e coli dna ligase  (New England Biolabs)


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    Structured Review

    New England Biolabs e coli dna ligase
    Individual replication fork progression is independent of primase A Micrographs showing replication products at 10 min where: i , all components present; or a component omitted: ii, DnaB and DnaC810; iii, Pol III*; iv, β; v, <t>SSB;</t> vi, primase. Composite, false-colored fields show anchor points for molecules that contain ssDNA, except i or v , where only long products were seen. In vi , surfaces were sparsely populated with <t>DNA</t> to avoid any ambiguity in molecule identification. Cyan, fields with flow off; magenta, same field with flow on showing fully-extended molecules. Molecules are bracketed for clarity. Scale bar: 10 μm, equal to 33.9 kb dsDNA or 80.3 knt SSB-bound ssDNA at 4,000 μl/h, without Mg 2+ ). B. Cartoon showing leading strand only product in a reaction lacking primase. C. Composite, false-colored image showing leading strand only replication without primase. Three replicating molecules ( 1, 2, 3 ). Molecules a, b and c are referred to later. D. Time-lapse, at 50-second intervals, of Molecules 1, 2 and 3 identified in C , colored by time-point as per C . E. Kymographs of molecules, numbered per C and D , showing fork progression without primase. Dashed grey line: position of anchor. Linear fits are from initiation to termination, yielding average fork rates. Pauses are included in the average here. F. Histograms of fork progression rates in the presence (grey) and absence of primase (light blue). Histograms fit to single Gaussians ( R 2 : with primase, 0.80; without primase, 0.94); no outliers were rejected. n , molecules. G. Processivities of single replisomes from live imaging experiments. Whisker plots of molecule lengths, with (320 nM) or without primase and/or β in flow. Data from 2 (primase, no β) or 3 (others) experiments. Horizontal bars, median; vertical bars, interquartile range. ***, significantly different pairs of populations (Kruskal-Wallis; P
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    Images

    1) Product Images from "Independent and Stochastic Action of DNA Polymerases in the Replisome"

    Article Title: Independent and Stochastic Action of DNA Polymerases in the Replisome

    Journal: Cell

    doi: 10.1016/j.cell.2017.05.041

    Individual replication fork progression is independent of primase A Micrographs showing replication products at 10 min where: i , all components present; or a component omitted: ii, DnaB and DnaC810; iii, Pol III*; iv, β; v, SSB; vi, primase. Composite, false-colored fields show anchor points for molecules that contain ssDNA, except i or v , where only long products were seen. In vi , surfaces were sparsely populated with DNA to avoid any ambiguity in molecule identification. Cyan, fields with flow off; magenta, same field with flow on showing fully-extended molecules. Molecules are bracketed for clarity. Scale bar: 10 μm, equal to 33.9 kb dsDNA or 80.3 knt SSB-bound ssDNA at 4,000 μl/h, without Mg 2+ ). B. Cartoon showing leading strand only product in a reaction lacking primase. C. Composite, false-colored image showing leading strand only replication without primase. Three replicating molecules ( 1, 2, 3 ). Molecules a, b and c are referred to later. D. Time-lapse, at 50-second intervals, of Molecules 1, 2 and 3 identified in C , colored by time-point as per C . E. Kymographs of molecules, numbered per C and D , showing fork progression without primase. Dashed grey line: position of anchor. Linear fits are from initiation to termination, yielding average fork rates. Pauses are included in the average here. F. Histograms of fork progression rates in the presence (grey) and absence of primase (light blue). Histograms fit to single Gaussians ( R 2 : with primase, 0.80; without primase, 0.94); no outliers were rejected. n , molecules. G. Processivities of single replisomes from live imaging experiments. Whisker plots of molecule lengths, with (320 nM) or without primase and/or β in flow. Data from 2 (primase, no β) or 3 (others) experiments. Horizontal bars, median; vertical bars, interquartile range. ***, significantly different pairs of populations (Kruskal-Wallis; P
    Figure Legend Snippet: Individual replication fork progression is independent of primase A Micrographs showing replication products at 10 min where: i , all components present; or a component omitted: ii, DnaB and DnaC810; iii, Pol III*; iv, β; v, SSB; vi, primase. Composite, false-colored fields show anchor points for molecules that contain ssDNA, except i or v , where only long products were seen. In vi , surfaces were sparsely populated with DNA to avoid any ambiguity in molecule identification. Cyan, fields with flow off; magenta, same field with flow on showing fully-extended molecules. Molecules are bracketed for clarity. Scale bar: 10 μm, equal to 33.9 kb dsDNA or 80.3 knt SSB-bound ssDNA at 4,000 μl/h, without Mg 2+ ). B. Cartoon showing leading strand only product in a reaction lacking primase. C. Composite, false-colored image showing leading strand only replication without primase. Three replicating molecules ( 1, 2, 3 ). Molecules a, b and c are referred to later. D. Time-lapse, at 50-second intervals, of Molecules 1, 2 and 3 identified in C , colored by time-point as per C . E. Kymographs of molecules, numbered per C and D , showing fork progression without primase. Dashed grey line: position of anchor. Linear fits are from initiation to termination, yielding average fork rates. Pauses are included in the average here. F. Histograms of fork progression rates in the presence (grey) and absence of primase (light blue). Histograms fit to single Gaussians ( R 2 : with primase, 0.80; without primase, 0.94); no outliers were rejected. n , molecules. G. Processivities of single replisomes from live imaging experiments. Whisker plots of molecule lengths, with (320 nM) or without primase and/or β in flow. Data from 2 (primase, no β) or 3 (others) experiments. Horizontal bars, median; vertical bars, interquartile range. ***, significantly different pairs of populations (Kruskal-Wallis; P

    Techniques Used: Flow Cytometry, Imaging, Whisker Assay

    2) Product Images from "Deoxyinosine repair in nuclear extracts of human cells"

    Article Title: Deoxyinosine repair in nuclear extracts of human cells

    Journal: Cell & Bioscience

    doi: 10.1186/s13578-015-0044-8

    Map of M13mp18 and f1PM based heteroduplex substrates. a The map of bacteriophage M13mp18 replicative form (RF) DNA shows restriction enzyme sites relevant to this study with derivatives M13LR1 and M13LR3 containing 22-bp insertions at the unique HindIII restriction site, and phage M13WX1 and M13X22 containing 26-bp and 22-bp insertions at Xba I site respectively. b The map of bacteriophage f1PM RF DNA with its derivative f1PMA with a 27-bp insertion at Xba I. ‘V’, phage viral strand. ‘C’, phage complementary strand. Underlines beneath each viral strand are the original insertion sequences. The C-strand from parental phage RF DNA was paired with viral strand of its insertion derivative to produce gapped duplex DNA, and the gap was sealed with dI or deoxyuridine containing synthetic oligodeoxyribonucleotide. A-I, C-I, G-I, T-I, and G-U are the resulting substrates and DNA sequence shown on each C-strand of the the synthetic linker used. In the presence of dI, the substrates were refractory to the restriction endonuclease scoring. After the repair, DNA products become sensitive to restriction endonuclease cleavage. The recognition sequence of corresponding restriction endonuclease markers for repair products are shown in bold on V-strands
    Figure Legend Snippet: Map of M13mp18 and f1PM based heteroduplex substrates. a The map of bacteriophage M13mp18 replicative form (RF) DNA shows restriction enzyme sites relevant to this study with derivatives M13LR1 and M13LR3 containing 22-bp insertions at the unique HindIII restriction site, and phage M13WX1 and M13X22 containing 26-bp and 22-bp insertions at Xba I site respectively. b The map of bacteriophage f1PM RF DNA with its derivative f1PMA with a 27-bp insertion at Xba I. ‘V’, phage viral strand. ‘C’, phage complementary strand. Underlines beneath each viral strand are the original insertion sequences. The C-strand from parental phage RF DNA was paired with viral strand of its insertion derivative to produce gapped duplex DNA, and the gap was sealed with dI or deoxyuridine containing synthetic oligodeoxyribonucleotide. A-I, C-I, G-I, T-I, and G-U are the resulting substrates and DNA sequence shown on each C-strand of the the synthetic linker used. In the presence of dI, the substrates were refractory to the restriction endonuclease scoring. After the repair, DNA products become sensitive to restriction endonuclease cleavage. The recognition sequence of corresponding restriction endonuclease markers for repair products are shown in bold on V-strands

    Techniques Used: Sequencing

    3) Product Images from "High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier"

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    Journal: bioRxiv

    doi: 10.1101/641506

    Transcriptional Maps of the Nucleosome Reveal that H2A.Z Enhances the Width and uH2B the Height of the Barrier (A) Median residence time histograms of Pol II transcription through bare NPS DNA (black), xWT (orange), hWT (red), H2A.Z (blue) and uH2B (green) nucleosomes. Bar width is 1 bp and major peak positions are labeled (in bp) above the corresponding peaks. NPS entry, dyad, NPS exit are marked with blue dashed lines. The polar plots on the right are the corresponding transcriptional maps of the nucleosome, formed by projecting the residence time histogram onto the surface of nucleosomal DNA. The top axis (red) indicates corresponding positions of the first half of nucleosome expressed as superhelical locations (SHL). n = 35, 23, 26, 21, 31, respectively for NPS DNA, xWT, hWT, H2A.Z and uH2B nucleosomes. (B) Crossing time (total time Pol II takes to cross the entire nucleosome region) distributions plotted using the complementary cumulative distribution function (CCDF, fraction of events longer than a given crossing time). Crossing times of Bare NPS DNA, Xenopus WT (xWT), human WT (hWT), uH2B and H2A.Z nucleosomes are plotted in black, orange, red, green and blue, respectively. See also Figure S6 on statistics of the crossing time, crossing probability, pause-free velocity and arrest position.
    Figure Legend Snippet: Transcriptional Maps of the Nucleosome Reveal that H2A.Z Enhances the Width and uH2B the Height of the Barrier (A) Median residence time histograms of Pol II transcription through bare NPS DNA (black), xWT (orange), hWT (red), H2A.Z (blue) and uH2B (green) nucleosomes. Bar width is 1 bp and major peak positions are labeled (in bp) above the corresponding peaks. NPS entry, dyad, NPS exit are marked with blue dashed lines. The polar plots on the right are the corresponding transcriptional maps of the nucleosome, formed by projecting the residence time histogram onto the surface of nucleosomal DNA. The top axis (red) indicates corresponding positions of the first half of nucleosome expressed as superhelical locations (SHL). n = 35, 23, 26, 21, 31, respectively for NPS DNA, xWT, hWT, H2A.Z and uH2B nucleosomes. (B) Crossing time (total time Pol II takes to cross the entire nucleosome region) distributions plotted using the complementary cumulative distribution function (CCDF, fraction of events longer than a given crossing time). Crossing times of Bare NPS DNA, Xenopus WT (xWT), human WT (hWT), uH2B and H2A.Z nucleosomes are plotted in black, orange, red, green and blue, respectively. See also Figure S6 on statistics of the crossing time, crossing probability, pause-free velocity and arrest position.

    Techniques Used: Labeling

    Topography Maps of the Nucleosome Revealed by Nucleosome Unzipping at Constant Force (A) Representative unzipping traces of bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes at 28 pN constant force. Unzipped bp are normalized to the beginning of the second NPS. Dashed lines mark entry, dyad and exit regions of the second NPS. Traces are shifted horizontally for clarity. (B) Mean residence time (RT) histogram of the unzipping fork along bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes during unzipping at a constant force of 28 pN. Bare NPS RTs are too short to see on the axes shown. Unzipped bp are normalized to the beginning of the second NPS core. Major peak positions are indicated above each peak (in bp). n = 33, 17, 20, 20, respectively for NPS DNA, WT, H2A.Z and uH2B nucleosomes. See also Figure S2 on assembly cooperativity of H2A.Z nucleosomes.
    Figure Legend Snippet: Topography Maps of the Nucleosome Revealed by Nucleosome Unzipping at Constant Force (A) Representative unzipping traces of bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes at 28 pN constant force. Unzipped bp are normalized to the beginning of the second NPS. Dashed lines mark entry, dyad and exit regions of the second NPS. Traces are shifted horizontally for clarity. (B) Mean residence time (RT) histogram of the unzipping fork along bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes during unzipping at a constant force of 28 pN. Bare NPS RTs are too short to see on the axes shown. Unzipped bp are normalized to the beginning of the second NPS core. Major peak positions are indicated above each peak (in bp). n = 33, 17, 20, 20, respectively for NPS DNA, WT, H2A.Z and uH2B nucleosomes. See also Figure S2 on assembly cooperativity of H2A.Z nucleosomes.

    Techniques Used:

