export complex TREX-2, is required for prevention of epigenetic gene silencing in Arabidopsis

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkw850

    Gene silencing in sac3b mutants is independent of R-loop accumulation. ( A ) A snapshot of RNAseq data showing FLC expression is decreased in the sac3b-7 mutant. ( B ) Reduced FLC gene expression in p31 and sac3b-7 mutants by RT-qPCR. ( C ) COOLAIR transcript levels in sac3b mutant alleles; RT-qPCR results are shown. Error bars are SEM, n = 3 biological replicates. ( D ) Schematic of d35S::LUC and FLC gene structure showing the examined subregions (I, II, III, FLC-h, FLC-e ) in DNA IP assays. ( E ) R-loop detection by DNA IP followed by qPCR. The S9.6 antibody was used for DNA IP. Values of R-loop enrichment were divided by input and normalized to FLC -h region. Error bars indicate SD from three technical repeats. Results were confirmed by three biological replicates. R-loop signals were confirmed by RNase H treatment that specifically degrades RNA within DNA:RNA hybrid. NoAb: no antibody control.
    Figure Legend Snippet: Gene silencing in sac3b mutants is independent of R-loop accumulation. ( A ) A snapshot of RNAseq data showing FLC expression is decreased in the sac3b-7 mutant. ( B ) Reduced FLC gene expression in p31 and sac3b-7 mutants by RT-qPCR. ( C ) COOLAIR transcript levels in sac3b mutant alleles; RT-qPCR results are shown. Error bars are SEM, n = 3 biological replicates. ( D ) Schematic of d35S::LUC and FLC gene structure showing the examined subregions (I, II, III, FLC-h, FLC-e ) in DNA IP assays. ( E ) R-loop detection by DNA IP followed by qPCR. The S9.6 antibody was used for DNA IP. Values of R-loop enrichment were divided by input and normalized to FLC -h region. Error bars indicate SD from three technical repeats. Results were confirmed by three biological replicates. R-loop signals were confirmed by RNase H treatment that specifically degrades RNA within DNA:RNA hybrid. NoAb: no antibody control.

    Techniques Used: Expressing, Mutagenesis, Quantitative RT-PCR, Real-time Polymerase Chain Reaction

    24) Product Images from "Scalable and cost-effective ribonuclease-based rRNA depletion for transcriptomics"

    Article Title: Scalable and cost-effective ribonuclease-based rRNA depletion for transcriptomics

    Journal: bioRxiv

    doi: 10.1101/645895

    Oligo probes can be applied to closely related species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples using an oligo probe library designed for B. dorei across closely related species ( B. uniformis and B. vulgatus ) and distantly related species ( C. aerofaciens and D. longicatena ). b) Scatter plot between probe-to-target sequence similarity and fold enrichment of non-rRNA reads for five species. Probe-to-target sequence similarity was calculated as the average of percentages of base with mismatches in 16S and 23S rRNA alignments.
    Figure Legend Snippet: Oligo probes can be applied to closely related species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples using an oligo probe library designed for B. dorei across closely related species ( B. uniformis and B. vulgatus ) and distantly related species ( C. aerofaciens and D. longicatena ). b) Scatter plot between probe-to-target sequence similarity and fold enrichment of non-rRNA reads for five species. Probe-to-target sequence similarity was calculated as the average of percentages of base with mismatches in 16S and 23S rRNA alignments.

    Techniques Used: Sequencing

    Workflow for bacterial RNase H based rRNA depletion. a) Probes used for depletion can be either designed and chemically synthesized from known rRNA sequences (oligo-based) or generated by PCR from genomic DNA with 5’-phosphorylated forward primers and subsequent lambda exonuclease digestion (amplicon-based). b) Probes are then hybridized to total RNA and the rRNA bound by the ssDNA probes is degraded by RNase H. Finally, all remaining probes are degraded by DNase I or removed by SPRI beads-based size selection, resulting in enriched mRNAs.
    Figure Legend Snippet: Workflow for bacterial RNase H based rRNA depletion. a) Probes used for depletion can be either designed and chemically synthesized from known rRNA sequences (oligo-based) or generated by PCR from genomic DNA with 5’-phosphorylated forward primers and subsequent lambda exonuclease digestion (amplicon-based). b) Probes are then hybridized to total RNA and the rRNA bound by the ssDNA probes is degraded by RNase H. Finally, all remaining probes are degraded by DNase I or removed by SPRI beads-based size selection, resulting in enriched mRNAs.

    Techniques Used: Synthesized, Generated, Polymerase Chain Reaction, Amplification, Selection

    RNase H based rRNA depletion with amplicons. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples with chemically synthesized oligonucleotides (oligo) or amplicon-based ssDNA (amplicon) probes. b, c) Scatter plot (b) and Quantile-Quantile (Q-Q) plot (c) for depleted sample using amplicon probes with 5:1 probe-to-RNA ratio and un-depleted sample.
    Figure Legend Snippet: RNase H based rRNA depletion with amplicons. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples with chemically synthesized oligonucleotides (oligo) or amplicon-based ssDNA (amplicon) probes. b, c) Scatter plot (b) and Quantile-Quantile (Q-Q) plot (c) for depleted sample using amplicon probes with 5:1 probe-to-RNA ratio and un-depleted sample.

    Techniques Used: Synthesized, Amplification

    High-throughput microbial RNA-seq screening of B. dorei on different substrates. a) The 5 short-chain fatty acids (SCFAs) and 14 carbohydrates tested. B. dorei cultures were treated with each substrate at 5 mg/mL final concentration during exponential growth phase. b) The RNase H based method showed efficient and consistent rRNA depletion on pooled RNA samples from different conditions. Three biological replicates were performed for each condition. c) Multidimensional scaling ordination of 20 conditions based on overall transcriptomes. Pairwise Spearman correlations between conditions were calculated using expression profiles and multidimensional scaling was then performed on normalized Spearman correlations to visualize the impact of substrates on bacterial transcriptome.
    Figure Legend Snippet: High-throughput microbial RNA-seq screening of B. dorei on different substrates. a) The 5 short-chain fatty acids (SCFAs) and 14 carbohydrates tested. B. dorei cultures were treated with each substrate at 5 mg/mL final concentration during exponential growth phase. b) The RNase H based method showed efficient and consistent rRNA depletion on pooled RNA samples from different conditions. Three biological replicates were performed for each condition. c) Multidimensional scaling ordination of 20 conditions based on overall transcriptomes. Pairwise Spearman correlations between conditions were calculated using expression profiles and multidimensional scaling was then performed on normalized Spearman correlations to visualize the impact of substrates on bacterial transcriptome.

    Techniques Used: High Throughput Screening Assay, RNA Sequencing Assay, Concentration Assay, Expressing

    Application of RNase H based rRNA depletion with oligos to three diverse gut microbiota species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples under optimized reaction condition across three microbiota species from different phyla. The same RNA samples used for Ribo-Zero depletion were pooled together to yield the RNA used for RNase H reaction. b, c) Consistency in transcriptome between rRNA depleted samples and un-depleted samples in terms of expression correlation (b) and expression distribution (c) . TPM indicates transcripts per million for each CDS. c) Points lie on a straight line in Quantile-Quantile (Q-Q) plots if there is no global shift in the distribution of expression profile between depleted samples and un-depleted samples.
    Figure Legend Snippet: Application of RNase H based rRNA depletion with oligos to three diverse gut microbiota species. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted, Ribo-Zero depleted, and RNase H depleted samples under optimized reaction condition across three microbiota species from different phyla. The same RNA samples used for Ribo-Zero depletion were pooled together to yield the RNA used for RNase H reaction. b, c) Consistency in transcriptome between rRNA depleted samples and un-depleted samples in terms of expression correlation (b) and expression distribution (c) . TPM indicates transcripts per million for each CDS. c) Points lie on a straight line in Quantile-Quantile (Q-Q) plots if there is no global shift in the distribution of expression profile between depleted samples and un-depleted samples.

    Techniques Used: Expressing

    Optimization of RNase H reaction conditions. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples for two RNase H enzymes and various reaction times with probe-to-RNA fixed to 1:1, or b) various probe-to-RNA ratios with reaction time fixed to 30 minutes. The same RNA sample isolated from Bacteroides dorei was split for rRNA depletion across different reaction conditions.
    Figure Legend Snippet: Optimization of RNase H reaction conditions. a) Proportion of rRNA-aligning reads of total mapped reads (% rRNA reads), for un-depleted and RNase H depleted samples for two RNase H enzymes and various reaction times with probe-to-RNA fixed to 1:1, or b) various probe-to-RNA ratios with reaction time fixed to 30 minutes. The same RNA sample isolated from Bacteroides dorei was split for rRNA depletion across different reaction conditions.

    Techniques Used: Isolation

    25) Product Images from "RNase H1 directs origin-specific initiation of DNA replication in human mitochondria"

    Article Title: RNase H1 directs origin-specific initiation of DNA replication in human mitochondria

    Journal: PLoS Genetics

    doi: 10.1371/journal.pgen.1007781

    DNA replication initiation defects in RNase H1 deficient cells. details). Control cell DNA (WT) in lanes 1-2 and RNase H1 patient cell DNA in lanes 3-4. Untreated DNA in lanes 1 and 3, and RNase H2 treated DNA in lanes 2 and 4. The NEB LMW ladder is indicated in black, mtDNA positions in green, mapped 5′-ends in blue and mapped RNA to DNA transition points in red. G, C, A and T sequencing ladders are found on the left-hand side. A schematic representation of the control region with OriH, CSBI-III and LSP is shown on the right-hand side. C. Primer extension of mtDNA from control and RNase H1 patient cells with primer 2 (corresponding to mtDNA positions 16,231-16,251, see panel A). Sample loading and colored indications as in panel B. D. 5′-end sequencing (5′-End-seq) of control cell mtDNA. E. Hydrolytic end sequencing (HydEn-seq) of control cell mtDNA to map 5′-ends with attached ribonucleotides. F. 5′-end sequencing (5′-End-seq) of RNase H1 patient cell mtDNA. G. Hydrolytic end sequencing (HydEn-seq) of RNase H1 patient cell mtDNA to map 5′-ends with attached ribonucleotides.
    Figure Legend Snippet: DNA replication initiation defects in RNase H1 deficient cells. details). Control cell DNA (WT) in lanes 1-2 and RNase H1 patient cell DNA in lanes 3-4. Untreated DNA in lanes 1 and 3, and RNase H2 treated DNA in lanes 2 and 4. The NEB LMW ladder is indicated in black, mtDNA positions in green, mapped 5′-ends in blue and mapped RNA to DNA transition points in red. G, C, A and T sequencing ladders are found on the left-hand side. A schematic representation of the control region with OriH, CSBI-III and LSP is shown on the right-hand side. C. Primer extension of mtDNA from control and RNase H1 patient cells with primer 2 (corresponding to mtDNA positions 16,231-16,251, see panel A). Sample loading and colored indications as in panel B. D. 5′-end sequencing (5′-End-seq) of control cell mtDNA. E. Hydrolytic end sequencing (HydEn-seq) of control cell mtDNA to map 5′-ends with attached ribonucleotides. F. 5′-end sequencing (5′-End-seq) of RNase H1 patient cell mtDNA. G. Hydrolytic end sequencing (HydEn-seq) of RNase H1 patient cell mtDNA to map 5′-ends with attached ribonucleotides.

    Techniques Used: Sequencing

    26) Product Images from "Isolation and genome-wide characterization of cellular DNA:RNA triplex structures"

    Article Title: Isolation and genome-wide characterization of cellular DNA:RNA triplex structures

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gky1305

    NEAT1 forms triplexes at numerous genomic sites. ( A ) NEAT1 profiles in TriplexRNA-seq (DNA-IP) (red) and nuclear RNA (blue) from HeLa S3 and U2OS cells with shaded TFR1 and TFR2. Minus (-) and plus (+) strands are shown. The position and sequence of NEAT1-TFR1 and -TFR2 are shown below. ( B ) EMSAs using 10 or 100 pmol of synthetic NEAT1 versions comprising TFR1 (40 or 52 nt) or TFR2 incubated with 0.25 pmol of double–stranded  32 P-labeled oligonucleotides which harbor sequences of NEAT1 target genes predicted from CHART-seq (  Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control, RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( C ) Schematic depiction of the TFR-based capture assay. Biotinylated RNA oligos covering NEAT1-TFR1 and NEAT1-TFR2 were used to capture genomic DNA. ( D ) MEME motif analysis identifying consensus motifs in DNA captured by NEAT1-TFR1 (399 of top 500 peaks) and by NEAT1-TFR2 (500 of top 500 peaks ranked by peak  P -value). Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( E ) TDF analysis of the triplex-forming potential of NEAT1-TFR1 and NEAT1-TFR2 RNAs with top 500 TFR-associated and control DNA peaks (ranked by peak  P -value) compared to 500 randomized regions ( N  = 1000, colored grey).  P -values were obtained from one-tailed Mann–Whitney test. ( F ) Scheme presenting antisense oligo (ASO)-based capture of NEAT1-associated DNA. ( G ) Consensus motif in NEAT1-associated DNA sites (314 of top 500 peaks ranked by peak  P -value). ( H ) TDF analysis predicting the triplex-forming potential of NEAT1 on ASO-captured DNA regions. Significant TFRs along NEAT1 are shown in orange, the number of target sites (DBS) for each TFR in purple. For TFR- and ASO-based capture assays nucleic acids isolated from HeLa S3 chromatin were used.
    Figure Legend Snippet: NEAT1 forms triplexes at numerous genomic sites. ( A ) NEAT1 profiles in TriplexRNA-seq (DNA-IP) (red) and nuclear RNA (blue) from HeLa S3 and U2OS cells with shaded TFR1 and TFR2. Minus (-) and plus (+) strands are shown. The position and sequence of NEAT1-TFR1 and -TFR2 are shown below. ( B ) EMSAs using 10 or 100 pmol of synthetic NEAT1 versions comprising TFR1 (40 or 52 nt) or TFR2 incubated with 0.25 pmol of double–stranded 32 P-labeled oligonucleotides which harbor sequences of NEAT1 target genes predicted from CHART-seq ( Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control, RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( C ) Schematic depiction of the TFR-based capture assay. Biotinylated RNA oligos covering NEAT1-TFR1 and NEAT1-TFR2 were used to capture genomic DNA. ( D ) MEME motif analysis identifying consensus motifs in DNA captured by NEAT1-TFR1 (399 of top 500 peaks) and by NEAT1-TFR2 (500 of top 500 peaks ranked by peak P -value). Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). ( E ) TDF analysis of the triplex-forming potential of NEAT1-TFR1 and NEAT1-TFR2 RNAs with top 500 TFR-associated and control DNA peaks (ranked by peak P -value) compared to 500 randomized regions ( N = 1000, colored grey). P -values were obtained from one-tailed Mann–Whitney test. ( F ) Scheme presenting antisense oligo (ASO)-based capture of NEAT1-associated DNA. ( G ) Consensus motif in NEAT1-associated DNA sites (314 of top 500 peaks ranked by peak P -value). ( H ) TDF analysis predicting the triplex-forming potential of NEAT1 on ASO-captured DNA regions. Significant TFRs along NEAT1 are shown in orange, the number of target sites (DBS) for each TFR in purple. For TFR- and ASO-based capture assays nucleic acids isolated from HeLa S3 chromatin were used.

    Techniques Used: Sequencing, Incubation, Labeling, One-tailed Test, MANN-WHITNEY, Allele-specific Oligonucleotide, Isolation

    Validation of triplex-forming RNA and DNAs. ( A ) TDF analysis predicting the potential of top 1000 enriched TriplexRNA (DNA-IP) regions (ranked by peak  P -value) to bind to active promoters defined by ChromHMM. Number of TFRs in RNA (per kilobase of RNA, left) and the number of putative DBSs at promoters (per kilobase of RNA, right) are shown. Boxplot borders are defined by the 1st and 3rd quantiles of the distributions, the middle line corresponds to the median value. The top whisker denotes the maximum value within the third quartile plus 1.5 times the interquartile range (bottom whisker is defined analogously). Dark gray dots represent outliers with values higher or lower than whiskers. Further box plots are based on the same definitions. ( B ) Motif analysis of triplexes formed between TriplexRNA (DNA-IP) and active promoters. The diagram depicts the fraction of antiparallel and parallel triplexes with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( C ) TDF analysis comparing the triplex-forming potential of top 2000 TriplexDNA-seq regions with top 1000 TriplexRNA (DNA-IP) (ranked by peak  P -value). The number of putative DBSs (per kilobase of RNA) is shown. ( D ) Motif analysis of predicted triplexes formed between TriplexRNAs (DNA-IP) and TriplexDNA. The diagram depicts the fraction of antiparallel and parallel triplexes, with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( E ) Box plot classifying triplex interactions between TriplexRNAs (DNA-IP) and TriplexDNA-seq regions as  cis  ( > 10 kb in the same chromosome) and  trans  (at different chromosomes) interactions, excluding underrepresented local interactions (within 10 kb distance). ( F ) EMSAs using 10 or 100 pmol of synthetic TriplexRNAs and 0.25 pmol of double–stranded  32 P-labeled oligonucleotides comprising target regions from TriplexDNA (  Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control (C), RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). TriplexRNA-seq and TriplexDNA-seq data are from HeLa S3 cells. Adjusted  P -values
    Figure Legend Snippet: Validation of triplex-forming RNA and DNAs. ( A ) TDF analysis predicting the potential of top 1000 enriched TriplexRNA (DNA-IP) regions (ranked by peak P -value) to bind to active promoters defined by ChromHMM. Number of TFRs in RNA (per kilobase of RNA, left) and the number of putative DBSs at promoters (per kilobase of RNA, right) are shown. Boxplot borders are defined by the 1st and 3rd quantiles of the distributions, the middle line corresponds to the median value. The top whisker denotes the maximum value within the third quartile plus 1.5 times the interquartile range (bottom whisker is defined analogously). Dark gray dots represent outliers with values higher or lower than whiskers. Further box plots are based on the same definitions. ( B ) Motif analysis of triplexes formed between TriplexRNA (DNA-IP) and active promoters. The diagram depicts the fraction of antiparallel and parallel triplexes with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( C ) TDF analysis comparing the triplex-forming potential of top 2000 TriplexDNA-seq regions with top 1000 TriplexRNA (DNA-IP) (ranked by peak P -value). The number of putative DBSs (per kilobase of RNA) is shown. ( D ) Motif analysis of predicted triplexes formed between TriplexRNAs (DNA-IP) and TriplexDNA. The diagram depicts the fraction of antiparallel and parallel triplexes, with the respective motif and nucleotide composition of TFRs in TriplexRNA. ( E ) Box plot classifying triplex interactions between TriplexRNAs (DNA-IP) and TriplexDNA-seq regions as cis ( > 10 kb in the same chromosome) and trans (at different chromosomes) interactions, excluding underrepresented local interactions (within 10 kb distance). ( F ) EMSAs using 10 or 100 pmol of synthetic TriplexRNAs and 0.25 pmol of double–stranded 32 P-labeled oligonucleotides comprising target regions from TriplexDNA ( Supplementary Table S2 ). Reactions marked with an asterisk (*) were treated with 0.5 U RNase H. As a control (C), RNA without a putative TFR was used. Potential Hoogsteen base pairing between motifs and respective TFR sequences are shown; mismatches are marked (*). TriplexRNA-seq and TriplexDNA-seq data are from HeLa S3 cells. Adjusted P -values

    Techniques Used: Whisker Assay, Labeling

    27) Product Images from "A Eukaryotic Translation Initiation Factor 4E-Binding Protein Promotes mRNA Decapping and Is Required for PUF Repression"

    Article Title: A Eukaryotic Translation Initiation Factor 4E-Binding Protein Promotes mRNA Decapping and Is Required for PUF Repression

    Journal: Molecular and Cellular Biology

    doi: 10.1128/MCB.00483-12

    Eap1p promotes decapping of HO mRNA. (A) HO mRNA was cleaved with RNase H and a DNA oligonucleotide to produce a 1,600-nucleotide 5′ fragment and a 253-nucleotide 3′ fragment with a poly(A) tail of up to 80 adenosines (pA 80 ). (B) Northern
    Figure Legend Snippet: Eap1p promotes decapping of HO mRNA. (A) HO mRNA was cleaved with RNase H and a DNA oligonucleotide to produce a 1,600-nucleotide 5′ fragment and a 253-nucleotide 3′ fragment with a poly(A) tail of up to 80 adenosines (pA 80 ). (B) Northern

    Techniques Used: Northern Blot

    28) Product Images from "Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter"

    Article Title: Strong transcription blockage mediated by R-loop formation within a G-rich homopurine–homopyrimidine sequence localized in the vicinity of the promoter

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkx403

    Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.
    Figure Legend Snippet: Effect of RNase H upon transcription. Substrates containing the G-rich sequence were used in these experiments. See the Results section for description of the experiment. ( A ) Gel image. ( B ) Quantitation of the results. All run-off signals are normalized to the signal for promoter–distal substrate transcribed without RNase H.