    Mechanical Model for Pol II Transcription Through the Nucleosome (A) Schematic of the mechanical model, showing three different lengths of unwrapped DNA for a given polymerase position along the DNA sequence. The steric spheres are shown in purple (polymerase) and beige (nucleosome), while the DNA is shown as a tube. (i) shows a configuration with a short, sharply bent DNA linker connecting Pol II and the nucleosome, which are in contact and sterically pushing on each other. (ii) shows a medium-length straighter linker, with Pol II still pushing on the nucleosome. (iii) shows a long straight linker without contact between Pol II and the nucleosome. Linker DNA color corresponds to overall energy for each configuration (given in C). Black arrows represent tangent orientations of the DNA backbone at the point of polymerase binding (top) and for the last contact with the nucleosome (bottom). Linker length and bending angle (between indicated tangents) are labeled on each polymerase-nucleosome pair. (B) Model of Pol II dynamics. Pairs (p,q) indicate the Pol II state: p indicates the length of the RNA transcript, and q the number of base pairs backtracked from the most recent main pathway state. Pol II steps forward one base pair with rate k 0 or can enter a backtracked pathway by stepping backward one base pair at rate k b1 . From backtracked positions, Pol II can move forward a base pair with rate k f n or can backtrack another base pair at rate k b n . Moving forward from the first backtracked state returns Pol II to the main pathway. (C) Energy landscape of nucleosome-Pol II interaction, for constant DNA-nucleosome interaction energies of 1k B T per base pair. DNA unwrapping decreases the DNA linker conformational energy, while removing favorable DNA-nucleosome interactions, overall providing a minimum energy a few base pairs ahead of the front edge of Pol II. Forward Pol II steps are unfavorable as they shorten the DNA linker. Points i , ii , and iii correspond to configurations illustrated in A. Inset shows cross-section of energy landscape at Pol II position of 47 bp, highlighting the minimum in the energy landscape a few bps ahead of Pol II, at ∼52 bps unwrapped. Pol II progress through the nucleosome is defined as the position of the Pol II center plus an additional 17 bp for consistency with the transcribed distance in Figure 6 . (D) Dwell time profiles for human WT, H2A.Z, and uH2B nucleosomes. Solid black lines are experimental mean dwell times and colored dotted lines are the best fitted mean dwell times according to the mechanical model. (E) Estimated DNA-octamer interaction energy profiles for human WT, H2A.Z, and uH2B nucleosomes. The energy values are found such that they give the best fitted dwell times shown in (D). Peak positions referenced in the text are labeled in bp, relative to the start of the NPS. See also Figure S7 for fitting of nucleosome energy profiles based on Pol II dwell times.
    Figure Legend Snippet: Mechanical Model for Pol II Transcription Through the Nucleosome (A) Schematic of the mechanical model, showing three different lengths of unwrapped DNA for a given polymerase position along the DNA sequence. The steric spheres are shown in purple (polymerase) and beige (nucleosome), while the DNA is shown as a tube. (i) shows a configuration with a short, sharply bent DNA linker connecting Pol II and the nucleosome, which are in contact and sterically pushing on each other. (ii) shows a medium-length straighter linker, with Pol II still pushing on the nucleosome. (iii) shows a long straight linker without contact between Pol II and the nucleosome. Linker DNA color corresponds to overall energy for each configuration (given in C). Black arrows represent tangent orientations of the DNA backbone at the point of polymerase binding (top) and for the last contact with the nucleosome (bottom). Linker length and bending angle (between indicated tangents) are labeled on each polymerase-nucleosome pair. (B) Model of Pol II dynamics. Pairs (p,q) indicate the Pol II state: p indicates the length of the RNA transcript, and q the number of base pairs backtracked from the most recent main pathway state. Pol II steps forward one base pair with rate k 0 or can enter a backtracked pathway by stepping backward one base pair at rate k b1 . From backtracked positions, Pol II can move forward a base pair with rate k f n or can backtrack another base pair at rate k b n . Moving forward from the first backtracked state returns Pol II to the main pathway. (C) Energy landscape of nucleosome-Pol II interaction, for constant DNA-nucleosome interaction energies of 1k B T per base pair. DNA unwrapping decreases the DNA linker conformational energy, while removing favorable DNA-nucleosome interactions, overall providing a minimum energy a few base pairs ahead of the front edge of Pol II. Forward Pol II steps are unfavorable as they shorten the DNA linker. Points i , ii , and iii correspond to configurations illustrated in A. Inset shows cross-section of energy landscape at Pol II position of 47 bp, highlighting the minimum in the energy landscape a few bps ahead of Pol II, at ∼52 bps unwrapped. Pol II progress through the nucleosome is defined as the position of the Pol II center plus an additional 17 bp for consistency with the transcribed distance in Figure 6 . (D) Dwell time profiles for human WT, H2A.Z, and uH2B nucleosomes. Solid black lines are experimental mean dwell times and colored dotted lines are the best fitted mean dwell times according to the mechanical model. (E) Estimated DNA-octamer interaction energy profiles for human WT, H2A.Z, and uH2B nucleosomes. The energy values are found such that they give the best fitted dwell times shown in (D). Peak positions referenced in the text are labeled in bp, relative to the start of the NPS. See also Figure S7 for fitting of nucleosome energy profiles based on Pol II dwell times.

    Techniques Used: Sequencing, Binding Assay, Labeling

    Unzipping Traces of Single Human WT, H2A.Z, M3_M7, uH2B Nucleosomes and Tetrasomes. (A-E) Representative unzipping traces of WT nucleosomes (A), tetrasomes (B), H2A.Z nucleosomes (C), M3_M7 nucleosomes (D) and uH2B nucleosomes (E). Rezipping traces are not shown and they match bare NPS DNA rezipping traces. The unzipped bp (basepairs) are normalized to the beginning of the second NPS core. (F) Number of transitions per trace at the second NPS region. H2A.Z nucleosomes have on average one more transition per trace than WT or uH2B nucleosomes. A transition event is counted when the residence time peak is above an arbitrary threshold. (G-H) Partial unzipping of H2A.Z (G) and WT (H) nucleosomes reveals no lateral mobility induced by multiple rounds of unzipping-rezipping. The unzipping fork repeatedly propagates to the proximal dimer region followed by rezipping (not shown for clarity). The inset shows zoomed-in view of the boxed region, where the position of initial force rise remains unchanged. The dwelling of the unzipping fork in alternative positions (labeled above the dashed lines in bp) is consistent with hopping observed in this region. (I) Native PAGE gels showing homogenous WT, H2A.Z and uH2B nucleosome samples used for single-molecule unzipping experiments.
    Figure Legend Snippet: Unzipping Traces of Single Human WT, H2A.Z, M3_M7, uH2B Nucleosomes and Tetrasomes. (A-E) Representative unzipping traces of WT nucleosomes (A), tetrasomes (B), H2A.Z nucleosomes (C), M3_M7 nucleosomes (D) and uH2B nucleosomes (E). Rezipping traces are not shown and they match bare NPS DNA rezipping traces. The unzipped bp (basepairs) are normalized to the beginning of the second NPS core. (F) Number of transitions per trace at the second NPS region. H2A.Z nucleosomes have on average one more transition per trace than WT or uH2B nucleosomes. A transition event is counted when the residence time peak is above an arbitrary threshold. (G-H) Partial unzipping of H2A.Z (G) and WT (H) nucleosomes reveals no lateral mobility induced by multiple rounds of unzipping-rezipping. The unzipping fork repeatedly propagates to the proximal dimer region followed by rezipping (not shown for clarity). The inset shows zoomed-in view of the boxed region, where the position of initial force rise remains unchanged. The dwelling of the unzipping fork in alternative positions (labeled above the dashed lines in bp) is consistent with hopping observed in this region. (I) Native PAGE gels showing homogenous WT, H2A.Z and uH2B nucleosome samples used for single-molecule unzipping experiments.

    Techniques Used: Labeling, Clear Native PAGE

    Observation of Multiple Nucleosomal States at the Proximal Dimer Region (A) Time traces of number of base pairs unzipped (relative to beginning of the second NPS) with hWT nucleosome for fixed trap separations of 1031 nm, 1045 nm, and 1060 nm (top to bottom). Color indicates increasing trap separation (purple to red), corresponding to clusters in Figure S3F . Grey dashed lines indicate 17, 23, and 28 base pairs unzipped. (B) Probability distributions for the number of DNA bps unzipped, computed from force-extension data in Figure S3F . Each curve is from a different trap separation, matching colors in A and Figure S3F . Distributions are shown for both bare DNA (top) and WT nucleosome (bottom). Vertical black dotted line indicates the start of the second NPS. Vertical grey dashed lines indicate peak positions for bare DNA (with position in bp labeled), showing that WT nucleosome shifts the first peak within the NPS, and gives rise to an additional peak at 28 bp. See Figure S3F for force-extension data. (C) Zoomed-in view of the black dashed box in (B). Peak positions are labeled in bp. (D) DNA unzipping energy computed by assuming the unzipped bp distributions from data in Figure S3F (including distributions in B) correspond to equilibrium Boltzmann statistics. Inset Δ E shows the DNA-octamer interaction energy, computed as the difference between unzipping energies in the presence of WT (red), H2A.Z (blue), and uH2B (green) nucleosomes and unzipping energies for bare DNA only (black). Vertical black dashed lines and * indicate peak positions (labeled in bp). See also Figure S3 on hopping traces and analysis of energy landscape from equilibrium data.
    Figure Legend Snippet: Observation of Multiple Nucleosomal States at the Proximal Dimer Region (A) Time traces of number of base pairs unzipped (relative to beginning of the second NPS) with hWT nucleosome for fixed trap separations of 1031 nm, 1045 nm, and 1060 nm (top to bottom). Color indicates increasing trap separation (purple to red), corresponding to clusters in Figure S3F . Grey dashed lines indicate 17, 23, and 28 base pairs unzipped. (B) Probability distributions for the number of DNA bps unzipped, computed from force-extension data in Figure S3F . Each curve is from a different trap separation, matching colors in A and Figure S3F . Distributions are shown for both bare DNA (top) and WT nucleosome (bottom). Vertical black dotted line indicates the start of the second NPS. Vertical grey dashed lines indicate peak positions for bare DNA (with position in bp labeled), showing that WT nucleosome shifts the first peak within the NPS, and gives rise to an additional peak at 28 bp. See Figure S3F for force-extension data. (C) Zoomed-in view of the black dashed box in (B). Peak positions are labeled in bp. (D) DNA unzipping energy computed by assuming the unzipped bp distributions from data in Figure S3F (including distributions in B) correspond to equilibrium Boltzmann statistics. Inset Δ E shows the DNA-octamer interaction energy, computed as the difference between unzipping energies in the presence of WT (red), H2A.Z (blue), and uH2B (green) nucleosomes and unzipping energies for bare DNA only (black). Vertical black dashed lines and * indicate peak positions (labeled in bp). See also Figure S3 on hopping traces and analysis of energy landscape from equilibrium data.

    Techniques Used: Labeling

    H2A.Z Nucleosomes Assemble More Cooperatively than WT nucleosomes (A) Sequence swaps between H2A and H2A.Z reveal important regions for hexasome formation. The native PAGE gel shows the propensity to form hexasomes during assembly of H2A, H2A.Z and swapped mutant nucleosomes. DNA is Cy5-labeled 70N0 where “N” denotes the 601 NPS. We found that this DNA configuration is more prone to hexasome formation due to the asymmetric nature of the 601 sequence. Two octamer-to-DNA ratios are tested for each sample and are shown below its corresponding lanes. The nucleosome, hexasome or DNA bands are indicated on the right. (B) Sequence alignment of H2A and H2A.Z swap mutants. Nomenclature of the swap mutants follows Clarkson et al .
    Figure Legend Snippet: H2A.Z Nucleosomes Assemble More Cooperatively than WT nucleosomes (A) Sequence swaps between H2A and H2A.Z reveal important regions for hexasome formation. The native PAGE gel shows the propensity to form hexasomes during assembly of H2A, H2A.Z and swapped mutant nucleosomes. DNA is Cy5-labeled 70N0 where “N” denotes the 601 NPS. We found that this DNA configuration is more prone to hexasome formation due to the asymmetric nature of the 601 sequence. Two octamer-to-DNA ratios are tested for each sample and are shown below its corresponding lanes. The nucleosome, hexasome or DNA bands are indicated on the right. (B) Sequence alignment of H2A and H2A.Z swap mutants. Nomenclature of the swap mutants follows Clarkson et al .

    Techniques Used: Sequencing, Clear Native PAGE, Mutagenesis, Labeling

    A ‘Molecular Ruler’ Gauges the Positions of an Elongating Pol II with Near-Basepair Accuracy (A) Experimental design of an improved single-molecule nucleosomal transcription assay. A single biotinylated Pol II (purple molecular structure) is tethered between two optical traps. Pol II transcription is measured as increases in distance between the two beads at 10 pN constant force. The inset box shows the composition of the template, which is constructed by ligating Pol II stalled complex (cyan), the molecular ruler (green), NPS DNA (or nucleosome, yellow-grey), and a short inter-strand crosslinked DNA (for stalling Pol II, red). The ‘molecular ruler’ consists of eight tandem repeats of a 64-bp DNA (green), each harboring a single sequence-encoded pause site. (B) A representative trace of a single Pol II transcribing through a Xenopus WT nucleosome. The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The inset shows a zoomed-in view of the boxed region, highlighting the repeating pause patterns within the ‘molecular ruler’. The grey dashed lines are the predicted pause sites, whereas the short green lines mark the actual pauses of Pol II. (C) Zoomed-in view of Pol II dynamics within the NPS region of (B). The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The right y-axis (in bp) is normalized to the beginning of the NPS. The left y-axis shows regions preceding the dyad as SHL in red. Black arrows indicates representative events of backtracking, pausing, productive elongation, and hopping. Regions corresponding to Pol II located at SHL(-5) and SHL(-1) are indicated with green and cyan dashed lines, with the corresponding Pol II-nucleosome complex structures plotted on top (PDB 6A5P for PolII-SHL(-5), 6A5 T for PolII-SHL(-1)). Pol II, histones, template DNA, non-template DNA are colored in grey, green, red and blue, respectively. See also Figure S4 on detailed characterization of the ‘molecular ruler’.
    Figure Legend Snippet: A ‘Molecular Ruler’ Gauges the Positions of an Elongating Pol II with Near-Basepair Accuracy (A) Experimental design of an improved single-molecule nucleosomal transcription assay. A single biotinylated Pol II (purple molecular structure) is tethered between two optical traps. Pol II transcription is measured as increases in distance between the two beads at 10 pN constant force. The inset box shows the composition of the template, which is constructed by ligating Pol II stalled complex (cyan), the molecular ruler (green), NPS DNA (or nucleosome, yellow-grey), and a short inter-strand crosslinked DNA (for stalling Pol II, red). The ‘molecular ruler’ consists of eight tandem repeats of a 64-bp DNA (green), each harboring a single sequence-encoded pause site. (B) A representative trace of a single Pol II transcribing through a Xenopus WT nucleosome. The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The inset shows a zoomed-in view of the boxed region, highlighting the repeating pause patterns within the ‘molecular ruler’. The grey dashed lines are the predicted pause sites, whereas the short green lines mark the actual pauses of Pol II. (C) Zoomed-in view of Pol II dynamics within the NPS region of (B). The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The right y-axis (in bp) is normalized to the beginning of the NPS. The left y-axis shows regions preceding the dyad as SHL in red. Black arrows indicates representative events of backtracking, pausing, productive elongation, and hopping. Regions corresponding to Pol II located at SHL(-5) and SHL(-1) are indicated with green and cyan dashed lines, with the corresponding Pol II-nucleosome complex structures plotted on top (PDB 6A5P for PolII-SHL(-5), 6A5 T for PolII-SHL(-1)). Pol II, histones, template DNA, non-template DNA are colored in grey, green, red and blue, respectively. See also Figure S4 on detailed characterization of the ‘molecular ruler’.