    Techniques Used: Sequencing, Quantitation Assay

    Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).
    Figure Legend Snippet: Model for transcription blockage by R-loop formation in the vicinity of the promoter. The R-loop-prone (G-rich) DNA sequence is shown in turquoise, the rest of DNA is shown in gray, transcript from the R-loop-prone sequence is shown in dark blue, the rest of RNA is shown in black, a bent arrow indicates the transcription start site. RNA polymerase (RNAP) is shown as a gray circle. During transcription, an R-loop is formed with a certain probability p , while transcription proceeds without R-loop formation with probability 1 – p . R-loop formation could be initiated somewhere within the R-loop-prone sequence, but then the nascent RNA tail is likely to invade the entire R-loop-prone sequence (probably, even further upstream to the very start of transcription) as shown. The RNAP that created the R-loop could continue transcription in the ‘R-loop mode’, and then stall, either within, or at some distance downstream from the R-loop-prone sequence. At least some of the stalled RNAPs may remain bound to the DNA template (as shown), or could dissociate (not shown). In any case, R-loop formation blocks further rounds of transcription (the blockage is symbolized by the red crisscross). Addition of RNase H during transcription (all arrows that symbolize transitions within RNase H-related pathway are shown in green) leads to R-loop removal and, consequently, eliminates the blockage (blockage elimination is symbolized by the green path parallel to the crisscrossed path). The substrate DNA molecules from which R-loop was removed, then become available for further rounds of transcription, and would produce some number of normal full-sized transcripts, before an R-loop would form again. In addition, an RNAP stalled within an R-loop could resume transcription upon R-loop removal, producing a shorter transcript. That accounts for the pattern of transcription products obtained in the presence of RNase H (lane 4 in Figure 5 , the relevant part of it is placed in the present figure.).

    Techniques Used: Sequencing

    29) Product Images from "Rapid differentiation of avian infectious bronchitis virus isolates by sample to residual ratio quantitation using real-time reverse transcriptase-polymerase chain reaction"

    Article Title: Rapid differentiation of avian infectious bronchitis virus isolates by sample to residual ratio quantitation using real-time reverse transcriptase-polymerase chain reaction

    Journal: Journal of Virological Methods

    doi: 10.1016/j.jviromet.2004.11.022

    Native agarose gel analysis of Massachusetts 41 S1 runoff RNA cleavage as mediated by RNase H and chimeric oligonucleotides specific for strains in the Massachusetts, Arkansas, Connecticut, and Delaware/Georgia 98 serotypes. Lane 1 = RNA ladder, sizes from top to bottom are 9000, 7000, 5000, 3000, 2000, 1000, and 500 bases (New England Biolabs, Beverly, MA); lane 2 = uncleaved Massachusetts 41 S1 runoff RNA; lane 3 = Massachusetts 41 S1 runoff RNA incubated with anti-Massachusetts chimeric oligonucleotide and RNase H; lane 4 = same as lane 3 except, anti-Arkansas chimeric oligonucleotide used; lane 5 = same as lane 3, except anti-Connecticut chimeric oligonucleotide used; lane 6 = same as lane 3, except anti-Delaware chimeric oligonucleotide used. Arrows indicate cleavage products of ∼1500 and 300 bases.
    Figure Legend Snippet: Native agarose gel analysis of Massachusetts 41 S1 runoff RNA cleavage as mediated by RNase H and chimeric oligonucleotides specific for strains in the Massachusetts, Arkansas, Connecticut, and Delaware/Georgia 98 serotypes. Lane 1 = RNA ladder, sizes from top to bottom are 9000, 7000, 5000, 3000, 2000, 1000, and 500 bases (New England Biolabs, Beverly, MA); lane 2 = uncleaved Massachusetts 41 S1 runoff RNA; lane 3 = Massachusetts 41 S1 runoff RNA incubated with anti-Massachusetts chimeric oligonucleotide and RNase H; lane 4 = same as lane 3 except, anti-Arkansas chimeric oligonucleotide used; lane 5 = same as lane 3, except anti-Connecticut chimeric oligonucleotide used; lane 6 = same as lane 3, except anti-Delaware chimeric oligonucleotide used. Arrows indicate cleavage products of ∼1500 and 300 bases.

    Techniques Used: Agarose Gel Electrophoresis, Incubation

    Design of specific chimeric oligonucleotides for the Massachusetts and Arkansas serotypes. (A) An S1 gene sequence alignment of IBV strains belonging to the Massachusetts and Arkansas serotypes was performed for the region flanked by the primer set NewS1OLIGO5′ and M41L328. Only nucleotides 42–204 (ATG start site = 1) are shown. The sequences targeted by the anti-Massachusetts and anti-Arkansas chimeric oligonucleotides are boxed in black. Dots indicate nucleotides that are identical to the majority sequence, while letters indicate nucleotides that differ from the majority sequence. (B) The structure of a representative chimeric oligonucleotide hybridized to a complimentary strand of RNA. The solid black line with letters is the target RNA. Below is the chimeric oligonucleotide (uppercase = DNA bases, lowercase = 2′-O-Me RNA bases). The arrow denotes the site at which RNase H will cleave the strand of target RNA ( Bogdanova et al., 1995 , Yu and Steitz, 1997 ).
    Figure Legend Snippet: Design of specific chimeric oligonucleotides for the Massachusetts and Arkansas serotypes. (A) An S1 gene sequence alignment of IBV strains belonging to the Massachusetts and Arkansas serotypes was performed for the region flanked by the primer set NewS1OLIGO5′ and M41L328. Only nucleotides 42–204 (ATG start site = 1) are shown. The sequences targeted by the anti-Massachusetts and anti-Arkansas chimeric oligonucleotides are boxed in black. Dots indicate nucleotides that are identical to the majority sequence, while letters indicate nucleotides that differ from the majority sequence. (B) The structure of a representative chimeric oligonucleotide hybridized to a complimentary strand of RNA. The solid black line with letters is the target RNA. Below is the chimeric oligonucleotide (uppercase = DNA bases, lowercase = 2′-O-Me RNA bases). The arrow denotes the site at which RNase H will cleave the strand of target RNA ( Bogdanova et al., 1995 , Yu and Steitz, 1997 ).

    Techniques Used: Sequencing

    30) Product Images from "Comparative analysis of RNA enrichment methods for preparation of Cryptococcus neoformans RNA sequencing libraries"

    Article Title: Comparative analysis of RNA enrichment methods for preparation of Cryptococcus neoformans RNA sequencing libraries

    Journal: bioRxiv

    doi: 10.1101/2021.03.01.433483

    rRNA depletion efficiency: The percentage of rRNA reads in each library is graphed. The RNase H depletion method has the most efficient depletion (lowest percentage of rRNA reads), with the Poly(A) isolation method a close second, and the Ribo-Zero depletion method a distant third. Unenriched libraries show that rRNA makes up most of the RNA in C. neoformans .
    Figure Legend Snippet: rRNA depletion efficiency: The percentage of rRNA reads in each library is graphed. The RNase H depletion method has the most efficient depletion (lowest percentage of rRNA reads), with the Poly(A) isolation method a close second, and the Ribo-Zero depletion method a distant third. Unenriched libraries show that rRNA makes up most of the RNA in C. neoformans .

    Techniques Used: Isolation

    Specificity of rRNA depletion for protein-coding genes: Pearson correlations were calculated in the same way as Figure 2, but only for protein-coding genes, excluding genes containing coding-strand rRNA duplications. A. Unenriched libraries have high internal consistency for protein-coding genes. B. The RNase H depletion method has the best rRNA depletion specificity for protein-coding genes.
    Figure Legend Snippet: Specificity of rRNA depletion for protein-coding genes: Pearson correlations were calculated in the same way as Figure 2, but only for protein-coding genes, excluding genes containing coding-strand rRNA duplications. A. Unenriched libraries have high internal consistency for protein-coding genes. B. The RNase H depletion method has the best rRNA depletion specificity for protein-coding genes.

    Techniques Used:

    Specificity of rRNA depletion for all genes: Pearson correlations were calculated for normalized read counts of all annotated genes in the C. neoformans genome, excluding rRNA genes and genes containing coding-strand rRNA duplications. A. Unenriched libraries have high internal consistency as determined by leave-one-out cross correlation of each Unenriched library with the mean of other Unenriched libraries. B. The RNase H depletion method has the best overall rRNA depletion specificity, as determined by Pearson correlation of read counts for all genes with the Unenriched libraries. Pearson correlation coefficient (R) was calculated between each enriched library and the gold standard, the gene-wise average of counts across all Unenriched library replicates.
    Figure Legend Snippet: Specificity of rRNA depletion for all genes: Pearson correlations were calculated for normalized read counts of all annotated genes in the C. neoformans genome, excluding rRNA genes and genes containing coding-strand rRNA duplications. A. Unenriched libraries have high internal consistency as determined by leave-one-out cross correlation of each Unenriched library with the mean of other Unenriched libraries. B. The RNase H depletion method has the best overall rRNA depletion specificity, as determined by Pearson correlation of read counts for all genes with the Unenriched libraries. Pearson correlation coefficient (R) was calculated between each enriched library and the gold standard, the gene-wise average of counts across all Unenriched library replicates.

    Techniques Used:

    31) Product Images from "ADAR1 RNA editing enzyme regulates R-loop formation and genome stability at telomeres in cancer cells"

    Article Title: ADAR1 RNA editing enzyme regulates R-loop formation and genome stability at telomeres in cancer cells

    Journal: Nature Communications

    doi: 10.1038/s41467-021-21921-x

    Increased telomeric RNA:DNA hybrids containing variant repeats in ADAR1-depleted cells. Formation of telomeric repeat RNA:DNA hybrids containing A–C mismatches by in  cis  slipped hybridization ( a ,  b ).  a  TERRA RNAs transcribed from the region containing four T C AGGG (green) variant repeats surrounded by TTAGGG (gray) canonical repeats form an RNA:DNA hybrid containing four C–A mismatches by in  cis  slipped hybridization to the C-strand DNA containing canonical TTAGGG (CCCTAA) repeats.  b  TERRA RNAs transcribed from the region containing four TT G GGG (orange) variant repeats surrounded by TTAGGG (gray) canonical repeats form an RNA:DNA hybrid containing four A–C mismatches by in  cis  slipped hybridization to the C-strand DNA containing TT G GGG (CCC C AA) variant repeats.  c ,  d  Detection of increased RNA:DNA hybrids containing T C AGGG and TT G GGG variant repeats in ADAR1-depleted HeLa cells.  c  DRIP products were examined for G-strand RNAs of U C AGGG variant and UUAGGG canonical repeats by dot blot analysis using high-affinity LNA-oligonucleotide probes capable of distinguishing a single-nucleotide mismatch (Supplementary Fig.   4c ).  d  Similarly, DRIP products were examined for C-strand DNAs of TTAGGG (CCCTAA) canonical and TT G GGG (CCC C AA) variant repeats using LNA-oligonucleotide probes capable of distinguishing a single-nucleotide mismatch (Supplementary Fig.   4d ).  c ,  d  Dot blot signals were abolished by  E. coli -RNase H treatment prior to DRIP. The significance of the increase in RNA:DNA hybrids containing telomeric canonical and variant repeats (RNA and DNA strands) was confirmed by conducting three independent dot blot hybridization analysis of DRIP products. Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s  t  tests: * P
    Figure Legend Snippet: Increased telomeric RNA:DNA hybrids containing variant repeats in ADAR1-depleted cells. Formation of telomeric repeat RNA:DNA hybrids containing A–C mismatches by in cis slipped hybridization ( a , b ). a TERRA RNAs transcribed from the region containing four T C AGGG (green) variant repeats surrounded by TTAGGG (gray) canonical repeats form an RNA:DNA hybrid containing four C–A mismatches by in cis slipped hybridization to the C-strand DNA containing canonical TTAGGG (CCCTAA) repeats. b TERRA RNAs transcribed from the region containing four TT G GGG (orange) variant repeats surrounded by TTAGGG (gray) canonical repeats form an RNA:DNA hybrid containing four A–C mismatches by in cis slipped hybridization to the C-strand DNA containing TT G GGG (CCC C AA) variant repeats. c , d Detection of increased RNA:DNA hybrids containing T C AGGG and TT G GGG variant repeats in ADAR1-depleted HeLa cells. c DRIP products were examined for G-strand RNAs of U C AGGG variant and UUAGGG canonical repeats by dot blot analysis using high-affinity LNA-oligonucleotide probes capable of distinguishing a single-nucleotide mismatch (Supplementary Fig.  4c ). d Similarly, DRIP products were examined for C-strand DNAs of TTAGGG (CCCTAA) canonical and TT G GGG (CCC C AA) variant repeats using LNA-oligonucleotide probes capable of distinguishing a single-nucleotide mismatch (Supplementary Fig.  4d ). c , d  Dot blot signals were abolished by E. coli -RNase H treatment prior to DRIP. The significance of the increase in RNA:DNA hybrids containing telomeric canonical and variant repeats (RNA and DNA strands) was confirmed by conducting three independent dot blot hybridization analysis of DRIP products. Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s t tests: * P

    Techniques Used: Variant Assay, Hybridization, Countercurrent Chromatography, Dot Blot, Two Tailed Test

    A-to-I editing activity of ADAR1p110, not ADAR1p150 or ADAR2, is required for suppression of R-loops. a – e  Dot blot analysis for RNA:DNA hybrids was conducted using control oligos ( a ) or genomic DNA ( b–e ).  a  The S9.6 antibody recognized specifically RNA:DNA but not DNA:DNA or RNA:RNA oligo duplex controls.  b ,  c  Increased RNA:DNA hybrids were detected only in ADAR1-depleted but not in ADAR2-depleted HeLa cells.  b  The S9.6 antibody signals were abolished by  E. coli -RNase H treatment, confirming specific detection of RNA:DNA hybrids.  d  Comparison of RNA:DNA hybrid levels between depletion of ADAR1 versus depletion of known R-loop regulators.  e  Increased RNA:DNA hybrid formation resulting from depletion of endogenous ADAR1 was rescued by infection of ADAR1p110-WT (wild type) but not by infection of ADAR1p110-E912A deamination defective mutant or ADAR1p150-WT.  c – e  Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s  t  tests: * P
    Figure Legend Snippet: A-to-I editing activity of ADAR1p110, not ADAR1p150 or ADAR2, is required for suppression of R-loops. a – e Dot blot analysis for RNA:DNA hybrids was conducted using control oligos ( a ) or genomic DNA ( b–e ). a The S9.6 antibody recognized specifically RNA:DNA but not DNA:DNA or RNA:RNA oligo duplex controls. b , c Increased RNA:DNA hybrids were detected only in ADAR1-depleted but not in ADAR2-depleted HeLa cells. b The S9.6 antibody signals were abolished by E. coli -RNase H treatment, confirming specific detection of RNA:DNA hybrids. d Comparison of RNA:DNA hybrid levels between depletion of ADAR1 versus depletion of known R-loop regulators. e Increased RNA:DNA hybrid formation resulting from depletion of endogenous ADAR1 was rescued by infection of ADAR1p110-WT (wild type) but not by infection of ADAR1p110-E912A deamination defective mutant or ADAR1p150-WT. c – e Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s t tests: * P

    Techniques Used: Activity Assay, Dot Blot, Infection, Mutagenesis, Two Tailed Test

    Accumulation of R-loops at telomeres in ADAR1-depleted cells. a  ADAR1 depletion had no effects on already known sites prone to the formation of R-loops. Six sites were examined by qPCR analysis of DRIP products. PCR primers used are listed in Supplementary Data   1 . Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s  t  tests: n.s., not significant.  b  DRIP products were subjected to genomic DNA dot blot hybridization analysis with a probe containing the G-rich-telomere canonical repeat (TTAGGG), or a probe for α-satellite repeat,  Alu , or  LINE1  consensus sequence (Supplementary Data   1 ). ADAR1 depletion resulted in increased the formation of RNA:DNA hybrids specifically at telomeric repeats, which was abolished by  E. coli -RNase H treatment prior to DRIP.  c  Significance of the increased R-loop formation at telomeric repeats was confirmed by conducting three independent dot blot hybridization analyses of DRIP products. Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s  t  tests: ** P
    Figure Legend Snippet: Accumulation of R-loops at telomeres in ADAR1-depleted cells. a ADAR1 depletion had no effects on already known sites prone to the formation of R-loops. Six sites were examined by qPCR analysis of DRIP products. PCR primers used are listed in Supplementary Data  1 . Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s t tests: n.s., not significant. b DRIP products were subjected to genomic DNA dot blot hybridization analysis with a probe containing the G-rich-telomere canonical repeat (TTAGGG), or a probe for α-satellite repeat, Alu , or LINE1 consensus sequence (Supplementary Data  1 ). ADAR1 depletion resulted in increased the formation of RNA:DNA hybrids specifically at telomeric repeats, which was abolished by E. coli -RNase H treatment prior to DRIP. c Significance of the increased R-loop formation at telomeric repeats was confirmed by conducting three independent dot blot hybridization analyses of DRIP products. Data are mean ± SD ( n  = 3, biological replicates); significant differences were identified by two-tailed Student’s t tests: ** P

    Techniques Used: Real-time Polymerase Chain Reaction, Polymerase Chain Reaction, Two Tailed Test, Dot Blot, Hybridization, Sequencing

    32) Product Images from "RNase H1 Cooperates with DNA Gyrases to Restrict R-Loops and Maintain Genome Integrity in Arabidopsis Chloroplasts"

    Article Title: RNase H1 Cooperates with DNA Gyrases to Restrict R-Loops and Maintain Genome Integrity in Arabidopsis Chloroplasts