    Techniques Used: Construct, Sequencing

    Dual-trap Optical Tweezers Single-molecule Unzipping Assay Unwinds Nucleosomal DNA and Maps Histone-DNA Interactions (A) Geometry of the single-molecule unzipping assay. Dashed arrows denote directions of trap movement (20 nm/s) during unzipping (red arrow) or rezipping (black arrow). Two DNA handles connect to the template DNA, which consists of two tandem NPS repeats and an end hairpin. Diagram illustrates nucleosome unzipping, with the second NPS replaced with a pre-assembled mononucleosome. For simplicity, linkers and restriction sites flanking the NPS are not shown. (B, C) Unzipping (red) and rezipping (black) traces of bare NPS DNA (B) and a single WT human nucleosome (C). The presence of the nucleosome on the second NPS causes characteristic high force (20-40 pN) transitions that correspond to disruption of histone-DNA contacts. The unzipped basepairs (bp) are normalized to the beginning of the second NPS. The nucleosome rezipping trace matches that of bare NPS DNA, indicating complete histone removal during unzipping. (D) Representative unzipping traces of tetrasome (cyan), WT (red), H2A.Z (blue), and uH2B (green) nucleosomes. For clarity, only the region after entering the second NPS (corresponding to the boxed region in (C)) is shown, with the unzipped bp normalized to the beginning of the second NPS. The three dashed lines are entry, dyad, and exit of the second NPS, respectively. Rezipping traces, identical to those of B and C, are not shown. (E) Topography maps are plotted as force-weighted residence time (RT) histograms of the unzipping fork along bare NPS DNA, tetrasome and different types of nucleosomes during unzipping at constant trap separation speed of 20 nm/s. The grey histograms with colored stripes (excluding Bare NPS DNA and WT Nucleosome) are residual plots found by subtracting the WT nucleosome RTs. Unzipped bp are normalized to the beginning of the second NPS core. Major peaks are highlighted with grey dashed lines, with the peak positions (in bp) labeled above the peaks. (Left to right: 17, 22, 26, 31, 35, 41, 52, 61, 69, 109, 112, 122 bp). n = 34, 41, 34, 39, 35, 10, respectively for NPS DNA, hWT, H2A.Z, M3_M7, uH2B nucleosome and tetrasome. See also Figure S1 for representative unzipping traces and analysis.
    Figure Legend Snippet: Dual-trap Optical Tweezers Single-molecule Unzipping Assay Unwinds Nucleosomal DNA and Maps Histone-DNA Interactions (A) Geometry of the single-molecule unzipping assay. Dashed arrows denote directions of trap movement (20 nm/s) during unzipping (red arrow) or rezipping (black arrow). Two DNA handles connect to the template DNA, which consists of two tandem NPS repeats and an end hairpin. Diagram illustrates nucleosome unzipping, with the second NPS replaced with a pre-assembled mononucleosome. For simplicity, linkers and restriction sites flanking the NPS are not shown. (B, C) Unzipping (red) and rezipping (black) traces of bare NPS DNA (B) and a single WT human nucleosome (C). The presence of the nucleosome on the second NPS causes characteristic high force (20-40 pN) transitions that correspond to disruption of histone-DNA contacts. The unzipped basepairs (bp) are normalized to the beginning of the second NPS. The nucleosome rezipping trace matches that of bare NPS DNA, indicating complete histone removal during unzipping. (D) Representative unzipping traces of tetrasome (cyan), WT (red), H2A.Z (blue), and uH2B (green) nucleosomes. For clarity, only the region after entering the second NPS (corresponding to the boxed region in (C)) is shown, with the unzipped bp normalized to the beginning of the second NPS. The three dashed lines are entry, dyad, and exit of the second NPS, respectively. Rezipping traces, identical to those of B and C, are not shown. (E) Topography maps are plotted as force-weighted residence time (RT) histograms of the unzipping fork along bare NPS DNA, tetrasome and different types of nucleosomes during unzipping at constant trap separation speed of 20 nm/s. The grey histograms with colored stripes (excluding Bare NPS DNA and WT Nucleosome) are residual plots found by subtracting the WT nucleosome RTs. Unzipped bp are normalized to the beginning of the second NPS core. Major peaks are highlighted with grey dashed lines, with the peak positions (in bp) labeled above the peaks. (Left to right: 17, 22, 26, 31, 35, 41, 52, 61, 69, 109, 112, 122 bp). n = 34, 41, 34, 39, 35, 10, respectively for NPS DNA, hWT, H2A.Z, M3_M7, uH2B nucleosome and tetrasome. See also Figure S1 for representative unzipping traces and analysis.

    Techniques Used: Labeling

    Hopping of the Unzipping Fork Near the Proximal Dimer Region of the Nucleosome (A, B) Unzipping traces of human WT nucleosome (A) and bare NPS DNA (B). Hopping near the proximal dimer region of WT nucleosome is indicated with a dashed blue square box; no similar hopping was observed in the corresponding region during unzipping of bare NPS DNA. Insets are the zoomed-in view of the dashed square boxes. Unzipped bp is normalized to the beginning of the second NPS. Rezipping traces are not shown for clarity. (C) Aligned individual force-extension curves (thin colored curves) and mean force-extension curve (thick black curve), for bare DNA. (D) Energy of DNA unzipping for each base pair, calculated from mean force-extension curve. (E) Comparison of experimental mean force-extension curve (blue) to the force-extension calculated from the extracted DNA unzipping energy (red). (F) Force-extension traces obtained at fixed trap separations with WT nucleosome. Color indicates increasing trap separation (purple to red), with number indicating the trap separation in nm. (G) DNA unzipping energy for each base pair, calculated from equilibrium hopping data at multiple fixed trap separations as in (F). (H) Comparison of experimental mean force-extension curve for bare DNA (black) and DNA with a WT nucleosome (red) to the force-extension curve predicted by the apparent DNA unzipping energy from equilibrium hopping data for the WT nucleosome (cyan).
    Figure Legend Snippet: Hopping of the Unzipping Fork Near the Proximal Dimer Region of the Nucleosome (A, B) Unzipping traces of human WT nucleosome (A) and bare NPS DNA (B). Hopping near the proximal dimer region of WT nucleosome is indicated with a dashed blue square box; no similar hopping was observed in the corresponding region during unzipping of bare NPS DNA. Insets are the zoomed-in view of the dashed square boxes. Unzipped bp is normalized to the beginning of the second NPS. Rezipping traces are not shown for clarity. (C) Aligned individual force-extension curves (thin colored curves) and mean force-extension curve (thick black curve), for bare DNA. (D) Energy of DNA unzipping for each base pair, calculated from mean force-extension curve. (E) Comparison of experimental mean force-extension curve (blue) to the force-extension calculated from the extracted DNA unzipping energy (red). (F) Force-extension traces obtained at fixed trap separations with WT nucleosome. Color indicates increasing trap separation (purple to red), with number indicating the trap separation in nm. (G) DNA unzipping energy for each base pair, calculated from equilibrium hopping data at multiple fixed trap separations as in (F). (H) Comparison of experimental mean force-extension curve for bare DNA (black) and DNA with a WT nucleosome (red) to the force-extension curve predicted by the apparent DNA unzipping energy from equilibrium hopping data for the WT nucleosome (cyan).

    Techniques Used:

    High-resolution Trajectories of Individual Pol II Enzymes Transcribing through WT, H2A.Z and uH2B Nucleosomes (A, B) Representative traces of single Pol II enzymes transcribing through single human WT nucleosomes. The grey dotted lines are the pause sites within the ‘molecular ruler’. The inset (black) is the residence time of Pol II within the ‘molecular ruler’, highlighting repeating pausing signatures of Pol II. The three black dashed lines indicate NPS entry, dyad and NPS exit. Relative positions of Pol II on the template DNA are shown as a cartoon on the right. The traces in blue, green, red and cyan are examples of successful nucleosome crossing, while the trace in grey is an example of Pol II arrest in the nucleosome. For comparison, a trace of Pol II transcribing through bare NPS DNA (black) is shown on the left. Zoomed in traces of high-resolution Pol II dynamics within the NPS are shown in (B), highlighting (black arrowheads) long-lived pausing, backtracking and hopping events. The traces are shifted horizontally for clarity. The right y-axis is normalized to the beginning of the NPS, with the major pause positions marked (in bp) on the right. (C, D) Representative traces of single Pol II enzymes transcribing through single human H2A.Z nucleosomes. (C) shows the full traces and (D) is a zoomed-in view of the high-resolution dynamics within the NPS region. (E, F) Representative traces of single Pol II enzymes transcribing through single human uH2B nucleosomes. (E) shows the full traces and (F) is a zoomed-in view of the high-resolution dynamics within the NPS region. See also Figure S5 on backtracking and hopping dynamics.
    Figure Legend Snippet: High-resolution Trajectories of Individual Pol II Enzymes Transcribing through WT, H2A.Z and uH2B Nucleosomes (A, B) Representative traces of single Pol II enzymes transcribing through single human WT nucleosomes. The grey dotted lines are the pause sites within the ‘molecular ruler’. The inset (black) is the residence time of Pol II within the ‘molecular ruler’, highlighting repeating pausing signatures of Pol II. The three black dashed lines indicate NPS entry, dyad and NPS exit. Relative positions of Pol II on the template DNA are shown as a cartoon on the right. The traces in blue, green, red and cyan are examples of successful nucleosome crossing, while the trace in grey is an example of Pol II arrest in the nucleosome. For comparison, a trace of Pol II transcribing through bare NPS DNA (black) is shown on the left. Zoomed in traces of high-resolution Pol II dynamics within the NPS are shown in (B), highlighting (black arrowheads) long-lived pausing, backtracking and hopping events. The traces are shifted horizontally for clarity. The right y-axis is normalized to the beginning of the NPS, with the major pause positions marked (in bp) on the right. (C, D) Representative traces of single Pol II enzymes transcribing through single human H2A.Z nucleosomes. (C) shows the full traces and (D) is a zoomed-in view of the high-resolution dynamics within the NPS region. (E, F) Representative traces of single Pol II enzymes transcribing through single human uH2B nucleosomes. (E) shows the full traces and (F) is a zoomed-in view of the high-resolution dynamics within the NPS region. See also Figure S5 on backtracking and hopping dynamics.

    Techniques Used:

    Biochemical and Single-molecule Characterization of the “Molecular Ruler” (A) In vitro transcription assay identifies a major pause site within a single repeat sequence (64 bp). The band corresponding to the pause site is highlighted with a dotted red box. The sequence of the single repeat template DNA is shown above the gel, with the identified pause site highlighted in red. (B) Histogram of the length of one repeat unit (periodicity, d). From aligned traces of Pol II transcription through xWT nucleosomes, d is calculated to be 21.1 ± 0.3 nm. (C) Mean (black) and median (red) residence time (in log scale) of Pol II transcribing through the repeat sequence confirms a single major pause site at 59 bp in the repeat sequence, matching the site identified in (A). (D) Zoomed in view of the alignment of traces using the “molecular ruler” (cartoon on the right). The major pause site within each repeat sequence is marked with a grey horizontal line and a red dot next to the “molecular ruler”. Short horizontal black lines indicate identified pauses and vertical black lines (with the exception of few cases where the tether breaks in the middle) indicate the entry and exit of the “molecular ruler”.
    Figure Legend Snippet: Biochemical and Single-molecule Characterization of the “Molecular Ruler” (A) In vitro transcription assay identifies a major pause site within a single repeat sequence (64 bp). The band corresponding to the pause site is highlighted with a dotted red box. The sequence of the single repeat template DNA is shown above the gel, with the identified pause site highlighted in red. (B) Histogram of the length of one repeat unit (periodicity, d). From aligned traces of Pol II transcription through xWT nucleosomes, d is calculated to be 21.1 ± 0.3 nm. (C) Mean (black) and median (red) residence time (in log scale) of Pol II transcribing through the repeat sequence confirms a single major pause site at 59 bp in the repeat sequence, matching the site identified in (A). (D) Zoomed in view of the alignment of traces using the “molecular ruler” (cartoon on the right). The major pause site within each repeat sequence is marked with a grey horizontal line and a red dot next to the “molecular ruler”. Short horizontal black lines indicate identified pauses and vertical black lines (with the exception of few cases where the tether breaks in the middle) indicate the entry and exit of the “molecular ruler”.

    Techniques Used: In Vitro, Sequencing

    Crossing Time, Crossing Probability and Pause-free Velocity of Pol II during Transcription through NPS DNA or Nucleosomes (A-E) Histograms of crossing time of Pol II transcription through bare NPS DNA (A), xWT (B), hWT (C), H2A.Z (D) and uH2B (E) nucleosomes. See also Figure 6B . (F) Relative percentage of Pol II molecules that are arrested or crossed during transcription through bare NPS DNA or nucleosomes. (G) Pol II arrest positions within the NPS. The positions are normalized to the beginning of the NPS. Each dot is a single arresting event. The percentages of arresting before or after dyad are shown below the dots. (H) Pause-free velocity of Pol II molecules before, inside and after NPS during transcription through bare NPS DNA, and xWT, hWT, H2A.Z and uH2B nucleosomes. Only traces that reached the stall site at the end of the template are considered. Pause-free velocities are calculated in three fastest regions (to partially correct for velocity differences due to sequence variations) up to100 bp before, inside, and up to 100 bp after NPS.
    Figure Legend Snippet: Crossing Time, Crossing Probability and Pause-free Velocity of Pol II during Transcription through NPS DNA or Nucleosomes (A-E) Histograms of crossing time of Pol II transcription through bare NPS DNA (A), xWT (B), hWT (C), H2A.Z (D) and uH2B (E) nucleosomes. See also Figure 6B . (F) Relative percentage of Pol II molecules that are arrested or crossed during transcription through bare NPS DNA or nucleosomes. (G) Pol II arrest positions within the NPS. The positions are normalized to the beginning of the NPS. Each dot is a single arresting event. The percentages of arresting before or after dyad are shown below the dots. (H) Pause-free velocity of Pol II molecules before, inside and after NPS during transcription through bare NPS DNA, and xWT, hWT, H2A.Z and uH2B nucleosomes. Only traces that reached the stall site at the end of the template are considered. Pause-free velocities are calculated in three fastest regions (to partially correct for velocity differences due to sequence variations) up to100 bp before, inside, and up to 100 bp after NPS.

    Techniques Used: Sequencing

    4) Product Images from "Cas12a-assisted precise targeted cloning using in vivo Cre-lox recombination"

    Article Title: Cas12a-assisted precise targeted cloning using in vivo Cre-lox recombination

    Journal: Nature Communications

    doi: 10.1038/s41467-021-21275-4

    Characterization of various genomic DNA digestion/DNA assembly combinations in the CAPTURE method. a Schematics of T4 DNA polymerase exo + fill-in DNA assembly. In step 1, DNA molecules ends are chewed back by T4 DNA polymerase to create ssDNA overhangs. The reaction mixture’s temperature is increased to 75 °C to inactivate T4 DNA polymerase and potentially remove ssDNA secondary structures. Temperature is then decreased to 50 °C to allow for ssDNA overhang hybridization. In step 2, by addition of fresh T4 DNA polymerase, and dNTPs, DNA gaps in the hybridized DNA molecule are filled. E. coli DNA ligase is then used to ligate the nicks and produce the final assembly product. b Comparison of different digestion/DNA assembly combinations in cloning four high GC-content BGCs from Actinomycetes. The Fn Cas12a/T4 exo + fill-in strategy showed ~100% cloning efficiency for all four target BGCs. RE: restriction enzymes. For each cloning experiment, at least seven colonies were selected and the purified plasmids from each colony were analyzed by restriction digestion. The cloning efficiencies were calculated as the ratio of correct colonies to the total number of checked colonies. Each experiment was performed in three biological replicates and data are presented as mean values ± standard error (SEM). c Summary of results for cloning uncharacterized BGCs using CAPTURE. BGCs ranging from 10 to 113 kb can be robustly cloned using the CAPTURE method at close to 100% efficiency regardless of their GC-content. Source data are provided as a Source Data file.
    Figure Legend Snippet: Characterization of various genomic DNA digestion/DNA assembly combinations in the CAPTURE method. a Schematics of T4 DNA polymerase exo + fill-in DNA assembly. In step 1, DNA molecules ends are chewed back by T4 DNA polymerase to create ssDNA overhangs. The reaction mixture’s temperature is increased to 75 °C to inactivate T4 DNA polymerase and potentially remove ssDNA secondary structures. Temperature is then decreased to 50 °C to allow for ssDNA overhang hybridization. In step 2, by addition of fresh T4 DNA polymerase, and dNTPs, DNA gaps in the hybridized DNA molecule are filled. E. coli DNA ligase is then used to ligate the nicks and produce the final assembly product. b Comparison of different digestion/DNA assembly combinations in cloning four high GC-content BGCs from Actinomycetes. The Fn Cas12a/T4 exo + fill-in strategy showed ~100% cloning efficiency for all four target BGCs. RE: restriction enzymes. For each cloning experiment, at least seven colonies were selected and the purified plasmids from each colony were analyzed by restriction digestion. The cloning efficiencies were calculated as the ratio of correct colonies to the total number of checked colonies. Each experiment was performed in three biological replicates and data are presented as mean values ± standard error (SEM). c Summary of results for cloning uncharacterized BGCs using CAPTURE. BGCs ranging from 10 to 113 kb can be robustly cloned using the CAPTURE method at close to 100% efficiency regardless of their GC-content. Source data are provided as a Source Data file.