    Journal: The Plant Cell

    doi: 10.1105/tpc.17.00305

    AtRNH1C Is Important for R-Loop Homeostasis and Genome Integrity in Chloroplast. (A) Slot-blot assay analyzing overall R-loop levels in the chloroplast genome (left panel). Serial dilution of chloroplast DNAs (50, 100, and 200 ng) extracted from Col-0 and atrnh1c plants with or without RNase H treatment were slotted onto a nylon membrane and detected using the DNA:RNA hybrid antibody S9.6. DNA was stained using DuRed to show equal loading (right panel). (B) PFGE of cpDNA obtained from the same (panel 1 from left) and different numbers (panel 3 from left) of chloroplasts (“1×” = 1.5 × 10 7 ) from 2-week-old Col-0 and atrnh1c plants after staining with ethidium bromide. Panel 2 and panels 4 and 5 are blot hybridization of the probe (a 505-bp rbcL gene fragment) that corresponds to panels 1 and 3, respectively (panels 4 and 5 correspond to panel 3 with a different exposure time). Red dotted lines indicate the destabilized cpDNA, and arrows indicate DNA molecular size. A Lambda Ladder (New England Biolabs; N0341) was used to indicate the molecular weight. (C) PCR-based detection of chloroplast DNA rearrangement events in 2-week-old Col-0 and atrnh1c plants. Information on primers (a to f) used for the PCR reactions is referred to in the upper panel. ycf2 .
    Figure Legend Snippet: AtRNH1C Is Important for R-Loop Homeostasis and Genome Integrity in Chloroplast. (A) Slot-blot assay analyzing overall R-loop levels in the chloroplast genome (left panel). Serial dilution of chloroplast DNAs (50, 100, and 200 ng) extracted from Col-0 and atrnh1c plants with or without RNase H treatment were slotted onto a nylon membrane and detected using the DNA:RNA hybrid antibody S9.6. DNA was stained using DuRed to show equal loading (right panel). (B) PFGE of cpDNA obtained from the same (panel 1 from left) and different numbers (panel 3 from left) of chloroplasts (“1×” = 1.5 × 10 7 ) from 2-week-old Col-0 and atrnh1c plants after staining with ethidium bromide. Panel 2 and panels 4 and 5 are blot hybridization of the probe (a 505-bp rbcL gene fragment) that corresponds to panels 1 and 3, respectively (panels 4 and 5 correspond to panel 3 with a different exposure time). Red dotted lines indicate the destabilized cpDNA, and arrows indicate DNA molecular size. A Lambda Ladder (New England Biolabs; N0341) was used to indicate the molecular weight. (C) PCR-based detection of chloroplast DNA rearrangement events in 2-week-old Col-0 and atrnh1c plants. Information on primers (a to f) used for the PCR reactions is referred to in the upper panel. ycf2 .

    Techniques Used: Slot Blot Assay, Serial Dilution, Staining, Hybridization, Molecular Weight, Polymerase Chain Reaction

    RNase H1-Like Proteins in Arabidopsis. (A) Phylogenetic analysis and protein structures of RNase H1 proteins. RNase H1 proteins from Arabidopsis are marked with red dot. Protein structures are drawn to scale. The RNase H domains are highlighted in blue, DNA:RNA hybrid binding domains (HBD) are highlighted in red, and mitochondrion and/or chloroplast targeting sequences are predicted using TargetP and highlighted in green. (B) Subcellular localization of RNase H1 proteins AtRNH1A, AtRNH1B, and AtRNH1C in Arabidopsis are shown. Subcellular localization of AtRNH1A and AtRNH1B is analyzed from the 2-week-old stably transformed seedling roots and protoplast, respectively; transiently expressed GFP-tagged AtRNH1C were analyzed from protoplasts. Mitochondrion was stained by MitoTracker Red CMXRos (magenta). Chloroplasts can be distinguished by the chlorophyll autofluorescence (red). Bars = 10 µm.
    Figure Legend Snippet: RNase H1-Like Proteins in Arabidopsis. (A) Phylogenetic analysis and protein structures of RNase H1 proteins. RNase H1 proteins from Arabidopsis are marked with red dot. Protein structures are drawn to scale. The RNase H domains are highlighted in blue, DNA:RNA hybrid binding domains (HBD) are highlighted in red, and mitochondrion and/or chloroplast targeting sequences are predicted using TargetP and highlighted in green. (B) Subcellular localization of RNase H1 proteins AtRNH1A, AtRNH1B, and AtRNH1C in Arabidopsis are shown. Subcellular localization of AtRNH1A and AtRNH1B is analyzed from the 2-week-old stably transformed seedling roots and protoplast, respectively; transiently expressed GFP-tagged AtRNH1C were analyzed from protoplasts. Mitochondrion was stained by MitoTracker Red CMXRos (magenta). Chloroplasts can be distinguished by the chlorophyll autofluorescence (red). Bars = 10 µm.

    Techniques Used: Binding Assay, Stable Transfection, Transformation Assay, Staining

    33) Product Images from "Ornithine capture by a translating ribosome controls bacterial polyamine synthesis"

    Article Title: Ornithine capture by a translating ribosome controls bacterial polyamine synthesis

    Journal: bioRxiv

    doi: 10.1101/604074

    Purification of a SpeFL-70S complex stalled in the presence of ornithine. a , Overlaid absorbance profiles of sucrose gradients containing a translation mixture incubated without ornithine (black), in the presence of 10 mM L-ornithine (red) or in the presence of 10 mM L-ornithine followed by treatment with 100 µM puromycin (blue). A schematic diagram depicting the expected ribosomal species in each fraction is shown on the right. b , Overlaid absorbance profiles of sucrose gradients loaded with polysomal fractions from a, with (blue) or without (black) RNase H treatment. Expected ribosomal species for each fraction are shown on the right. c , Schematic representation of the purification strategy for SpeFL-70S. The collected fractions are indicated with gray boxes.
    Figure Legend Snippet: Purification of a SpeFL-70S complex stalled in the presence of ornithine. a , Overlaid absorbance profiles of sucrose gradients containing a translation mixture incubated without ornithine (black), in the presence of 10 mM L-ornithine (red) or in the presence of 10 mM L-ornithine followed by treatment with 100 µM puromycin (blue). A schematic diagram depicting the expected ribosomal species in each fraction is shown on the right. b , Overlaid absorbance profiles of sucrose gradients loaded with polysomal fractions from a, with (blue) or without (black) RNase H treatment. Expected ribosomal species for each fraction are shown on the right. c , Schematic representation of the purification strategy for SpeFL-70S. The collected fractions are indicated with gray boxes.

    Techniques Used: Purification, Incubation

    34) Product Images from "Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †"

    Article Title: Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †

    Journal:

    doi: 10.1128/MCB.01410-07

    RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)
    Figure Legend Snippet: RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)

    Techniques Used:

    35) Product Images from "Telomeres in ICF syndrome cells are vulnerable to DNA damage due to elevated DNA:RNA hybrids"

    Article Title: Telomeres in ICF syndrome cells are vulnerable to DNA damage due to elevated DNA:RNA hybrids

    Journal: Nature Communications

    doi: 10.1038/ncomms14015

    Sequence characteristics and potential for DNA:RNA hybrid formation in human distal subtelomeres. ( a ) 2 kb distal subtelomeric regions immediately adjacent to the telomere repeat tract were analysed for CpG density, GC content and GC skew over tiled overlapping 200 bp windows. The value for each window is depicted using a colour heatmap, as indicated below; each tick mark corresponds to a window. The analysed subtelomeres, indicated on the right, are clustered by the GC skew downstream of the annotated TERRA promoter, when available (grey: no annotation). Predicted Pol2 promoters and putative TSSs are indicated by boxes and black vertical lines, respectively. ( b ) DRIP-seq signal over each subtelomeric region is indicated by a colour heatmap reflecting signal enrichment over input for each 2 kb window (colour scheme is indicated at the right). A, B, and C refer to three distinct DRIP-seq datasets in human fibroblast (Fibro - A) and human Ntera2 cells (NT2 - B and C) measured relative to input. Each dataset included two independent technical replicates. The presence of consistent signal peaks in each replicate within a dataset is noted by an asterisk (*), as indicated. D corresponds to C but measured relative to an RNase H-treated control. Subtelomeric regions are arranged in the same order as  a . See ‘Results' for details. ( c ) Representative screenshots of DRIP-seq data over three distinct subtelomeric regions (22q, 15q and 10q, as indicated). Normalized DRIP-seq signal densities are displayed for each region over two distinct replicates (indicated by (I) and (II)) as well as input and, when available, RNase H-treated controls. The position of the Pol2 promoters and putative TSSs is shown at the top. Vertical dashed lines indicate the position of restriction enzyme sites used to fragment the genomic DNA before DRIP-seq. The grey shaded area highlights a TRF encompassing the TERRA promoter and/or downstream regions showing significant DRIP-seq signal.
    Figure Legend Snippet: Sequence characteristics and potential for DNA:RNA hybrid formation in human distal subtelomeres. ( a ) 2 kb distal subtelomeric regions immediately adjacent to the telomere repeat tract were analysed for CpG density, GC content and GC skew over tiled overlapping 200 bp windows. The value for each window is depicted using a colour heatmap, as indicated below; each tick mark corresponds to a window. The analysed subtelomeres, indicated on the right, are clustered by the GC skew downstream of the annotated TERRA promoter, when available (grey: no annotation). Predicted Pol2 promoters and putative TSSs are indicated by boxes and black vertical lines, respectively. ( b ) DRIP-seq signal over each subtelomeric region is indicated by a colour heatmap reflecting signal enrichment over input for each 2 kb window (colour scheme is indicated at the right). A, B, and C refer to three distinct DRIP-seq datasets in human fibroblast (Fibro - A) and human Ntera2 cells (NT2 - B and C) measured relative to input. Each dataset included two independent technical replicates. The presence of consistent signal peaks in each replicate within a dataset is noted by an asterisk (*), as indicated. D corresponds to C but measured relative to an RNase H-treated control. Subtelomeric regions are arranged in the same order as a . See ‘Results' for details. ( c ) Representative screenshots of DRIP-seq data over three distinct subtelomeric regions (22q, 15q and 10q, as indicated). Normalized DRIP-seq signal densities are displayed for each region over two distinct replicates (indicated by (I) and (II)) as well as input and, when available, RNase H-treated controls. The position of the Pol2 promoters and putative TSSs is shown at the top. Vertical dashed lines indicate the position of restriction enzyme sites used to fragment the genomic DNA before DRIP-seq. The grey shaded area highlights a TRF encompassing the TERRA promoter and/or downstream regions showing significant DRIP-seq signal.

    Techniques Used: Sequencing

    36) Product Images from "The RNA helicase RHAU (DHX36) suppresses expression of the transcription factor PITX1"

    Article Title: The RNA helicase RHAU (DHX36) suppresses expression of the transcription factor PITX1

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkt1340

    RHAU binds to the PITX1 3′-UTR between nucleotides 2110 and 2283. ( A ) Schematic representing the PITX1 mRNA. Primer binding sites are indicated by horizontal arrows, and DNA oligonucleotide complementary regions are indicated by solid horizontal lines. Relative locations of the predicted quadruplex-forming regions are indicated by vertical arrows. DNA oligonucleotide pairs used for RNase H-directed RNA digestion and their respective digestion sites on the full-length mRNA are indicated above. ( B ) RT-PCR analysis of bead-bound RNA following oligonucleotide-directed RNase digestion of the RNA–protein complex. Fold enrichment is expressed relative to 25 ng of total RNA for primer sets within the PITX1 coding sequence (CDS Primers) and 3′-UTR (UTR Primers). High fold enrichment is indicative of sustained binding of the primer template to RHAU following specific RNA digestion. Data represent the mean of three replicates ± standard error.
    Figure Legend Snippet: RHAU binds to the PITX1 3′-UTR between nucleotides 2110 and 2283. ( A ) Schematic representing the PITX1 mRNA. Primer binding sites are indicated by horizontal arrows, and DNA oligonucleotide complementary regions are indicated by solid horizontal lines. Relative locations of the predicted quadruplex-forming regions are indicated by vertical arrows. DNA oligonucleotide pairs used for RNase H-directed RNA digestion and their respective digestion sites on the full-length mRNA are indicated above. ( B ) RT-PCR analysis of bead-bound RNA following oligonucleotide-directed RNase digestion of the RNA–protein complex. Fold enrichment is expressed relative to 25 ng of total RNA for primer sets within the PITX1 coding sequence (CDS Primers) and 3′-UTR (UTR Primers). High fold enrichment is indicative of sustained binding of the primer template to RHAU following specific RNA digestion. Data represent the mean of three replicates ± standard error.

    Techniques Used: Binding Assay, Reverse Transcription Polymerase Chain Reaction, Sequencing

    37) Product Images from "RADICL-seq identifies general and cell type-specific principles of genome-wide RNA-chromatin interactions"

    Article Title: RADICL-seq identifies general and cell type-specific principles of genome-wide RNA-chromatin interactions

    Journal: bioRxiv

    doi: 10.1101/681924

    RADICL-seq method for the identification of RNA-chromatin interactions. a) Schematic representation of the RADICL-seq protocol. Top: sequence of enzymatic reactions taking place in fixed nuclei after partial lysis of the nuclear membrane. The adduct formed by genomic DNA (black), RNA (red) and proteins (blue circles) goes through controlled DNase digestion and chromatin preparation. After RNase H digestion, an adapter (dark blue) containing an internally biotinylated residue (black dot) bridges RNA and DNA in close proximity. Bottom: sequence of enzymatic reactions performed in solution. After reversal of crosslinks, the RNA-DNA chimera is converted into a fully dsDNA molecule and digested by EcoP15I enzyme to a designated length. After ligation of the sequencing linker and biotin pull-down, the library is PCR-amplified and high-throughput sequenced. b) Reproducibility of the RNA-DNA interaction frequencies across replicates, assessed by counting the occurrences of transcribed genes and 25 kb genomic bins pairs. c) RNA and d) DNA tags origin. The inner pie charts represent a broader classification into intergenic and genic (annotated genes), while the outer circles show a finer classification of the genic portion. e) Nuclear and f) cytosolic RNA-seq tags comparison with RADICL-seq RNA reads gene counts. The former are normalized to tags per million (TPM), while the latter are normalised to reads per kilobase (RPK). The linear regression lines are shown in red. g) Density of the normalized counts of DNA reads detected by RADICL-seq around ATAC-seq (red), DHS-seq (green) and H3K9me3 ChIP-seq (blue) peaks; dashed lines represent the density profiles of random genomic reads.
    Figure Legend Snippet: RADICL-seq method for the identification of RNA-chromatin interactions. a) Schematic representation of the RADICL-seq protocol. Top: sequence of enzymatic reactions taking place in fixed nuclei after partial lysis of the nuclear membrane. The adduct formed by genomic DNA (black), RNA (red) and proteins (blue circles) goes through controlled DNase digestion and chromatin preparation. After RNase H digestion, an adapter (dark blue) containing an internally biotinylated residue (black dot) bridges RNA and DNA in close proximity. Bottom: sequence of enzymatic reactions performed in solution. After reversal of crosslinks, the RNA-DNA chimera is converted into a fully dsDNA molecule and digested by EcoP15I enzyme to a designated length. After ligation of the sequencing linker and biotin pull-down, the library is PCR-amplified and high-throughput sequenced. b) Reproducibility of the RNA-DNA interaction frequencies across replicates, assessed by counting the occurrences of transcribed genes and 25 kb genomic bins pairs. c) RNA and d) DNA tags origin. The inner pie charts represent a broader classification into intergenic and genic (annotated genes), while the outer circles show a finer classification of the genic portion. e) Nuclear and f) cytosolic RNA-seq tags comparison with RADICL-seq RNA reads gene counts. The former are normalized to tags per million (TPM), while the latter are normalised to reads per kilobase (RPK). The linear regression lines are shown in red. g) Density of the normalized counts of DNA reads detected by RADICL-seq around ATAC-seq (red), DHS-seq (green) and H3K9me3 ChIP-seq (blue) peaks; dashed lines represent the density profiles of random genomic reads.

    Techniques Used: Sequencing, Lysis, Ligation, Polymerase Chain Reaction, Amplification, High Throughput Screening Assay, RNA Sequencing Assay, Chromatin Immunoprecipitation

    38) Product Images from "RNA from a simple-tandem repeat is required for sperm maturation and male fertility in Drosophila melanogaster"

    Article Title: RNA from a simple-tandem repeat is required for sperm maturation and male fertility in Drosophila melanogaster

    Journal: eLife

    doi: 10.7554/eLife.48940

    AAGAG RNA-FISH localizes RNA and not DNA. Confocal sections of cycle 14 nuclei treated with either ( a ) RNAseH or ( b ) RNAseIII after AAGAG RNA probe (magenta) hybridization. A higher laser intensity for the AAGAG probe channel was used in b to demonstrate abolishment of AAGAG signal.
    Figure Legend Snippet: AAGAG RNA-FISH localizes RNA and not DNA. Confocal sections of cycle 14 nuclei treated with either ( a ) RNAseH or ( b ) RNAseIII after AAGAG RNA probe (magenta) hybridization. A higher laser intensity for the AAGAG probe channel was used in b to demonstrate abolishment of AAGAG signal.

    Techniques Used: Fluorescence In Situ Hybridization, Hybridization

    AAGAG RNA foci contain single-stranded RNA and are not associated with R-loops. Confocal sections of embryonic nuclei in cycle 14 (with exception of left panel in ‘b’), nuclear periphery outlined in dotted circles. ( a ) No RNase control. ( b ) Treated with RNaseIII (left nucleus is cycle 12) ( c ) RNaseH ( d ) RNase1 and ( e ) RNaseA.
    Figure Legend Snippet: AAGAG RNA foci contain single-stranded RNA and are not associated with R-loops. Confocal sections of embryonic nuclei in cycle 14 (with exception of left panel in ‘b’), nuclear periphery outlined in dotted circles. ( a ) No RNase control. ( b ) Treated with RNaseIII (left nucleus is cycle 12) ( c ) RNaseH ( d ) RNase1 and ( e ) RNaseA.

    Techniques Used:

    39) Product Images from "Formation and Repair of Mismatches Containing Ribonucleotides and Oxidized Bases at Repeated DNA Sequences *"

    Article Title: Formation and Repair of Mismatches Containing Ribonucleotides and Oxidized Bases at Repeated DNA Sequences *

    Journal: The Journal of Biological Chemistry

    doi: 10.1074/jbc.M115.679209

    BER and RER activity on complex mispairs containing oxidized bases and ribonucleotides. When POL β incorporates rCMP opposite 8-oxodG (dG*), RNase H2 is going to efficiently remove rC from the resulting 8-oxodG:rC mispair, whereas OGG1 repair of dG* is slightly reduced. Should 8-oxodG:rA arise after rAMP incorporation, RER will process the rA containing strand, whereas MUTYH-mediated BER will be inhibited. Possible interference on RER activity might occur by concurrent recognition of the lesion. In the likelihood of limiting MTH1 hydrolytic activity, 8-oxorGTP (rG*TP) might be used by POL β to produce rG*:dA mispairs. These substrates will be efficiently processed by MUTYH and RNase H2. Simultaneous BER and RER activities might lead to the formation of double strand breaks ( DSB ) or intermediate repair products of unknown reparability.
    Figure Legend Snippet: BER and RER activity on complex mispairs containing oxidized bases and ribonucleotides. When POL β incorporates rCMP opposite 8-oxodG (dG*), RNase H2 is going to efficiently remove rC from the resulting 8-oxodG:rC mispair, whereas OGG1 repair of dG* is slightly reduced. Should 8-oxodG:rA arise after rAMP incorporation, RER will process the rA containing strand, whereas MUTYH-mediated BER will be inhibited. Possible interference on RER activity might occur by concurrent recognition of the lesion. In the likelihood of limiting MTH1 hydrolytic activity, 8-oxorGTP (rG*TP) might be used by POL β to produce rG*:dA mispairs. These substrates will be efficiently processed by MUTYH and RNase H2. Simultaneous BER and RER activities might lead to the formation of double strand breaks ( DSB ) or intermediate repair products of unknown reparability.