    Techniques Used: Hybridization, Clone Assay, Purification

    Development of the CAPTURE method. a Overview of the workflow. In the first step, purified genomic DNA is digested by Cas12a enzyme to release the target BGC fragment. In the second step, digestion products are mixed with two DNA receivers containing lox P sites at their ends. The target BGC fragment and DNA receivers are assembled together using T4 DNA polymerase exo + fill-in DNA assembly. In the final step, the assembly mixture is transformed into E. coli cells harboring a circularization helper plasmid. The linear DNA is able to circularize in vivo by Cre- lox recombination. b DNA map of helper plasmid pBE14. tcr : tetracycline resistance marker; araBAD : L-arabinose inducible promoter and its regulator; gam : phage lambda Red gam gene; pSC101: temperature-sensitive origin of replication; recA1 : mutated E. coli recA gene to increase transformation efficiency. c Comparison of recombination frequency between Flp (pBE11) and Cre (pBE12) helper plasmids. -: without L-arabinose induction, +: with L-arabinose induction. Recombination frequencies were calculated based on the ratio of white colonies to the total number of acquired colonies. d Linear DNA transformation efficiency for E. coli cells harboring pBE11 (Flp), pBE12 (Cre), pBE14 (Cre and recA1) helper plasmids. Both pBE12 and pBE14 E. coli cells exhibited transformation efficiencies similar to circular DNA. e Comparison of in vitro versus in vivo circularization for two large (50 kb, 73 kb) linear DNA molecules. In vivo circularization showed ~33-fold and 150-fold higher frequency than in vitro circularization for 50 kb and 73 kb molecules, respectively. Circularization frequencies were calculated based on the number of colonies acquired for each circularization experiment in comparison to the number of colonies acquired after transformation of the original circular DNA (see Methods for full description). Each experiment was performed in three biological replicates and data are presented as mean values ± standard deviation (SD). Source data are provided as a Source Data file.
    Figure Legend Snippet: Development of the CAPTURE method. a Overview of the workflow. In the first step, purified genomic DNA is digested by Cas12a enzyme to release the target BGC fragment. In the second step, digestion products are mixed with two DNA receivers containing lox P sites at their ends. The target BGC fragment and DNA receivers are assembled together using T4 DNA polymerase exo + fill-in DNA assembly. In the final step, the assembly mixture is transformed into E. coli cells harboring a circularization helper plasmid. The linear DNA is able to circularize in vivo by Cre- lox recombination. b DNA map of helper plasmid pBE14. tcr : tetracycline resistance marker; araBAD : L-arabinose inducible promoter and its regulator; gam : phage lambda Red gam gene; pSC101: temperature-sensitive origin of replication; recA1 : mutated E. coli recA gene to increase transformation efficiency. c Comparison of recombination frequency between Flp (pBE11) and Cre (pBE12) helper plasmids. -: without L-arabinose induction, +: with L-arabinose induction. Recombination frequencies were calculated based on the ratio of white colonies to the total number of acquired colonies. d Linear DNA transformation efficiency for E. coli cells harboring pBE11 (Flp), pBE12 (Cre), pBE14 (Cre and recA1) helper plasmids. Both pBE12 and pBE14 E. coli cells exhibited transformation efficiencies similar to circular DNA. e Comparison of in vitro versus in vivo circularization for two large (50 kb, 73 kb) linear DNA molecules. In vivo circularization showed ~33-fold and 150-fold higher frequency than in vitro circularization for 50 kb and 73 kb molecules, respectively. Circularization frequencies were calculated based on the number of colonies acquired for each circularization experiment in comparison to the number of colonies acquired after transformation of the original circular DNA (see Methods for full description). Each experiment was performed in three biological replicates and data are presented as mean values ± standard deviation (SD). Source data are provided as a Source Data file.

    Techniques Used: Purification, Transformation Assay, Plasmid Preparation, In Vivo, Marker, In Vitro, Standard Deviation

    5) Product Images from "Regulation by interdomain communication of a headful packaging nuclease from bacteriophage T4"

    Article Title: Regulation by interdomain communication of a headful packaging nuclease from bacteriophage T4

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkq1191

    The large terminase gp17 is a weak endonuclease. ( A ) Time course of cleavage of circular pET28b plasmid (100 ng) by gp17 (1.5 µM). ( B ) Cleavage of topoisomerase 1 relaxed DNA by gp17. ( C ) Increasing concentrations of gp17 (9–900 nM) were incubated with circular pAd10 DNA (400 ng, 0.9 nM) and the amount of undigested circular DNA in each lane was quantified by laser densitometry. ( D ) Comparison of the nuclease activity of T4 gp17 with DNase I (non-specific nickase) and Sau3A1 (frequent cutting restriction endonuclease). Increasing concentrations of each enzyme were incubated with circular pAd10 DNA (400 ng, 0.9 nM) with the enzyme (monomer) to DNA ratio (number of molecules of each) varied over a range of 10–1000:1. The enzyme:DNA ratio at 50% cleavage was determined by quantifying the amount of undigested circular DNA in each lane. Values represent average of duplicates from two independent experiments. The ‘C’ lanes represent untreated DNA. See ‘Materials and Methods’ section for additional details.
    Figure Legend Snippet: The large terminase gp17 is a weak endonuclease. ( A ) Time course of cleavage of circular pET28b plasmid (100 ng) by gp17 (1.5 µM). ( B ) Cleavage of topoisomerase 1 relaxed DNA by gp17. ( C ) Increasing concentrations of gp17 (9–900 nM) were incubated with circular pAd10 DNA (400 ng, 0.9 nM) and the amount of undigested circular DNA in each lane was quantified by laser densitometry. ( D ) Comparison of the nuclease activity of T4 gp17 with DNase I (non-specific nickase) and Sau3A1 (frequent cutting restriction endonuclease). Increasing concentrations of each enzyme were incubated with circular pAd10 DNA (400 ng, 0.9 nM) with the enzyme (monomer) to DNA ratio (number of molecules of each) varied over a range of 10–1000:1. The enzyme:DNA ratio at 50% cleavage was determined by quantifying the amount of undigested circular DNA in each lane. Values represent average of duplicates from two independent experiments. The ‘C’ lanes represent untreated DNA. See ‘Materials and Methods’ section for additional details.

    Techniques Used: Plasmid Preparation, Incubation, Activity Assay

    ATP stimulates gp17 nuclease. ( A ) Nuclease activity of gp17 is stimulated in presence of ATP. gp17 (1 µM) was incubated with linear pAd10 DNA (100 ng) in the presence of increasing concentrations of ATP (0.05–5 mM) . Note that the DNA is degraded to small fragments in some of the lanes. Therefore, very little DNA smear is seen in these lanes. ( B ) Nuclease activity of gp17 in the presence of ATP analogs. gp17 (1.2 µM) was incubated with circular pAd10 DNA (200 ng) in the presence of ATP, ADP, ATP-γS or AMP-PNP (1 mM). ( C ) The T4 nuclease domain (C360, amino acids 360–577) (left panel; lanes 1–7) or the RB49 nuclease domain (C360, amino acids 358–607) (right panel; lanes 8–14) is not stimulated by ATP. T4 C360 (4 µM) or RB49 C360 (1 µM), either alone (lanes 2 and 9) or in the presence of ATP (lanes 3–7 and 10–14) was incubated with linear pAd10 DNA (100 ng) for 15 min and the samples were analyzed by 0.8% (w/v) agarose gel electrophoresis. Lanes 1 and 8 labeled as ‘C’ are control lanes having untreated DNA.
    Figure Legend Snippet: ATP stimulates gp17 nuclease. ( A ) Nuclease activity of gp17 is stimulated in presence of ATP. gp17 (1 µM) was incubated with linear pAd10 DNA (100 ng) in the presence of increasing concentrations of ATP (0.05–5 mM) . Note that the DNA is degraded to small fragments in some of the lanes. Therefore, very little DNA smear is seen in these lanes. ( B ) Nuclease activity of gp17 in the presence of ATP analogs. gp17 (1.2 µM) was incubated with circular pAd10 DNA (200 ng) in the presence of ATP, ADP, ATP-γS or AMP-PNP (1 mM). ( C ) The T4 nuclease domain (C360, amino acids 360–577) (left panel; lanes 1–7) or the RB49 nuclease domain (C360, amino acids 358–607) (right panel; lanes 8–14) is not stimulated by ATP. T4 C360 (4 µM) or RB49 C360 (1 µM), either alone (lanes 2 and 9) or in the presence of ATP (lanes 3–7 and 10–14) was incubated with linear pAd10 DNA (100 ng) for 15 min and the samples were analyzed by 0.8% (w/v) agarose gel electrophoresis. Lanes 1 and 8 labeled as ‘C’ are control lanes having untreated DNA.

    Techniques Used: Activity Assay, Incubation, Agarose Gel Electrophoresis, Labeling

    gp17 nuclease prefers long DNA substrates and cleaves at the ends of linear DNA. ( A ) Increasing concentrations of gp17 were incubated with 0.9 nM each of 29 kb pAd10 plasmid DNA or 2.6 kb pUC19 plasmid DNA. The undigested circular DNA was quantified and used to determine the percent of cleaved DNA at different gp17:DNA ratios. Values represent the average of duplicates from two independent experiments. ( B ) gp17 preference for longer DNA molecules was seen by incubating gp17 (3 µM, lanes 2–7) with a 2-log DNA ladder (400 ng, 0.1–10 kb, New England Biolabs) for 2–30 min. ( C ) Autoradiogram showing the cleavage of γ 32 P end-labeled λ-HindIII DNA fragments (0.5 pmol, 125–23 130 bp, Promega) by gp17 (1.2 µM) (lanes 2–6) or DNase I (0.0024 µM, 500-fold less than gp17) (lanes 7–11). Lane 1 has untreated DNA. ( D ) gp17 nuclease generates blunt ends. Circular pUC19 DNA (40 ng) was cleaved by gp17 (lanes 2–4) or BamH1 (lanes 5–7). The cleaved DNA was then treated with E. coli DNA ligase (lanes 3 and 6) or T4 DNA ligase (lanes 4 and 7). Lanes labeled as ‘C’ are control untreated lanes. See ‘Materials and Methods’ section for additional details.
    Figure Legend Snippet: gp17 nuclease prefers long DNA substrates and cleaves at the ends of linear DNA. ( A ) Increasing concentrations of gp17 were incubated with 0.9 nM each of 29 kb pAd10 plasmid DNA or 2.6 kb pUC19 plasmid DNA. The undigested circular DNA was quantified and used to determine the percent of cleaved DNA at different gp17:DNA ratios. Values represent the average of duplicates from two independent experiments. ( B ) gp17 preference for longer DNA molecules was seen by incubating gp17 (3 µM, lanes 2–7) with a 2-log DNA ladder (400 ng, 0.1–10 kb, New England Biolabs) for 2–30 min. ( C ) Autoradiogram showing the cleavage of γ 32 P end-labeled λ-HindIII DNA fragments (0.5 pmol, 125–23 130 bp, Promega) by gp17 (1.2 µM) (lanes 2–6) or DNase I (0.0024 µM, 500-fold less than gp17) (lanes 7–11). Lane 1 has untreated DNA. ( D ) gp17 nuclease generates blunt ends. Circular pUC19 DNA (40 ng) was cleaved by gp17 (lanes 2–4) or BamH1 (lanes 5–7). The cleaved DNA was then treated with E. coli DNA ligase (lanes 3 and 6) or T4 DNA ligase (lanes 4 and 7). Lanes labeled as ‘C’ are control untreated lanes. See ‘Materials and Methods’ section for additional details.

    Techniques Used: Incubation, Plasmid Preparation, Labeling

    6) Product Images from "Partial Reconstitution of Human DNA Mismatch Repair In Vitro: Characterization of the Role of Human Replication Protein A"

    Article Title: Partial Reconstitution of Human DNA Mismatch Repair In Vitro: Characterization of the Role of Human Replication Protein A

    Journal: Molecular and Cellular Biology

    doi: 10.1128/MCB.22.7.2037-2046.2002

    Reconstitution of MMR in vitro. (A) Fractionation of a HeLa nuclear extract into three components required for MMR. (B) Reconstitution of MMR requires SS1, SS2, and FII. The DNA substrate (100 ng of the 5′ G-T heteroduplex) was incubated for 15 min at 37°C in the reaction buffer with fractions as indicated. Amounts of protein used were 15 μg of SS1, 1.5 μg of SS2, or 30 μg of FII. DNA was extracted, treated with Hin dIII and Bsp 106, electrophoresed on an agarose gel, and visualized by ethidium bromide staining under UV illumination. ND, not detectable.
    Figure Legend Snippet: Reconstitution of MMR in vitro. (A) Fractionation of a HeLa nuclear extract into three components required for MMR. (B) Reconstitution of MMR requires SS1, SS2, and FII. The DNA substrate (100 ng of the 5′ G-T heteroduplex) was incubated for 15 min at 37°C in the reaction buffer with fractions as indicated. Amounts of protein used were 15 μg of SS1, 1.5 μg of SS2, or 30 μg of FII. DNA was extracted, treated with Hin dIII and Bsp 106, electrophoresed on an agarose gel, and visualized by ethidium bromide staining under UV illumination. ND, not detectable.

    Techniques Used: In Vitro, Fractionation, Incubation, Agarose Gel Electrophoresis, Staining

    7) Product Images from "NAD+ is not utilized as a co-factor for DNA ligation by human DNA ligase IV"

    Article Title: NAD+ is not utilized as a co-factor for DNA ligation by human DNA ligase IV

    Journal: Nucleic Acids Research

    doi: 10.1093/nar/gkaa1118

    NAD + does not stimulate nick ligation by X4L4. ( A ) Representative gel of nick ligation. X4L4 (50 nM) and T4 DNA ligase (10 units, NEB) were incubated with 100 nM labeled nicked DNA duplex as described in ‘Materials and Methods’ section in the absence or presence of 0.5 mM NAD + and ATP as indicated. The duplex DNA substrate with a single ligatable nick is represented schematically. ( B ) Quantification of nick ligation. Data are represented as the mean ± SD of three independent replicates. Two-tailed Student’s t –test was used for P value calculations.
    Figure Legend Snippet: NAD + does not stimulate nick ligation by X4L4. ( A ) Representative gel of nick ligation. X4L4 (50 nM) and T4 DNA ligase (10 units, NEB) were incubated with 100 nM labeled nicked DNA duplex as described in ‘Materials and Methods’ section in the absence or presence of 0.5 mM NAD + and ATP as indicated. The duplex DNA substrate with a single ligatable nick is represented schematically. ( B ) Quantification of nick ligation. Data are represented as the mean ± SD of three independent replicates. Two-tailed Student’s t –test was used for P value calculations.

    Techniques Used: Ligation, Incubation, Labeling, Two Tailed Test

    8) Product Images from "Cas12a-assisted precise targeted cloning using in vivo Cre-lox recombination"

    Article Title: Cas12a-assisted precise targeted cloning using in vivo Cre-lox recombination

    Journal: Nature Communications

    doi: 10.1038/s41467-021-21275-4

    Characterization of various genomic DNA digestion/DNA assembly combinations in the CAPTURE method. a Schematics of T4 DNA polymerase exo + fill-in DNA assembly. In step 1, DNA molecules ends are chewed back by T4 DNA polymerase to create ssDNA overhangs. The reaction mixture’s temperature is increased to 75 °C to inactivate T4 DNA polymerase and potentially remove ssDNA secondary structures. Temperature is then decreased to 50 °C to allow for ssDNA overhang hybridization. In step 2, by addition of fresh T4 DNA polymerase, and dNTPs, DNA gaps in the hybridized DNA molecule are filled. E. coli DNA ligase is then used to ligate the nicks and produce the final assembly product. b Comparison of different digestion/DNA assembly combinations in cloning four high GC-content BGCs from Actinomycetes. The Fn Cas12a/T4 exo + fill-in strategy showed ~100% cloning efficiency for all four target BGCs. RE: restriction enzymes. For each cloning experiment, at least seven colonies were selected and the purified plasmids from each colony were analyzed by restriction digestion. The cloning efficiencies were calculated as the ratio of correct colonies to the total number of checked colonies. Each experiment was performed in three biological replicates and data are presented as mean values ± standard error (SEM). c Summary of results for cloning uncharacterized BGCs using CAPTURE. BGCs ranging from 10 to 113 kb can be robustly cloned using the CAPTURE method at close to 100% efficiency regardless of their GC-content. Source data are provided as a Source Data file.
    Figure Legend Snippet: Characterization of various genomic DNA digestion/DNA assembly combinations in the CAPTURE method. a Schematics of T4 DNA polymerase exo + fill-in DNA assembly. In step 1, DNA molecules ends are chewed back by T4 DNA polymerase to create ssDNA overhangs. The reaction mixture’s temperature is increased to 75 °C to inactivate T4 DNA polymerase and potentially remove ssDNA secondary structures. Temperature is then decreased to 50 °C to allow for ssDNA overhang hybridization. In step 2, by addition of fresh T4 DNA polymerase, and dNTPs, DNA gaps in the hybridized DNA molecule are filled. E. coli DNA ligase is then used to ligate the nicks and produce the final assembly product. b Comparison of different digestion/DNA assembly combinations in cloning four high GC-content BGCs from Actinomycetes. The Fn Cas12a/T4 exo + fill-in strategy showed ~100% cloning efficiency for all four target BGCs. RE: restriction enzymes. For each cloning experiment, at least seven colonies were selected and the purified plasmids from each colony were analyzed by restriction digestion. The cloning efficiencies were calculated as the ratio of correct colonies to the total number of checked colonies. Each experiment was performed in three biological replicates and data are presented as mean values ± standard error (SEM). c Summary of results for cloning uncharacterized BGCs using CAPTURE. BGCs ranging from 10 to 113 kb can be robustly cloned using the CAPTURE method at close to 100% efficiency regardless of their GC-content. Source data are provided as a Source Data file.

    Techniques Used: Hybridization, Clone Assay, Purification

    Development of the CAPTURE method. a Overview of the workflow. In the first step, purified genomic DNA is digested by Cas12a enzyme to release the target BGC fragment. In the second step, digestion products are mixed with two DNA receivers containing lox P sites at their ends. The target BGC fragment and DNA receivers are assembled together using T4 DNA polymerase exo + fill-in DNA assembly. In the final step, the assembly mixture is transformed into E. coli cells harboring a circularization helper plasmid. The linear DNA is able to circularize in vivo by Cre- lox recombination. b DNA map of helper plasmid pBE14. tcr : tetracycline resistance marker; araBAD : L-arabinose inducible promoter and its regulator; gam : phage lambda Red gam gene; pSC101: temperature-sensitive origin of replication; recA1 : mutated E. coli recA gene to increase transformation efficiency. c Comparison of recombination frequency between Flp (pBE11) and Cre (pBE12) helper plasmids. -: without L-arabinose induction, +: with L-arabinose induction. Recombination frequencies were calculated based on the ratio of white colonies to the total number of acquired colonies. d Linear DNA transformation efficiency for E. coli cells harboring pBE11 (Flp), pBE12 (Cre), pBE14 (Cre and recA1) helper plasmids. Both pBE12 and pBE14 E. coli cells exhibited transformation efficiencies similar to circular DNA. e Comparison of in vitro versus in vivo circularization for two large (50 kb, 73 kb) linear DNA molecules. In vivo circularization showed ~33-fold and 150-fold higher frequency than in vitro circularization for 50 kb and 73 kb molecules, respectively. Circularization frequencies were calculated based on the number of colonies acquired for each circularization experiment in comparison to the number of colonies acquired after transformation of the original circular DNA (see Methods for full description). Each experiment was performed in three biological replicates and data are presented as mean values ± standard deviation (SD). Source data are provided as a Source Data file.
    Figure Legend Snippet: Development of the CAPTURE method. a Overview of the workflow. In the first step, purified genomic DNA is digested by Cas12a enzyme to release the target BGC fragment. In the second step, digestion products are mixed with two DNA receivers containing lox P sites at their ends. The target BGC fragment and DNA receivers are assembled together using T4 DNA polymerase exo + fill-in DNA assembly. In the final step, the assembly mixture is transformed into E. coli cells harboring a circularization helper plasmid. The linear DNA is able to circularize in vivo by Cre- lox recombination. b DNA map of helper plasmid pBE14. tcr : tetracycline resistance marker; araBAD : L-arabinose inducible promoter and its regulator; gam : phage lambda Red gam gene; pSC101: temperature-sensitive origin of replication; recA1 : mutated E. coli recA gene to increase transformation efficiency. c Comparison of recombination frequency between Flp (pBE11) and Cre (pBE12) helper plasmids. -: without L-arabinose induction, +: with L-arabinose induction. Recombination frequencies were calculated based on the ratio of white colonies to the total number of acquired colonies. d Linear DNA transformation efficiency for E. coli cells harboring pBE11 (Flp), pBE12 (Cre), pBE14 (Cre and recA1) helper plasmids. Both pBE12 and pBE14 E. coli cells exhibited transformation efficiencies similar to circular DNA. e Comparison of in vitro versus in vivo circularization for two large (50 kb, 73 kb) linear DNA molecules. In vivo circularization showed ~33-fold and 150-fold higher frequency than in vitro circularization for 50 kb and 73 kb molecules, respectively. Circularization frequencies were calculated based on the number of colonies acquired for each circularization experiment in comparison to the number of colonies acquired after transformation of the original circular DNA (see Methods for full description). Each experiment was performed in three biological replicates and data are presented as mean values ± standard deviation (SD). Source data are provided as a Source Data file.

    Techniques Used: Purification, Transformation Assay, Plasmid Preparation, In Vivo, Marker, In Vitro, Standard Deviation

    9) Product Images from "PlasmidMaker: a Versatile, Automated, and High Throughput End-to-End Platform for Plasmid Construction"

    Article Title: PlasmidMaker: a Versatile, Automated, and High Throughput End-to-End Platform for Plasmid Construction

    Journal: bioRxiv

    doi: 10.1101/2021.12.31.474679

    Overview of the workflow of the automated DNA assembly process. a) An online ordering system receives plasmid orders to be assembled. The received plasmid sequence is annotated and processed to generate primers, guides, and liquid handling worklists. b) Algorithm for design of DNA guides and choice of enzyme for efficient Pf Ago-based assembly. Using annotated fragments of the plasmid as input, guide search space is created from the junction of the fragments. Based on the GC content of the fragments and guide search space, a suggestion is provided to use either WT Pf Ago and Pf Ago* or only Pf Ago*. The guide library created from the guide search space is filtered based on the design rules for Pf Ago digestion and Hifi Taq ligation to identify 24 bp recognition sequences for high-fidelity plasmid assembly. c) Workflow for generation of liquid handling worklists for Pf Ago-based plasmid assembly. Using reference csv file containing the location of primers/guides in 96-well/384-well plate for each plasmid, liquid handling steps are created for mixing primers and templates, mixing guides, followed by equimolar mixing of purified PCR fragments.
    Figure Legend Snippet: Overview of the workflow of the automated DNA assembly process. a) An online ordering system receives plasmid orders to be assembled. The received plasmid sequence is annotated and processed to generate primers, guides, and liquid handling worklists. b) Algorithm for design of DNA guides and choice of enzyme for efficient Pf Ago-based assembly. Using annotated fragments of the plasmid as input, guide search space is created from the junction of the fragments. Based on the GC content of the fragments and guide search space, a suggestion is provided to use either WT Pf Ago and Pf Ago* or only Pf Ago*. The guide library created from the guide search space is filtered based on the design rules for Pf Ago digestion and Hifi Taq ligation to identify 24 bp recognition sequences for high-fidelity plasmid assembly. c) Workflow for generation of liquid handling worklists for Pf Ago-based plasmid assembly. Using reference csv file containing the location of primers/guides in 96-well/384-well plate for each plasmid, liquid handling steps are created for mixing primers and templates, mixing guides, followed by equimolar mixing of purified PCR fragments.

    Techniques Used: Plasmid Preparation, Sequencing, Ligation, Purification, Polymerase Chain Reaction

    Characterization of Pf Ago/AREs capabilities in creation of user defined sticky ends of different sizes on linear DNA ends. a) The strategy used for creation of varying sticky end sizes on DNA ends. One guide DNA was designed to target the lower strand of the dsDNA molecule and create a nick after its 10 th nucleotide position. This guide DNA was kept constant for creation of all sticky end sizes. The second guide DNA was designed to target the upper strand of the dsDNA. By changing the second guide’s position, sticky ends of varying sizes can be created. b) Characterization of Pf Ago/AREs cleavage efficiencies in creation of 5-12 nt sticky ends on linear dsDNA molecules ends. amp and CrtI genes were amplified by PCR to share 24 bp sequence homology at their ends. The homology sequence is shown in part a . The amplified fragments were digested by Pf Ago/AREs and ligated by T4 DNA ligase. The assembly product was then analyzed using agarose gel electrophoresis. Except for 10 nt sticky ends, all other sticky ends can be efficiently used for DNA assembly applications. M: 1 kb DNA ladder.
    Figure Legend Snippet: Characterization of Pf Ago/AREs capabilities in creation of user defined sticky ends of different sizes on linear DNA ends. a) The strategy used for creation of varying sticky end sizes on DNA ends. One guide DNA was designed to target the lower strand of the dsDNA molecule and create a nick after its 10 th nucleotide position. This guide DNA was kept constant for creation of all sticky end sizes. The second guide DNA was designed to target the upper strand of the dsDNA. By changing the second guide’s position, sticky ends of varying sizes can be created. b) Characterization of Pf Ago/AREs cleavage efficiencies in creation of 5-12 nt sticky ends on linear dsDNA molecules ends. amp and CrtI genes were amplified by PCR to share 24 bp sequence homology at their ends. The homology sequence is shown in part a . The amplified fragments were digested by Pf Ago/AREs and ligated by T4 DNA ligase. The assembly product was then analyzed using agarose gel electrophoresis. Except for 10 nt sticky ends, all other sticky ends can be efficiently used for DNA assembly applications. M: 1 kb DNA ladder.

    Techniques Used: Amplification, Polymerase Chain Reaction, Sequencing, Agarose Gel Electrophoresis

    10) Product Images from "Escherichia coli β-clamp slows down DNA polymerase I dependent nick translation while accelerating ligation"

    Article Title: Escherichia coli β-clamp slows down DNA polymerase I dependent nick translation while accelerating ligation

    Journal: bioRxiv

    doi: 10.1101/256537

    β-clamp inhibits the contact between the 5’end of nicked DNA and Pol-I. ( A ) Structural model of E. coli Klenow /DNA complex shows that finger domain of Klenow makes a contact with downstream nick site. The model shows that the conserved F771 (in blue surface representation), which participates in the strand displacement, is positioned between the downstream nicked DNA strands. ( B ) A fraction of BrdU base containing radiolabelled 28 bases oligonucleotide shows a gel shift under the near UV light, suggesting that the oligonucleotide makes cross-linked adduct with Klenow during strand displacement (lane 2). The presence of β-clamp and γ-complex in the absence of ATP (lane 3), or ATP alone (lane 4) also exhibited similar crosslinking adduct. However, β-clamp, γ-complex, and ATP together suppressed the formation of crosslinking product (lane 6), suggesting that loading of the β-clamp on the template blocks the crosslinking. * Represents a minor contaminated band with the custom synthesized 28 bases oligonucleotide.
    Figure Legend Snippet: β-clamp inhibits the contact between the 5’end of nicked DNA and Pol-I. ( A ) Structural model of E. coli Klenow /DNA complex shows that finger domain of Klenow makes a contact with downstream nick site. The model shows that the conserved F771 (in blue surface representation), which participates in the strand displacement, is positioned between the downstream nicked DNA strands. ( B ) A fraction of BrdU base containing radiolabelled 28 bases oligonucleotide shows a gel shift under the near UV light, suggesting that the oligonucleotide makes cross-linked adduct with Klenow during strand displacement (lane 2). The presence of β-clamp and γ-complex in the absence of ATP (lane 3), or ATP alone (lane 4) also exhibited similar crosslinking adduct. However, β-clamp, γ-complex, and ATP together suppressed the formation of crosslinking product (lane 6), suggesting that loading of the β-clamp on the template blocks the crosslinking. * Represents a minor contaminated band with the custom synthesized 28 bases oligonucleotide.

    Techniques Used: Electrophoretic Mobility Shift Assay, Synthesized

    β-clamp inhibits strand displacement activity of exo - Klenow and exo - Pol I ( A ) The urea PAGE shows that exo - Klenow exhibits a slower speed of strand displacement. Presence of the template-loaded β-clamp stalls the exo - Klenow pauses mostly at the 39 th nucleotide position. The exo - Klenow-mediated strand displacement coupled with ligation represents that the DNA ligase fails to ligate nicks. However, the presence of template-loaded β-clamp increases the ligation frequency. ( B ) exo - Pol I exhibits similar functional consequences on the strand displacement and ligation as observed in case of exo - Klenow (panel A). The values representing mean ± sd are calculated from three different experiments.
    Figure Legend Snippet: β-clamp inhibits strand displacement activity of exo - Klenow and exo - Pol I ( A ) The urea PAGE shows that exo - Klenow exhibits a slower speed of strand displacement. Presence of the template-loaded β-clamp stalls the exo - Klenow pauses mostly at the 39 th nucleotide position. The exo - Klenow-mediated strand displacement coupled with ligation represents that the DNA ligase fails to ligate nicks. However, the presence of template-loaded β-clamp increases the ligation frequency. ( B ) exo - Pol I exhibits similar functional consequences on the strand displacement and ligation as observed in case of exo - Klenow (panel A). The values representing mean ± sd are calculated from three different experiments.

    Techniques Used: Activity Assay, Polyacrylamide Gel Electrophoresis, Ligation, Functional Assay

    β-clamp inhibits Pol I-mediated nick translation. ( A ) The template used in the assay was prepared by assembling 67 bases, 5’-phosphorylated 28 bases and 5’-radiolabelled (asterisk) 19 bases oligonucleotides. ( B ) A representative urea denaturing PAGE and the rate calculation therein (see also S2 Fig ) indicate that the presence of β-clamp nominally affected nick translation traversing 28 bases downstream RNA. ( C ) The urea-denaturing gel shows that Pol I efficiently translated the nick, degrading 28 bases downstream DNA, but the presence of β-clamp dramatically reduced the speed of nick translation. The presence of ligase allowed early ligation when β-clamp slowed down Pol I-mediated nick translation. ( D ) The urea PAGE represents that the 3’-exo - Pol I-mediated nick translation was slowed down in the presence of template-loaded β-clamp, in a manner similar to the nick translation with β-clamp and Pol I combinations, as shown in panel C. Moreover, the presence of template-loaded β-clamp increases the ligation efficiency during the 3’-exo - Pol I-mediated nick translation. (E) The urea denaturing gel represents that the 5’-exo - Pol I intensely paused at the 39 th nucleotide and downstream positions, affecting the ligation process. The presence of template-loaded β-clamp moderately enhances these pauses, but facilitates ligation to some extent. The # indicates the position of truncated by-products that originated from the 67-nucleotide long product by the 3’ exonuclease function of Pol I and 5’-exo - Pol I (panel B, C, E). This truncated product is missing in the assay with 3’-exo - Pol I (panel D). All the experiments were performed at least three times. The values represent mean ± standard deviation (sd).
    Figure Legend Snippet: β-clamp inhibits Pol I-mediated nick translation. ( A ) The template used in the assay was prepared by assembling 67 bases, 5’-phosphorylated 28 bases and 5’-radiolabelled (asterisk) 19 bases oligonucleotides. ( B ) A representative urea denaturing PAGE and the rate calculation therein (see also S2 Fig ) indicate that the presence of β-clamp nominally affected nick translation traversing 28 bases downstream RNA. ( C ) The urea-denaturing gel shows that Pol I efficiently translated the nick, degrading 28 bases downstream DNA, but the presence of β-clamp dramatically reduced the speed of nick translation. The presence of ligase allowed early ligation when β-clamp slowed down Pol I-mediated nick translation. ( D ) The urea PAGE represents that the 3’-exo - Pol I-mediated nick translation was slowed down in the presence of template-loaded β-clamp, in a manner similar to the nick translation with β-clamp and Pol I combinations, as shown in panel C. Moreover, the presence of template-loaded β-clamp increases the ligation efficiency during the 3’-exo - Pol I-mediated nick translation. (E) The urea denaturing gel represents that the 5’-exo - Pol I intensely paused at the 39 th nucleotide and downstream positions, affecting the ligation process. The presence of template-loaded β-clamp moderately enhances these pauses, but facilitates ligation to some extent. The # indicates the position of truncated by-products that originated from the 67-nucleotide long product by the 3’ exonuclease function of Pol I and 5’-exo - Pol I (panel B, C, E). This truncated product is missing in the assay with 3’-exo - Pol I (panel D). All the experiments were performed at least three times. The values represent mean ± standard deviation (sd).

    Techniques Used: Nick Translation, Polyacrylamide Gel Electrophoresis, Ligation, Standard Deviation

    11) Product Images from "A Method for Selectively Enriching Microbial DNA from Contaminating Vertebrate Host DNA"

    Article Title: A Method for Selectively Enriching Microbial DNA from Contaminating Vertebrate Host DNA

    Journal: PLoS ONE

    doi: 10.1371/journal.pone.0076096

    MBD-Fc enriches E. coli DNA from mixed E. coli and human DNA (IMR-90) samples. Graphs showing the percentage of mapped reads from Ion Torrent PGM experiments to either the E. coli MG1655 or human hg19 reference genome from libraries made with different ratios of human to E. coli DNA. The ratio between E. coli to human DNA in the premixed samples is indicated above the figure. “Unenriched” refers to untreated, control mixtures. “Bound” indicates DNA that remained bound to MBD-Fc beads and “Enriched” corresponds to unbound DNA remaining in the supernatant.
    Figure Legend Snippet: MBD-Fc enriches E. coli DNA from mixed E. coli and human DNA (IMR-90) samples. Graphs showing the percentage of mapped reads from Ion Torrent PGM experiments to either the E. coli MG1655 or human hg19 reference genome from libraries made with different ratios of human to E. coli DNA. The ratio between E. coli to human DNA in the premixed samples is indicated above the figure. “Unenriched” refers to untreated, control mixtures. “Bound” indicates DNA that remained bound to MBD-Fc beads and “Enriched” corresponds to unbound DNA remaining in the supernatant.

    Techniques Used:

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    New England Biolabs e coli dna ligase
    Individual replication fork progression is independent of primase A Micrographs showing replication products at 10 min where: i , all components present; or a component omitted: ii, DnaB and DnaC810; iii, Pol III*; iv, β; v, <t>SSB;</t> vi, primase. Composite, false-colored fields show anchor points for molecules that contain ssDNA, except i or v , where only long products were seen. In vi , surfaces were sparsely populated with <t>DNA</t> to avoid any ambiguity in molecule identification. Cyan, fields with flow off; magenta, same field with flow on showing fully-extended molecules. Molecules are bracketed for clarity. Scale bar: 10 μm, equal to 33.9 kb dsDNA or 80.3 knt SSB-bound ssDNA at 4,000 μl/h, without Mg 2+ ). B. Cartoon showing leading strand only product in a reaction lacking primase. C. Composite, false-colored image showing leading strand only replication without primase. Three replicating molecules ( 1, 2, 3 ). Molecules a, b and c are referred to later. D. Time-lapse, at 50-second intervals, of Molecules 1, 2 and 3 identified in C , colored by time-point as per C . E. Kymographs of molecules, numbered per C and D , showing fork progression without primase. Dashed grey line: position of anchor. Linear fits are from initiation to termination, yielding average fork rates. Pauses are included in the average here. F. Histograms of fork progression rates in the presence (grey) and absence of primase (light blue). Histograms fit to single Gaussians ( R 2 : with primase, 0.80; without primase, 0.94); no outliers were rejected. n , molecules. G. Processivities of single replisomes from live imaging experiments. Whisker plots of molecule lengths, with (320 nM) or without primase and/or β in flow. Data from 2 (primase, no β) or 3 (others) experiments. Horizontal bars, median; vertical bars, interquartile range. ***, significantly different pairs of populations (Kruskal-Wallis; P
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    Individual replication fork progression is independent of primase A Micrographs showing replication products at 10 min where: i , all components present; or a component omitted: ii, DnaB and DnaC810; iii, Pol III*; iv, β; v, SSB; vi, primase. Composite, false-colored fields show anchor points for molecules that contain ssDNA, except i or v , where only long products were seen. In vi , surfaces were sparsely populated with DNA to avoid any ambiguity in molecule identification. Cyan, fields with flow off; magenta, same field with flow on showing fully-extended molecules. Molecules are bracketed for clarity. Scale bar: 10 μm, equal to 33.9 kb dsDNA or 80.3 knt SSB-bound ssDNA at 4,000 μl/h, without Mg 2+ ). B. Cartoon showing leading strand only product in a reaction lacking primase. C. Composite, false-colored image showing leading strand only replication without primase. Three replicating molecules ( 1, 2, 3 ). Molecules a, b and c are referred to later. D. Time-lapse, at 50-second intervals, of Molecules 1, 2 and 3 identified in C , colored by time-point as per C . E. Kymographs of molecules, numbered per C and D , showing fork progression without primase. Dashed grey line: position of anchor. Linear fits are from initiation to termination, yielding average fork rates. Pauses are included in the average here. F. Histograms of fork progression rates in the presence (grey) and absence of primase (light blue). Histograms fit to single Gaussians ( R 2 : with primase, 0.80; without primase, 0.94); no outliers were rejected. n , molecules. G. Processivities of single replisomes from live imaging experiments. Whisker plots of molecule lengths, with (320 nM) or without primase and/or β in flow. Data from 2 (primase, no β) or 3 (others) experiments. Horizontal bars, median; vertical bars, interquartile range. ***, significantly different pairs of populations (Kruskal-Wallis; P

    Journal: Cell

    Article Title: Independent and Stochastic Action of DNA Polymerases in the Replisome

    doi: 10.1016/j.cell.2017.05.041

    Figure Lengend Snippet: Individual replication fork progression is independent of primase A Micrographs showing replication products at 10 min where: i , all components present; or a component omitted: ii, DnaB and DnaC810; iii, Pol III*; iv, β; v, SSB; vi, primase. Composite, false-colored fields show anchor points for molecules that contain ssDNA, except i or v , where only long products were seen. In vi , surfaces were sparsely populated with DNA to avoid any ambiguity in molecule identification. Cyan, fields with flow off; magenta, same field with flow on showing fully-extended molecules. Molecules are bracketed for clarity. Scale bar: 10 μm, equal to 33.9 kb dsDNA or 80.3 knt SSB-bound ssDNA at 4,000 μl/h, without Mg 2+ ). B. Cartoon showing leading strand only product in a reaction lacking primase. C. Composite, false-colored image showing leading strand only replication without primase. Three replicating molecules ( 1, 2, 3 ). Molecules a, b and c are referred to later. D. Time-lapse, at 50-second intervals, of Molecules 1, 2 and 3 identified in C , colored by time-point as per C . E. Kymographs of molecules, numbered per C and D , showing fork progression without primase. Dashed grey line: position of anchor. Linear fits are from initiation to termination, yielding average fork rates. Pauses are included in the average here. F. Histograms of fork progression rates in the presence (grey) and absence of primase (light blue). Histograms fit to single Gaussians ( R 2 : with primase, 0.80; without primase, 0.94); no outliers were rejected. n , molecules. G. Processivities of single replisomes from live imaging experiments. Whisker plots of molecule lengths, with (320 nM) or without primase and/or β in flow. Data from 2 (primase, no β) or 3 (others) experiments. Horizontal bars, median; vertical bars, interquartile range. ***, significantly different pairs of populations (Kruskal-Wallis; P

    Article Snippet: Replication products were pulse-labeled by injecting 50 μl of a mix comprising: 30 mM Tris-Cl (pH 8.0 at 25°C), 10 mM magnesium acetate, 10 μg/ml BSA, 40 μM each of dATP, dCTP and dGTP, 20 μM digoxygenin-11-dUTP (alkali-stable, Roche), 26 μM NAD+ , 3.3 U DNA Pol I (wild-type, full-length, NEB), 3.3 U E. coli DNA ligase (NEB), and 250 nM SSB (as tetramer), at a flow-rate of 500 μl•h−1 for 5 min. At 5 min, a further 50 μl SMB + 1 M NaCl was introduced to remove bound Pol I and ligase, immediately followed by a further wash of 400 μl WB at 4,000 μl•h−1 , again without stopping flow.

    Techniques: Flow Cytometry, Imaging, Whisker Assay

    Map of M13mp18 and f1PM based heteroduplex substrates. a The map of bacteriophage M13mp18 replicative form (RF) DNA shows restriction enzyme sites relevant to this study with derivatives M13LR1 and M13LR3 containing 22-bp insertions at the unique HindIII restriction site, and phage M13WX1 and M13X22 containing 26-bp and 22-bp insertions at Xba I site respectively. b The map of bacteriophage f1PM RF DNA with its derivative f1PMA with a 27-bp insertion at Xba I. ‘V’, phage viral strand. ‘C’, phage complementary strand. Underlines beneath each viral strand are the original insertion sequences. The C-strand from parental phage RF DNA was paired with viral strand of its insertion derivative to produce gapped duplex DNA, and the gap was sealed with dI or deoxyuridine containing synthetic oligodeoxyribonucleotide. A-I, C-I, G-I, T-I, and G-U are the resulting substrates and DNA sequence shown on each C-strand of the the synthetic linker used. In the presence of dI, the substrates were refractory to the restriction endonuclease scoring. After the repair, DNA products become sensitive to restriction endonuclease cleavage. The recognition sequence of corresponding restriction endonuclease markers for repair products are shown in bold on V-strands

    Journal: Cell & Bioscience

    Article Title: Deoxyinosine repair in nuclear extracts of human cells

    doi: 10.1186/s13578-015-0044-8

    Figure Lengend Snippet: Map of M13mp18 and f1PM based heteroduplex substrates. a The map of bacteriophage M13mp18 replicative form (RF) DNA shows restriction enzyme sites relevant to this study with derivatives M13LR1 and M13LR3 containing 22-bp insertions at the unique HindIII restriction site, and phage M13WX1 and M13X22 containing 26-bp and 22-bp insertions at Xba I site respectively. b The map of bacteriophage f1PM RF DNA with its derivative f1PMA with a 27-bp insertion at Xba I. ‘V’, phage viral strand. ‘C’, phage complementary strand. Underlines beneath each viral strand are the original insertion sequences. The C-strand from parental phage RF DNA was paired with viral strand of its insertion derivative to produce gapped duplex DNA, and the gap was sealed with dI or deoxyuridine containing synthetic oligodeoxyribonucleotide. A-I, C-I, G-I, T-I, and G-U are the resulting substrates and DNA sequence shown on each C-strand of the the synthetic linker used. In the presence of dI, the substrates were refractory to the restriction endonuclease scoring. After the repair, DNA products become sensitive to restriction endonuclease cleavage. The recognition sequence of corresponding restriction endonuclease markers for repair products are shown in bold on V-strands

    Article Snippet: E. coli DNA ligase, T4 polynucleotide kinase, HindIII-HFTM and other restriction endonucleases were obtained from New England Biolabs.

    Techniques: Sequencing

    Transcriptional Maps of the Nucleosome Reveal that H2A.Z Enhances the Width and uH2B the Height of the Barrier (A) Median residence time histograms of Pol II transcription through bare NPS DNA (black), xWT (orange), hWT (red), H2A.Z (blue) and uH2B (green) nucleosomes. Bar width is 1 bp and major peak positions are labeled (in bp) above the corresponding peaks. NPS entry, dyad, NPS exit are marked with blue dashed lines. The polar plots on the right are the corresponding transcriptional maps of the nucleosome, formed by projecting the residence time histogram onto the surface of nucleosomal DNA. The top axis (red) indicates corresponding positions of the first half of nucleosome expressed as superhelical locations (SHL). n = 35, 23, 26, 21, 31, respectively for NPS DNA, xWT, hWT, H2A.Z and uH2B nucleosomes. (B) Crossing time (total time Pol II takes to cross the entire nucleosome region) distributions plotted using the complementary cumulative distribution function (CCDF, fraction of events longer than a given crossing time). Crossing times of Bare NPS DNA, Xenopus WT (xWT), human WT (hWT), uH2B and H2A.Z nucleosomes are plotted in black, orange, red, green and blue, respectively. See also Figure S6 on statistics of the crossing time, crossing probability, pause-free velocity and arrest position.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: Transcriptional Maps of the Nucleosome Reveal that H2A.Z Enhances the Width and uH2B the Height of the Barrier (A) Median residence time histograms of Pol II transcription through bare NPS DNA (black), xWT (orange), hWT (red), H2A.Z (blue) and uH2B (green) nucleosomes. Bar width is 1 bp and major peak positions are labeled (in bp) above the corresponding peaks. NPS entry, dyad, NPS exit are marked with blue dashed lines. The polar plots on the right are the corresponding transcriptional maps of the nucleosome, formed by projecting the residence time histogram onto the surface of nucleosomal DNA. The top axis (red) indicates corresponding positions of the first half of nucleosome expressed as superhelical locations (SHL). n = 35, 23, 26, 21, 31, respectively for NPS DNA, xWT, hWT, H2A.Z and uH2B nucleosomes. (B) Crossing time (total time Pol II takes to cross the entire nucleosome region) distributions plotted using the complementary cumulative distribution function (CCDF, fraction of events longer than a given crossing time). Crossing times of Bare NPS DNA, Xenopus WT (xWT), human WT (hWT), uH2B and H2A.Z nucleosomes are plotted in black, orange, red, green and blue, respectively. See also Figure S6 on statistics of the crossing time, crossing probability, pause-free velocity and arrest position.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques: Labeling

    Topography Maps of the Nucleosome Revealed by Nucleosome Unzipping at Constant Force (A) Representative unzipping traces of bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes at 28 pN constant force. Unzipped bp are normalized to the beginning of the second NPS. Dashed lines mark entry, dyad and exit regions of the second NPS. Traces are shifted horizontally for clarity. (B) Mean residence time (RT) histogram of the unzipping fork along bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes during unzipping at a constant force of 28 pN. Bare NPS RTs are too short to see on the axes shown. Unzipped bp are normalized to the beginning of the second NPS core. Major peak positions are indicated above each peak (in bp). n = 33, 17, 20, 20, respectively for NPS DNA, WT, H2A.Z and uH2B nucleosomes. See also Figure S2 on assembly cooperativity of H2A.Z nucleosomes.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: Topography Maps of the Nucleosome Revealed by Nucleosome Unzipping at Constant Force (A) Representative unzipping traces of bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes at 28 pN constant force. Unzipped bp are normalized to the beginning of the second NPS. Dashed lines mark entry, dyad and exit regions of the second NPS. Traces are shifted horizontally for clarity. (B) Mean residence time (RT) histogram of the unzipping fork along bare NPS DNA (black), WT (red), H2A.Z (blue) and uH2B (green) nucleosomes during unzipping at a constant force of 28 pN. Bare NPS RTs are too short to see on the axes shown. Unzipped bp are normalized to the beginning of the second NPS core. Major peak positions are indicated above each peak (in bp). n = 33, 17, 20, 20, respectively for NPS DNA, WT, H2A.Z and uH2B nucleosomes. See also Figure S2 on assembly cooperativity of H2A.Z nucleosomes.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques:

    Mechanical Model for Pol II Transcription Through the Nucleosome (A) Schematic of the mechanical model, showing three different lengths of unwrapped DNA for a given polymerase position along the DNA sequence. The steric spheres are shown in purple (polymerase) and beige (nucleosome), while the DNA is shown as a tube. (i) shows a configuration with a short, sharply bent DNA linker connecting Pol II and the nucleosome, which are in contact and sterically pushing on each other. (ii) shows a medium-length straighter linker, with Pol II still pushing on the nucleosome. (iii) shows a long straight linker without contact between Pol II and the nucleosome. Linker DNA color corresponds to overall energy for each configuration (given in C). Black arrows represent tangent orientations of the DNA backbone at the point of polymerase binding (top) and for the last contact with the nucleosome (bottom). Linker length and bending angle (between indicated tangents) are labeled on each polymerase-nucleosome pair. (B) Model of Pol II dynamics. Pairs (p,q) indicate the Pol II state: p indicates the length of the RNA transcript, and q the number of base pairs backtracked from the most recent main pathway state. Pol II steps forward one base pair with rate k 0 or can enter a backtracked pathway by stepping backward one base pair at rate k b1 . From backtracked positions, Pol II can move forward a base pair with rate k f n or can backtrack another base pair at rate k b n . Moving forward from the first backtracked state returns Pol II to the main pathway. (C) Energy landscape of nucleosome-Pol II interaction, for constant DNA-nucleosome interaction energies of 1k B T per base pair. DNA unwrapping decreases the DNA linker conformational energy, while removing favorable DNA-nucleosome interactions, overall providing a minimum energy a few base pairs ahead of the front edge of Pol II. Forward Pol II steps are unfavorable as they shorten the DNA linker. Points i , ii , and iii correspond to configurations illustrated in A. Inset shows cross-section of energy landscape at Pol II position of 47 bp, highlighting the minimum in the energy landscape a few bps ahead of Pol II, at ∼52 bps unwrapped. Pol II progress through the nucleosome is defined as the position of the Pol II center plus an additional 17 bp for consistency with the transcribed distance in Figure 6 . (D) Dwell time profiles for human WT, H2A.Z, and uH2B nucleosomes. Solid black lines are experimental mean dwell times and colored dotted lines are the best fitted mean dwell times according to the mechanical model. (E) Estimated DNA-octamer interaction energy profiles for human WT, H2A.Z, and uH2B nucleosomes. The energy values are found such that they give the best fitted dwell times shown in (D). Peak positions referenced in the text are labeled in bp, relative to the start of the NPS. See also Figure S7 for fitting of nucleosome energy profiles based on Pol II dwell times.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: Mechanical Model for Pol II Transcription Through the Nucleosome (A) Schematic of the mechanical model, showing three different lengths of unwrapped DNA for a given polymerase position along the DNA sequence. The steric spheres are shown in purple (polymerase) and beige (nucleosome), while the DNA is shown as a tube. (i) shows a configuration with a short, sharply bent DNA linker connecting Pol II and the nucleosome, which are in contact and sterically pushing on each other. (ii) shows a medium-length straighter linker, with Pol II still pushing on the nucleosome. (iii) shows a long straight linker without contact between Pol II and the nucleosome. Linker DNA color corresponds to overall energy for each configuration (given in C). Black arrows represent tangent orientations of the DNA backbone at the point of polymerase binding (top) and for the last contact with the nucleosome (bottom). Linker length and bending angle (between indicated tangents) are labeled on each polymerase-nucleosome pair. (B) Model of Pol II dynamics. Pairs (p,q) indicate the Pol II state: p indicates the length of the RNA transcript, and q the number of base pairs backtracked from the most recent main pathway state. Pol II steps forward one base pair with rate k 0 or can enter a backtracked pathway by stepping backward one base pair at rate k b1 . From backtracked positions, Pol II can move forward a base pair with rate k f n or can backtrack another base pair at rate k b n . Moving forward from the first backtracked state returns Pol II to the main pathway. (C) Energy landscape of nucleosome-Pol II interaction, for constant DNA-nucleosome interaction energies of 1k B T per base pair. DNA unwrapping decreases the DNA linker conformational energy, while removing favorable DNA-nucleosome interactions, overall providing a minimum energy a few base pairs ahead of the front edge of Pol II. Forward Pol II steps are unfavorable as they shorten the DNA linker. Points i , ii , and iii correspond to configurations illustrated in A. Inset shows cross-section of energy landscape at Pol II position of 47 bp, highlighting the minimum in the energy landscape a few bps ahead of Pol II, at ∼52 bps unwrapped. Pol II progress through the nucleosome is defined as the position of the Pol II center plus an additional 17 bp for consistency with the transcribed distance in Figure 6 . (D) Dwell time profiles for human WT, H2A.Z, and uH2B nucleosomes. Solid black lines are experimental mean dwell times and colored dotted lines are the best fitted mean dwell times according to the mechanical model. (E) Estimated DNA-octamer interaction energy profiles for human WT, H2A.Z, and uH2B nucleosomes. The energy values are found such that they give the best fitted dwell times shown in (D). Peak positions referenced in the text are labeled in bp, relative to the start of the NPS. See also Figure S7 for fitting of nucleosome energy profiles based on Pol II dwell times.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques: Sequencing, Binding Assay, Labeling

    Unzipping Traces of Single Human WT, H2A.Z, M3_M7, uH2B Nucleosomes and Tetrasomes. (A-E) Representative unzipping traces of WT nucleosomes (A), tetrasomes (B), H2A.Z nucleosomes (C), M3_M7 nucleosomes (D) and uH2B nucleosomes (E). Rezipping traces are not shown and they match bare NPS DNA rezipping traces. The unzipped bp (basepairs) are normalized to the beginning of the second NPS core. (F) Number of transitions per trace at the second NPS region. H2A.Z nucleosomes have on average one more transition per trace than WT or uH2B nucleosomes. A transition event is counted when the residence time peak is above an arbitrary threshold. (G-H) Partial unzipping of H2A.Z (G) and WT (H) nucleosomes reveals no lateral mobility induced by multiple rounds of unzipping-rezipping. The unzipping fork repeatedly propagates to the proximal dimer region followed by rezipping (not shown for clarity). The inset shows zoomed-in view of the boxed region, where the position of initial force rise remains unchanged. The dwelling of the unzipping fork in alternative positions (labeled above the dashed lines in bp) is consistent with hopping observed in this region. (I) Native PAGE gels showing homogenous WT, H2A.Z and uH2B nucleosome samples used for single-molecule unzipping experiments.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: Unzipping Traces of Single Human WT, H2A.Z, M3_M7, uH2B Nucleosomes and Tetrasomes. (A-E) Representative unzipping traces of WT nucleosomes (A), tetrasomes (B), H2A.Z nucleosomes (C), M3_M7 nucleosomes (D) and uH2B nucleosomes (E). Rezipping traces are not shown and they match bare NPS DNA rezipping traces. The unzipped bp (basepairs) are normalized to the beginning of the second NPS core. (F) Number of transitions per trace at the second NPS region. H2A.Z nucleosomes have on average one more transition per trace than WT or uH2B nucleosomes. A transition event is counted when the residence time peak is above an arbitrary threshold. (G-H) Partial unzipping of H2A.Z (G) and WT (H) nucleosomes reveals no lateral mobility induced by multiple rounds of unzipping-rezipping. The unzipping fork repeatedly propagates to the proximal dimer region followed by rezipping (not shown for clarity). The inset shows zoomed-in view of the boxed region, where the position of initial force rise remains unchanged. The dwelling of the unzipping fork in alternative positions (labeled above the dashed lines in bp) is consistent with hopping observed in this region. (I) Native PAGE gels showing homogenous WT, H2A.Z and uH2B nucleosome samples used for single-molecule unzipping experiments.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques: Labeling, Clear Native PAGE

    Observation of Multiple Nucleosomal States at the Proximal Dimer Region (A) Time traces of number of base pairs unzipped (relative to beginning of the second NPS) with hWT nucleosome for fixed trap separations of 1031 nm, 1045 nm, and 1060 nm (top to bottom). Color indicates increasing trap separation (purple to red), corresponding to clusters in Figure S3F . Grey dashed lines indicate 17, 23, and 28 base pairs unzipped. (B) Probability distributions for the number of DNA bps unzipped, computed from force-extension data in Figure S3F . Each curve is from a different trap separation, matching colors in A and Figure S3F . Distributions are shown for both bare DNA (top) and WT nucleosome (bottom). Vertical black dotted line indicates the start of the second NPS. Vertical grey dashed lines indicate peak positions for bare DNA (with position in bp labeled), showing that WT nucleosome shifts the first peak within the NPS, and gives rise to an additional peak at 28 bp. See Figure S3F for force-extension data. (C) Zoomed-in view of the black dashed box in (B). Peak positions are labeled in bp. (D) DNA unzipping energy computed by assuming the unzipped bp distributions from data in Figure S3F (including distributions in B) correspond to equilibrium Boltzmann statistics. Inset Δ E shows the DNA-octamer interaction energy, computed as the difference between unzipping energies in the presence of WT (red), H2A.Z (blue), and uH2B (green) nucleosomes and unzipping energies for bare DNA only (black). Vertical black dashed lines and * indicate peak positions (labeled in bp). See also Figure S3 on hopping traces and analysis of energy landscape from equilibrium data.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: Observation of Multiple Nucleosomal States at the Proximal Dimer Region (A) Time traces of number of base pairs unzipped (relative to beginning of the second NPS) with hWT nucleosome for fixed trap separations of 1031 nm, 1045 nm, and 1060 nm (top to bottom). Color indicates increasing trap separation (purple to red), corresponding to clusters in Figure S3F . Grey dashed lines indicate 17, 23, and 28 base pairs unzipped. (B) Probability distributions for the number of DNA bps unzipped, computed from force-extension data in Figure S3F . Each curve is from a different trap separation, matching colors in A and Figure S3F . Distributions are shown for both bare DNA (top) and WT nucleosome (bottom). Vertical black dotted line indicates the start of the second NPS. Vertical grey dashed lines indicate peak positions for bare DNA (with position in bp labeled), showing that WT nucleosome shifts the first peak within the NPS, and gives rise to an additional peak at 28 bp. See Figure S3F for force-extension data. (C) Zoomed-in view of the black dashed box in (B). Peak positions are labeled in bp. (D) DNA unzipping energy computed by assuming the unzipped bp distributions from data in Figure S3F (including distributions in B) correspond to equilibrium Boltzmann statistics. Inset Δ E shows the DNA-octamer interaction energy, computed as the difference between unzipping energies in the presence of WT (red), H2A.Z (blue), and uH2B (green) nucleosomes and unzipping energies for bare DNA only (black). Vertical black dashed lines and * indicate peak positions (labeled in bp). See also Figure S3 on hopping traces and analysis of energy landscape from equilibrium data.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques: Labeling

    A ‘Molecular Ruler’ Gauges the Positions of an Elongating Pol II with Near-Basepair Accuracy (A) Experimental design of an improved single-molecule nucleosomal transcription assay. A single biotinylated Pol II (purple molecular structure) is tethered between two optical traps. Pol II transcription is measured as increases in distance between the two beads at 10 pN constant force. The inset box shows the composition of the template, which is constructed by ligating Pol II stalled complex (cyan), the molecular ruler (green), NPS DNA (or nucleosome, yellow-grey), and a short inter-strand crosslinked DNA (for stalling Pol II, red). The ‘molecular ruler’ consists of eight tandem repeats of a 64-bp DNA (green), each harboring a single sequence-encoded pause site. (B) A representative trace of a single Pol II transcribing through a Xenopus WT nucleosome. The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The inset shows a zoomed-in view of the boxed region, highlighting the repeating pause patterns within the ‘molecular ruler’. The grey dashed lines are the predicted pause sites, whereas the short green lines mark the actual pauses of Pol II. (C) Zoomed-in view of Pol II dynamics within the NPS region of (B). The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The right y-axis (in bp) is normalized to the beginning of the NPS. The left y-axis shows regions preceding the dyad as SHL in red. Black arrows indicates representative events of backtracking, pausing, productive elongation, and hopping. Regions corresponding to Pol II located at SHL(-5) and SHL(-1) are indicated with green and cyan dashed lines, with the corresponding Pol II-nucleosome complex structures plotted on top (PDB 6A5P for PolII-SHL(-5), 6A5 T for PolII-SHL(-1)). Pol II, histones, template DNA, non-template DNA are colored in grey, green, red and blue, respectively. See also Figure S4 on detailed characterization of the ‘molecular ruler’.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: A ‘Molecular Ruler’ Gauges the Positions of an Elongating Pol II with Near-Basepair Accuracy (A) Experimental design of an improved single-molecule nucleosomal transcription assay. A single biotinylated Pol II (purple molecular structure) is tethered between two optical traps. Pol II transcription is measured as increases in distance between the two beads at 10 pN constant force. The inset box shows the composition of the template, which is constructed by ligating Pol II stalled complex (cyan), the molecular ruler (green), NPS DNA (or nucleosome, yellow-grey), and a short inter-strand crosslinked DNA (for stalling Pol II, red). The ‘molecular ruler’ consists of eight tandem repeats of a 64-bp DNA (green), each harboring a single sequence-encoded pause site. (B) A representative trace of a single Pol II transcribing through a Xenopus WT nucleosome. The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The inset shows a zoomed-in view of the boxed region, highlighting the repeating pause patterns within the ‘molecular ruler’. The grey dashed lines are the predicted pause sites, whereas the short green lines mark the actual pauses of Pol II. (C) Zoomed-in view of Pol II dynamics within the NPS region of (B). The three black dashed lines indicate NPS entry, dyad and NPS exit, respectively. The right y-axis (in bp) is normalized to the beginning of the NPS. The left y-axis shows regions preceding the dyad as SHL in red. Black arrows indicates representative events of backtracking, pausing, productive elongation, and hopping. Regions corresponding to Pol II located at SHL(-5) and SHL(-1) are indicated with green and cyan dashed lines, with the corresponding Pol II-nucleosome complex structures plotted on top (PDB 6A5P for PolII-SHL(-5), 6A5 T for PolII-SHL(-1)). Pol II, histones, template DNA, non-template DNA are colored in grey, green, red and blue, respectively. See also Figure S4 on detailed characterization of the ‘molecular ruler’.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques: Construct, Sequencing

    High-resolution Trajectories of Individual Pol II Enzymes Transcribing through WT, H2A.Z and uH2B Nucleosomes (A, B) Representative traces of single Pol II enzymes transcribing through single human WT nucleosomes. The grey dotted lines are the pause sites within the ‘molecular ruler’. The inset (black) is the residence time of Pol II within the ‘molecular ruler’, highlighting repeating pausing signatures of Pol II. The three black dashed lines indicate NPS entry, dyad and NPS exit. Relative positions of Pol II on the template DNA are shown as a cartoon on the right. The traces in blue, green, red and cyan are examples of successful nucleosome crossing, while the trace in grey is an example of Pol II arrest in the nucleosome. For comparison, a trace of Pol II transcribing through bare NPS DNA (black) is shown on the left. Zoomed in traces of high-resolution Pol II dynamics within the NPS are shown in (B), highlighting (black arrowheads) long-lived pausing, backtracking and hopping events. The traces are shifted horizontally for clarity. The right y-axis is normalized to the beginning of the NPS, with the major pause positions marked (in bp) on the right. (C, D) Representative traces of single Pol II enzymes transcribing through single human H2A.Z nucleosomes. (C) shows the full traces and (D) is a zoomed-in view of the high-resolution dynamics within the NPS region. (E, F) Representative traces of single Pol II enzymes transcribing through single human uH2B nucleosomes. (E) shows the full traces and (F) is a zoomed-in view of the high-resolution dynamics within the NPS region. See also Figure S5 on backtracking and hopping dynamics.

    Journal: bioRxiv

    Article Title: High-resolution and High-accuracy Topographic and Transcriptional Maps of the Nucleosome Barrier

    doi: 10.1101/641506

    Figure Lengend Snippet: High-resolution Trajectories of Individual Pol II Enzymes Transcribing through WT, H2A.Z and uH2B Nucleosomes (A, B) Representative traces of single Pol II enzymes transcribing through single human WT nucleosomes. The grey dotted lines are the pause sites within the ‘molecular ruler’. The inset (black) is the residence time of Pol II within the ‘molecular ruler’, highlighting repeating pausing signatures of Pol II. The three black dashed lines indicate NPS entry, dyad and NPS exit. Relative positions of Pol II on the template DNA are shown as a cartoon on the right. The traces in blue, green, red and cyan are examples of successful nucleosome crossing, while the trace in grey is an example of Pol II arrest in the nucleosome. For comparison, a trace of Pol II transcribing through bare NPS DNA (black) is shown on the left. Zoomed in traces of high-resolution Pol II dynamics within the NPS are shown in (B), highlighting (black arrowheads) long-lived pausing, backtracking and hopping events. The traces are shifted horizontally for clarity. The right y-axis is normalized to the beginning of the NPS, with the major pause positions marked (in bp) on the right. (C, D) Representative traces of single Pol II enzymes transcribing through single human H2A.Z nucleosomes. (C) shows the full traces and (D) is a zoomed-in view of the high-resolution dynamics within the NPS region. (E, F) Representative traces of single Pol II enzymes transcribing through single human uH2B nucleosomes. (E) shows the full traces and (F) is a zoomed-in view of the high-resolution dynamics within the NPS region. See also Figure S5 on backtracking and hopping dynamics.

    Article Snippet: Pol II sample beads were prepared by ligating the 1 µm 2 kb spacer DNA beads, Pol II stalled complex, 8 × repeat DNA and nucleosome loaded on NPS-Xlink (or bare NPS-Xlink DNA) using E. Coli DNA ligase (NEB) at 16 °C for 2 hours.

    Techniques:

    Characterization of various genomic DNA digestion/DNA assembly combinations in the CAPTURE method. a Schematics of T4 DNA polymerase exo + fill-in DNA assembly. In step 1, DNA molecules ends are chewed back by T4 DNA polymerase to create ssDNA overhangs. The reaction mixture’s temperature is increased to 75 °C to inactivate T4 DNA polymerase and potentially remove ssDNA secondary structures. Temperature is then decreased to 50 °C to allow for ssDNA overhang hybridization. In step 2, by addition of fresh T4 DNA polymerase, and dNTPs, DNA gaps in the hybridized DNA molecule are filled. E. coli DNA ligase is then used to ligate the nicks and produce the final assembly product. b Comparison of different digestion/DNA assembly combinations in cloning four high GC-content BGCs from Actinomycetes. The Fn Cas12a/T4 exo + fill-in strategy showed ~100% cloning efficiency for all four target BGCs. RE: restriction enzymes. For each cloning experiment, at least seven colonies were selected and the purified plasmids from each colony were analyzed by restriction digestion. The cloning efficiencies were calculated as the ratio of correct colonies to the total number of checked colonies. Each experiment was performed in three biological replicates and data are presented as mean values ± standard error (SEM). c Summary of results for cloning uncharacterized BGCs using CAPTURE. BGCs ranging from 10 to 113 kb can be robustly cloned using the CAPTURE method at close to 100% efficiency regardless of their GC-content. Source data are provided as a Source Data file.

    Journal: Nature Communications

    Article Title: Cas12a-assisted precise targeted cloning using in vivo Cre-lox recombination

    doi: 10.1038/s41467-021-21275-4

    Figure Lengend Snippet: Characterization of various genomic DNA digestion/DNA assembly combinations in the CAPTURE method. a Schematics of T4 DNA polymerase exo + fill-in DNA assembly. In step 1, DNA molecules ends are chewed back by T4 DNA polymerase to create ssDNA overhangs. The reaction mixture’s temperature is increased to 75 °C to inactivate T4 DNA polymerase and potentially remove ssDNA secondary structures. Temperature is then decreased to 50 °C to allow for ssDNA overhang hybridization. In step 2, by addition of fresh T4 DNA polymerase, and dNTPs, DNA gaps in the hybridized DNA molecule are filled. E. coli DNA ligase is then used to ligate the nicks and produce the final assembly product. b Comparison of different digestion/DNA assembly combinations in cloning four high GC-content BGCs from Actinomycetes. The Fn Cas12a/T4 exo + fill-in strategy showed ~100% cloning efficiency for all four target BGCs. RE: restriction enzymes. For each cloning experiment, at least seven colonies were selected and the purified plasmids from each colony were analyzed by restriction digestion. The cloning efficiencies were calculated as the ratio of correct colonies to the total number of checked colonies. Each experiment was performed in three biological replicates and data are presented as mean values ± standard error (SEM). c Summary of results for cloning uncharacterized BGCs using CAPTURE. BGCs ranging from 10 to 113 kb can be robustly cloned using the CAPTURE method at close to 100% efficiency regardless of their GC-content. Source data are provided as a Source Data file.

    Article Snippet: Next, 1 µL of 1 mM NAD+ , 0.4 µl of 10 mM dNTPs, 1 µL (3 U) of T4 DNA polymerase, and 1 µL of E. coli DNA ligase were added and after gentle mixing using wide-bore pipette tips, the mixture was incubated at 37 °C for 1 h, 75 °C for 20 min, and stored at 10 °C until transformation.

    Techniques: Hybridization, Clone Assay, Purification

    Development of the CAPTURE method. a Overview of the workflow. In the first step, purified genomic DNA is digested by Cas12a enzyme to release the target BGC fragment. In the second step, digestion products are mixed with two DNA receivers containing lox P sites at their ends. The target BGC fragment and DNA receivers are assembled together using T4 DNA polymerase exo + fill-in DNA assembly. In the final step, the assembly mixture is transformed into E. coli cells harboring a circularization helper plasmid. The linear DNA is able to circularize in vivo by Cre- lox recombination. b DNA map of helper plasmid pBE14. tcr : tetracycline resistance marker; araBAD : L-arabinose inducible promoter and its regulator; gam : phage lambda Red gam gene; pSC101: temperature-sensitive origin of replication; recA1 : mutated E. coli recA gene to increase transformation efficiency. c Comparison of recombination frequency between Flp (pBE11) and Cre (pBE12) helper plasmids. -: without L-arabinose induction, +: with L-arabinose induction. Recombination frequencies were calculated based on the ratio of white colonies to the total number of acquired colonies. d Linear DNA transformation efficiency for E. coli cells harboring pBE11 (Flp), pBE12 (Cre), pBE14 (Cre and recA1) helper plasmids. Both pBE12 and pBE14 E. coli cells exhibited transformation efficiencies similar to circular DNA. e Comparison of in vitro versus in vivo circularization for two large (50 kb, 73 kb) linear DNA molecules. In vivo circularization showed ~33-fold and 150-fold higher frequency than in vitro circularization for 50 kb and 73 kb molecules, respectively. Circularization frequencies were calculated based on the number of colonies acquired for each circularization experiment in comparison to the number of colonies acquired after transformation of the original circular DNA (see Methods for full description). Each experiment was performed in three biological replicates and data are presented as mean values ± standard deviation (SD). Source data are provided as a Source Data file.

    Journal: Nature Communications

    Article Title: Cas12a-assisted precise targeted cloning using in vivo Cre-lox recombination

    doi: 10.1038/s41467-021-21275-4

    Figure Lengend Snippet: Development of the CAPTURE method. a Overview of the workflow. In the first step, purified genomic DNA is digested by Cas12a enzyme to release the target BGC fragment. In the second step, digestion products are mixed with two DNA receivers containing lox P sites at their ends. The target BGC fragment and DNA receivers are assembled together using T4 DNA polymerase exo + fill-in DNA assembly. In the final step, the assembly mixture is transformed into E. coli cells harboring a circularization helper plasmid. The linear DNA is able to circularize in vivo by Cre- lox recombination. b DNA map of helper plasmid pBE14. tcr : tetracycline resistance marker; araBAD : L-arabinose inducible promoter and its regulator; gam : phage lambda Red gam gene; pSC101: temperature-sensitive origin of replication; recA1 : mutated E. coli recA gene to increase transformation efficiency. c Comparison of recombination frequency between Flp (pBE11) and Cre (pBE12) helper plasmids. -: without L-arabinose induction, +: with L-arabinose induction. Recombination frequencies were calculated based on the ratio of white colonies to the total number of acquired colonies. d Linear DNA transformation efficiency for E. coli cells harboring pBE11 (Flp), pBE12 (Cre), pBE14 (Cre and recA1) helper plasmids. Both pBE12 and pBE14 E. coli cells exhibited transformation efficiencies similar to circular DNA. e Comparison of in vitro versus in vivo circularization for two large (50 kb, 73 kb) linear DNA molecules. In vivo circularization showed ~33-fold and 150-fold higher frequency than in vitro circularization for 50 kb and 73 kb molecules, respectively. Circularization frequencies were calculated based on the number of colonies acquired for each circularization experiment in comparison to the number of colonies acquired after transformation of the original circular DNA (see Methods for full description). Each experiment was performed in three biological replicates and data are presented as mean values ± standard deviation (SD). Source data are provided as a Source Data file.

    Article Snippet: Next, 1 µL of 1 mM NAD+ , 0.4 µl of 10 mM dNTPs, 1 µL (3 U) of T4 DNA polymerase, and 1 µL of E. coli DNA ligase were added and after gentle mixing using wide-bore pipette tips, the mixture was incubated at 37 °C for 1 h, 75 °C for 20 min, and stored at 10 °C until transformation.

    Techniques: Purification, Transformation Assay, Plasmid Preparation, In Vivo, Marker, In Vitro, Standard Deviation