    Techniques Used: Activity Assay

    40) Product Images from "Translation initiation of alphavirus mRNA reveals new insights into the topology of the 48S initiation complex"

    Article Title: Translation initiation of alphavirus mRNA reveals new insights into the topology of the 48S initiation complex

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gky071

    eIF4A activity within the 48S complex. ( A ) RNase H-mapping of RNA–RNA interactions between SV-DLP U1 and 18S rRNA. The analysis was carried out in the absence or presence of 1 μM hippuristanol, with identification of the resulting RNA fragments indicated. For clarity, a schematic diagram of the ES6S and h16–18 regions of rabbit 18S rRNA with the primers used for RNase H digestion is shown. The use of oligos 4 and 9 limited the region of 18S rRNA (509–830) where the crosslinkings concentrated. Bands corresponding to crosslinking of SV DLP U1 mRNA with the ES6S region (680–1863) and h16-h18 helices (1–662) were quantified by densitometry and expressed as a ratio. Data are the mean ± SEM from four independent experiments. ( B ). Reactivity to SHAPE reagent (NMIA) is higher for unpaired nucleotides (red) and low for those involved in pairings (black). Stops corresponding to toeprints are marked with arrowheads. Quantification of toeprint ratios (17–19/23–25) in absence or presence of hippuristanol is shown from three independent experiments; data are the mean ± SEM.
    Figure Legend Snippet: eIF4A activity within the 48S complex. ( A ) RNase H-mapping of RNA–RNA interactions between SV-DLP U1 and 18S rRNA. The analysis was carried out in the absence or presence of 1 μM hippuristanol, with identification of the resulting RNA fragments indicated. For clarity, a schematic diagram of the ES6S and h16–18 regions of rabbit 18S rRNA with the primers used for RNase H digestion is shown. The use of oligos 4 and 9 limited the region of 18S rRNA (509–830) where the crosslinkings concentrated. Bands corresponding to crosslinking of SV DLP U1 mRNA with the ES6S region (680–1863) and h16-h18 helices (1–662) were quantified by densitometry and expressed as a ratio. Data are the mean ± SEM from four independent experiments. ( B ). Reactivity to SHAPE reagent (NMIA) is higher for unpaired nucleotides (red) and low for those involved in pairings (black). Stops corresponding to toeprints are marked with arrowheads. Quantification of toeprint ratios (17–19/23–25) in absence or presence of hippuristanol is shown from three independent experiments; data are the mean ± SEM.

    Techniques Used: Activity Assay

    Related Articles

    Binding Assay:

    Article Title: Improvement of RNA secondary structure prediction using RNase H cleavage and randomized oligonucleotides
    Article Snippet: .. The RNase H method, however, also determines the binding site mitigating the inability to deconvolute sequence degeneracy. ..

    Sequencing:

    Article Title: Improvement of RNA secondary structure prediction using RNase H cleavage and randomized oligonucleotides
    Article Snippet: .. The RNase H method, however, also determines the binding site mitigating the inability to deconvolute sequence degeneracy. ..

    Incubation:

    Article Title: The final step of 40S ribosomal subunit maturation is controlled by a dual key lock
    Article Snippet: RNase H digestion assay and RNA analysis For RNase H digestion assays, 250 ng of pre-40S purified RNAs were denatured at 95°C for 6 min with a RNA/ DNA /RNA reverse probe hybridizing in the 3’ end of 18S rRNA (probe RnaseH_3_Hyb1: 5’-UGUUACGACUUUU ACTTCCUCUAGAUAGUCAAGUUC-3’; 0,5 μl at 100 μM). .. After annealing by cooling down to room temperature for 20 min, the reaction mixture was diluted to 30 μl with a reaction mix containing 1 × RNase H reaction buffer, 25 μM DTT, 0.5 U/l RNasin (Promega) and 50 U RNase H (New England Biolabs), and incubated at 37°C for 30 min. .. The reaction was then blocked by addition of 0.3 M sodium acetate, pH 5.2 and 0.2 mM EDTA, and the RNAs were recovered by ethanol precipitation after phenol–chloroform–isoamylalcohol (25:24:1) extraction.

    Activity Assay:

    Article Title: Argonaute-based programmable RNase as a tool for cleavage of highly-structured RNA
    Article Snippet: Based on these results, 1.5 units of RNase H was used in each 10 μl reaction for subsequent experiments. .. Comparing activity of DISC and RNase H on unstructured or structured targets DISC reactions were carried out as described above for unstructured and structured targets using 500 nM AGO207, 10 nM gDNA and 1 nM target RNA. ..

    other:

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide
    Article Snippet: Although there are alternative interpretations, this could indicate that RNase H binds poorly to these resistant sites.

    Article Title: Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †
    Article Snippet: This suggests that for some transcripts, the step in 3′-end processing apparatus assembly that enhances splicing occurred before the transcripts were cut by RNase H. If this interpretation is correct, it would predict that a poly(A) signal that is capable of more-rapid assembly of the 3′-end processing apparatus would be able to complete this enhancing step for a greater fraction of transcripts before RNase H cutting occurs.

    Purification:

    Article Title: BRCA2 controls DNA:RNA hybrid level at DSBs by mediating RNase H2 recruitment
    Article Snippet: Briefly, DNA was extracted gently with phenol:chloroform:isoamyl alcohol (Sigma-Aldrich cat. no. P2069) and digested with Hind III, Eco RI, Bsr GI, Xba I, and Ssp I (NEB). .. After purification from restriction enzymes, half of the DNA was treated overnight with RNase H (NEB). .. In the meantime, serum-free medium containing the S9.6 antibody (kind gift from D. Piccini and M. Foiani) was mixed with protein A and protein G Dynabeads (Invitrogen) and incubated on a rotating wheel overnight at 4 °C.

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    New England Biolabs rnase h
    Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing <t>RNase</t> H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.
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    Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing RNase H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.

    Journal: Nucleic Acids Research

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide

    doi: 10.1093/nar/gkaa500

    Figure Lengend Snippet: Preparing and evaluating synthetic RNA–DNA hybrids as spike-ins for DRIP. ( A ) Experimental scheme showing how hybrids were synthesized. Briefly, target regions were amplified from E. coli genomic DNA with a flanking T7 promoter. RNA was prepared from these templates by in vitro transcription, then hybridized to a synthetic ssDNA oligo. Hybrids were purified by agarose gel electrophoresis. ( B ) Gel image showing RNase H reversible size-shifts after hybridization of RNA and DNA. Unlabeled samples were separated on a 2.5% agarose gel which was then stained with RedSafe nucleic acid staining solution. ( C ) qPCR of genomic (left) and spike-in (right) hybrids following transcription inhibition with DRB. RNase H (RH) treatment demonstrates antibody specificity. Error bars represent 95% confidence interval (CI) of the mean. Results are significantly different as determined by non-overlapping 95% CIs. In primer name, GB indicates gene body.

    Article Snippet: Although there are alternative interpretations, this could indicate that RNase H binds poorly to these resistant sites.

    Techniques: Synthesized, Amplification, In Vitro, Purification, Agarose Gel Electrophoresis, Hybridization, Staining, Real-time Polymerase Chain Reaction, Inhibition

    qDRIP provides strand-specific, high resolution RNA–DNA hybrid mapping. ( A ) Schematic of qDRIP experimental process. ( B ) Representative genome browser view of qDRIP-seq signal. From top to bottom: two qDRIP-seq biological replicates, RNase H digested sample pooled prior to IP, and input pooled from replicates. All tracks normalized by reads per million mapped. Negative strand signal in red, positive in blue. Bent arrows represent TSS, while large triangular arrows represent TES (transcription end site). ( C ) Read counts from template strand (TS) and non-template strand (NTS) of hybrids, as well as from ssDNA and dsDNA negative controls. ( D ) GC (green) and AT (red) skew across coding strand of qDRIP peaks, including 600 bp flanking 5’- and 3’-ends. ( E ) Fractions of qDRIP peaks overlapping noted genomic features ( P = 2.5e–2798, chi-square test). ( F ) Scaled metaplot of sense hybrids between TSS and first-intron exon boundary, as well as 1 kb upstream of TSS and 1 kb downstream of first intron-exon boundary. Tracks shown are mean IP (blue) and pooled input (grey). Bands represent 95% CI of mean read signal.

    Journal: Nucleic Acids Research

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide

    doi: 10.1093/nar/gkaa500

    Figure Lengend Snippet: qDRIP provides strand-specific, high resolution RNA–DNA hybrid mapping. ( A ) Schematic of qDRIP experimental process. ( B ) Representative genome browser view of qDRIP-seq signal. From top to bottom: two qDRIP-seq biological replicates, RNase H digested sample pooled prior to IP, and input pooled from replicates. All tracks normalized by reads per million mapped. Negative strand signal in red, positive in blue. Bent arrows represent TSS, while large triangular arrows represent TES (transcription end site). ( C ) Read counts from template strand (TS) and non-template strand (NTS) of hybrids, as well as from ssDNA and dsDNA negative controls. ( D ) GC (green) and AT (red) skew across coding strand of qDRIP peaks, including 600 bp flanking 5’- and 3’-ends. ( E ) Fractions of qDRIP peaks overlapping noted genomic features ( P = 2.5e–2798, chi-square test). ( F ) Scaled metaplot of sense hybrids between TSS and first-intron exon boundary, as well as 1 kb upstream of TSS and 1 kb downstream of first intron-exon boundary. Tracks shown are mean IP (blue) and pooled input (grey). Bands represent 95% CI of mean read signal.

    Article Snippet: Although there are alternative interpretations, this could indicate that RNase H binds poorly to these resistant sites.

    Techniques:

    R-loop lifetimes. ( A ) Schematic of transcription with and without DRB. ( B ) Ratio of DRB to control signal in RNase H-sensitive peaks, compared to estimated time without transcription. Error bands are 95% CI of the mean. Horizontal dotted line indicates a 2-fold decrease in DRB signal. ( C ) qPCR measurements during a DRB timecourse at regions predicted to be unstable (top) or stable (bottom) by pseudo-timecourse obtained from sequencing data. Error bars represent 95% CI of the mean. In primer name, GB indicates gene body. ( D ) GC content across 500 bp regions with shorter, longer or close to average (NS) lifetimes ( P  = 2.5e–143, Kruskal–Wallis test). ( E ) Biochemically determined G-quadruplex counts (  37 ) over the same regions as (D) ( P  = 2.7e–7, ANOVA on Negative Binomial regression, likelihood ratio test). ( F ) Relative replication fork directionality (RFD) (  39 ) to transcription over the same regions as (D), where 1 represents fully co-directional and –1 represents fully head-on ( P  = 3.5e–12, Kruskal–Wallis test). ( G ) Distribution of half-lives assuming first-order decay.

    Journal: Nucleic Acids Research

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide

    doi: 10.1093/nar/gkaa500

    Figure Lengend Snippet: R-loop lifetimes. ( A ) Schematic of transcription with and without DRB. ( B ) Ratio of DRB to control signal in RNase H-sensitive peaks, compared to estimated time without transcription. Error bands are 95% CI of the mean. Horizontal dotted line indicates a 2-fold decrease in DRB signal. ( C ) qPCR measurements during a DRB timecourse at regions predicted to be unstable (top) or stable (bottom) by pseudo-timecourse obtained from sequencing data. Error bars represent 95% CI of the mean. In primer name, GB indicates gene body. ( D ) GC content across 500 bp regions with shorter, longer or close to average (NS) lifetimes ( P = 2.5e–143, Kruskal–Wallis test). ( E ) Biochemically determined G-quadruplex counts ( 37 ) over the same regions as (D) ( P = 2.7e–7, ANOVA on Negative Binomial regression, likelihood ratio test). ( F ) Relative replication fork directionality (RFD) ( 39 ) to transcription over the same regions as (D), where 1 represents fully co-directional and –1 represents fully head-on ( P = 3.5e–12, Kruskal–Wallis test). ( G ) Distribution of half-lives assuming first-order decay.

    Article Snippet: Although there are alternative interpretations, this could indicate that RNase H binds poorly to these resistant sites.

    Techniques: Real-time Polymerase Chain Reaction, Sequencing

    RNase H resistant signal. ( A ) Aggregate plot of qDRIP-seq signal around the TSS of top 10,000 expressed genes as determined by GRO-seq ( 36 ). Tracks are IP (blue), RHR (red) and input (grey). Error bands represent 95% CI of mean. ( B ) Heatmaps of mean IP signal, RNase H-resistant signal and GC-skew around top 10,000 promoters ranked by GC-skew immediately (0–500 bp) downstream of the TSS. Correlation coefficient between IP signal and GC-skew was 0.06, whereas correlation coefficient for RHR signal was 0.22 (Spearman's rho). ( C ) GC-skew around RNase H resistant regions within the full (unfiltered) qDRIP-seq peak set. qDRIP peaks (red) compared to regions of equal lengths randomly selected from non-resistant qDRIP-peaks (blue). As before, bands represent 95% CI of mean read signal. ( D ) Same as (C), but showing biochemically determined G-quadruplex density ( 37 ) over these regions. ( E ) RNase H-resistant signal around RH-resistant peak calls. Peaks lying 5’ in genes (which DRB should affect) are in blue, while peaks lying 3’ in genes (which DRB should not affect) are in red. Left panel is RNase H treatment in control cells, while right panel is RNase H treatment in DRB treated cells. As before, error bands represent 95% CI of the mean.

    Journal: Nucleic Acids Research

    Article Title: qDRIP: a method to quantitatively assess RNA–DNA hybrid formation genome-wide

    doi: 10.1093/nar/gkaa500

    Figure Lengend Snippet: RNase H resistant signal. ( A ) Aggregate plot of qDRIP-seq signal around the TSS of top 10,000 expressed genes as determined by GRO-seq ( 36 ). Tracks are IP (blue), RHR (red) and input (grey). Error bands represent 95% CI of mean. ( B ) Heatmaps of mean IP signal, RNase H-resistant signal and GC-skew around top 10,000 promoters ranked by GC-skew immediately (0–500 bp) downstream of the TSS. Correlation coefficient between IP signal and GC-skew was 0.06, whereas correlation coefficient for RHR signal was 0.22 (Spearman's rho). ( C ) GC-skew around RNase H resistant regions within the full (unfiltered) qDRIP-seq peak set. qDRIP peaks (red) compared to regions of equal lengths randomly selected from non-resistant qDRIP-peaks (blue). As before, bands represent 95% CI of mean read signal. ( D ) Same as (C), but showing biochemically determined G-quadruplex density ( 37 ) over these regions. ( E ) RNase H-resistant signal around RH-resistant peak calls. Peaks lying 5’ in genes (which DRB should affect) are in blue, while peaks lying 3’ in genes (which DRB should not affect) are in red. Left panel is RNase H treatment in control cells, while right panel is RNase H treatment in DRB treated cells. As before, error bands represent 95% CI of the mean.

    Article Snippet: Although there are alternative interpretations, this could indicate that RNase H binds poorly to these resistant sites.

    Techniques:

    Complex purification and cryo-EM data processing workflow. Stalled ribosomal complexes were prepared using a bicistronic operon containing two identical copies of tnaC or tnaC(R23F) . A first sucrose gradient was performed to collect polysomes, followed by a second sucrose gradient after RNase H treatment to collect the monosomal fraction, which was the used to prepare the grids for cryo-EM data acquisition. The flowchart shows the workflow used to process and analyze cryo-EM data. Cross-sections of the final reconstructions are shown with the 70S ribosome in white, the mRNA in violet, the P-site tRNA in pink, the TnaC peptide in red and the L-Trp molecule in orange. Detailed maps of the TnaC peptide and L-Trp density are also shown for the TnaC–70S and TnaC(R23F)–70S complexes using the same color scheme, along with fitted atomic models. Fourier Shell Correlation (FSC) curves of the final reconstructions are shown as calculated by the RELION 3 . 1 (Ref. 51) post-processing algorithm. Cross-sections of maps displaying the local resolution calculated by RELION 3 . 1 (Ref. 51) are shown.

    Journal: bioRxiv

    Article Title: Structural basis for the tryptophan sensitivity of TnaC-mediated ribosome stalling

    doi: 10.1101/2021.03.31.437805

    Figure Lengend Snippet: Complex purification and cryo-EM data processing workflow. Stalled ribosomal complexes were prepared using a bicistronic operon containing two identical copies of tnaC or tnaC(R23F) . A first sucrose gradient was performed to collect polysomes, followed by a second sucrose gradient after RNase H treatment to collect the monosomal fraction, which was the used to prepare the grids for cryo-EM data acquisition. The flowchart shows the workflow used to process and analyze cryo-EM data. Cross-sections of the final reconstructions are shown with the 70S ribosome in white, the mRNA in violet, the P-site tRNA in pink, the TnaC peptide in red and the L-Trp molecule in orange. Detailed maps of the TnaC peptide and L-Trp density are also shown for the TnaC–70S and TnaC(R23F)–70S complexes using the same color scheme, along with fitted atomic models. Fourier Shell Correlation (FSC) curves of the final reconstructions are shown as calculated by the RELION 3 . 1 (Ref. 51) post-processing algorithm. Cross-sections of maps displaying the local resolution calculated by RELION 3 . 1 (Ref. 51) are shown.

    Article Snippet: This was achieved by mixing 50 µL of polysomes with 50 µL of a 100 µM stock of RNase H oligonucleotide , 4 µL RNase H (New England Biolabs), 644 µL buffer A and 2 µL DTT (1M).

    Techniques: Purification, Cryo-EM Sample Prep

    Comparing cleavage activity of DISC and RNase H. ( A ) Schematic of matched and mismatched guide and target pairs used to target four TRs across the HIV-1 ΔDIS 5′UTR RNA. For each pair, the HIV-1 ΔDIS 5′UTR sequence is shown on top and the perfectly matched gDNA strand is shown on the bottom. Circle indicates target position complementary to the first position of the guide that does not pair due to structural restrains by the protein. Black arrowheads indicate cleavage site. Mismatches between the guide and target strands are indicated by a black box around the bases of the guide that are mutated to the bases shown below the box. ( B ) Quantified cleavage products from the assay using matched and mismatched guide and target pairs described in (A) are plotted with solid bars representing the average of three replicates and circles representing individual replicates. Cleavage that was not detectable by the assay is indicated by ‘nd’. ( C and D ) Comparing DISC (circles) and RNase H (triangles) cleavage of the unstructured 60-nt target (C) or of a structured 352-nt RNA target (D). Bars indicate average cleavage of three replicates.

    Journal: Nucleic Acids Research

    Article Title: Argonaute-based programmable RNase as a tool for cleavage of highly-structured RNA

    doi: 10.1093/nar/gky496

    Figure Lengend Snippet: Comparing cleavage activity of DISC and RNase H. ( A ) Schematic of matched and mismatched guide and target pairs used to target four TRs across the HIV-1 ΔDIS 5′UTR RNA. For each pair, the HIV-1 ΔDIS 5′UTR sequence is shown on top and the perfectly matched gDNA strand is shown on the bottom. Circle indicates target position complementary to the first position of the guide that does not pair due to structural restrains by the protein. Black arrowheads indicate cleavage site. Mismatches between the guide and target strands are indicated by a black box around the bases of the guide that are mutated to the bases shown below the box. ( B ) Quantified cleavage products from the assay using matched and mismatched guide and target pairs described in (A) are plotted with solid bars representing the average of three replicates and circles representing individual replicates. Cleavage that was not detectable by the assay is indicated by ‘nd’. ( C and D ) Comparing DISC (circles) and RNase H (triangles) cleavage of the unstructured 60-nt target (C) or of a structured 352-nt RNA target (D). Bars indicate average cleavage of three replicates.

    Article Snippet: Comparing activity of DISC and RNase H on unstructured or structured targets DISC reactions were carried out as described above for unstructured and structured targets using 500 nM AGO207, 10 nM gDNA and 1 nM target RNA.

    Techniques: Activity Assay, Sequencing

    RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)

    Journal:

    Article Title: Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿Functional Coupling of Last-Intron Splicing and 3?-End Processing to Transcription In Vitro: the Poly(A) Signal Couples to Splicing before Committing to Cleavage ▿ †

    doi: 10.1128/MCB.01410-07

    Figure Lengend Snippet: RNase H cutting has much less effect on second-intron splicing when the SV40 late poly(A) signal defines the terminal exon. (A) This experiment was done as described in the legend to Fig. except that transcripts were postcut at the poly(A)

    Article Snippet: This suggests that for some transcripts, the step in 3′-end processing apparatus assembly that enhances splicing occurred before the transcripts were cut by RNase H. If this interpretation is correct, it would predict that a poly(A) signal that is capable of more-rapid assembly of the 3′-end processing apparatus would be able to complete this enhancing step for a greater fraction of transcripts before RNase H cutting occurs.

    Techniques: