RESEARCH ARTICLE

High-resolution and high-accuracy
topographic and transcriptional maps ofthe nucleosome barrier
Zhijie Chen1,2†, Ronen Gabizon1†, Aidan I Brown3, Antony Lee4, Aixin Song5,
César Dı́az-Celis1,2, Craig D Kaplan6, Elena F Koslover3, Tingting Yao5*,
Carlos Bustamante1,2,4,7,8,9,10,11*1

Institute for Quantitative Biosciences-QB3, University of California, Berkeley,
Berkeley, United States; 2Howard Hughes Medical Institute, University of California,
Berkeley, Berkeley, United States; 3Department of Physics, University of California,
San Diego, San Diego, United States; 4Department of Physics, University ofCalifornia, Berkeley, Berkeley, United States; 5Department of Biochemistry andMolecular Biology, Colorado State University, Fort Collins, United States;
6
Department of Biological Sciences, University of Pittsburgh, Pittsburgh, UnitedStates; 7Jason L Choy Laboratory of Single-Molecule Biophysics, University ofCalifornia, Berkeley, Berkeley, United States; 8Biophysics Graduate Group,
University of California, Berkeley, Berkeley, United States; 9Department of
Molecular & Cell Biology, University of California, Berkeley, Berkeley, United States;
10
Department of Chemistry, University of California, Berkeley, Berkeley, UnitedStates; 11Kavli Energy Nanoscience Institute, University of California, Berkeley,
Berkeley, United States*For correspondence:
Tingting.Yao@Colostate.edu (TY);
carlosb@berkeley.edu (CB)†

These authors contributed
equally to this work

Competing interests: The
authors declare that nocompeting interests exist.
Funding: See page 34
Received: 08 May 2019
Accepted: 30 July 2019
Published: 31 July 2019
Reviewing editor: Taekjip Ha,
Johns Hopkins University, UnitedStatesCopyright Chen et al. This
article is distributed under theterms of the Creative Commons
Attribution License, which
permits unrestricted use andredistribution provided that theoriginal author and source arecredited.

Abstract Nucleosomes represent mechanical and energetic barriers that RNA Polymerase II (PolII) must overcome during transcription. A high-resolution description of the barrier topography, its
modulation by epigenetic modifications, and their effects on Pol II nucleosome crossing dynamics,
is still missing. Here, we obtain topographic and transcriptional (Pol II residence time) maps ofcanonical, H2A.Z, and monoubiquitinated H2B (uH2B) nucleosomes at near base-pair resolution andaccuracy. Pol II crossing dynamics are complex, displaying pauses at specific loci, backtracking, andnucleosome hopping between wrapped states. While H2A.Z widens the barrier, uH2B heightens it,
and both modifications greatly lengthen Pol II crossing time. Using the dwell times of Pol II at eachnucleosomal position we extract the energetics of the barrier. The orthogonal barrier modificationsof H2A.Z and uH2B, and their effects on Pol II dynamics rationalize their observed enrichment in +1
nucleosomes and suggest a mechanism for selective control of gene expression.
DOI: https://doi.org/10.7554/eLife.48281.001

Introduction
The organization of genomic DNA into nucleosomes represents the main physical barrier to transcription by Pol II and constitutes a fundamental mechanism for regulation of gene expression in
eukaryotes. In canonical (hereafter referred to as WT) nucleosomes, a core histone octamer made up
of two copies of histones H2A, H2B, H3 and H4, is wrapped by ~147 basepairs (bp) of DNA. Variations in DNA sequence, wrapping strength asymmetry, and position-dependent histone-DNA interactions are collectively responsible for the uneven character and polarity of the nucleosomal barrier
to an elongating polymerase (Bondarenko et al., 2006; Hall et al., 2009; Ngo et al., 2015). The

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topography of the nucleosomal barrier can be described using two parameters: its height at eachposition (i.e., the magnitude of the energy required to access the DNA) and its width (i.e., extension
along the DNA). Although Pol II has been shown to be capable of transcribing through the nucleosome both in vitro (Bondarenko et al., 2006) and in vivo (Weber et al., 2014), the detailed, highresolution dynamics of Pol II crossing the nucleosomal barrier have not been observed yet. Because
the properties of the barrier likely determine the dynamics of a transcribing polymerase, obtaining
high-resolution topographic and transcriptional maps of the barrier lies at the heart of understanding
the regulation of gene expression.
The majority of eukaryotic genes have a well-defined +1 nucleosome (the first nucleosome
encountered by Pol II following initiation), which is enriched in H2A.Z and uH2B histones(Rhee et al., 2014; Teves et al., 2014), and represents the highest barrier to transcription(Teves et al., 2014). Whether the high prevalence of H2A.Z and uH2B modifies the intrinsic barrierat the +1 nucleosome, results in a different local spatial organization of chromatin, plays a role inregulating the binding and/or activity of extrinsic transcription factors, or a combination of all ofthese, remains unknown. Early optical tweezers studies have shown that in front of a WT nucleosome, Pol II slows down, pauses, backtracks, and cannot actively ‘peel’ the DNA wrapped aroundthe histones (Hodges et al., 2009). Instead, the polymerase functions as a fluctuating ratchet that
advances by rectifying transient, spontaneous wrapping/unwrapping transitions of the nucleosomal
DNA around the histone core (Hodges et al., 2009). A similar study using tailless histones andmutated DNA sequences suggests that these nucleosomal elements modulate the topography ofthe barrier by affecting the density and duration of Pol II pauses (Bintu et al., 2012). However,
because of their low resolution, these studies failed to accurately map the topography of the barrier
and its effects on the dynamics of transcription. A high-resolution transcriptional map around thenucleosome is necessary to ultimately understand how the interaction of trans-acting factors at specific and selective positions of the polymerase around the octamer regulate transcription across thebarrier.
Prior attempts to characterize the nucleosomal barrier to transcription have suffered from two
substantial limitations. First, previous assessment of nucleosome stability relied on pulling andunwrapping the nucleosome from both ends (Mihardja et al., 2006). These experiments, while providing a measure of the strength of DNA/histone interactions, may not fully recapitulate the physicalprocess of nucleosome invasion by Pol II, which unidirectionally unwinds the nucleosomal DNA. Second, although we can now obtain transcription trajectories with millisecond temporal and near bp
spatial resolution using optical tweezers (Righini et al., 2018), it is very difficult to determine theabsolute location of the polymerase on the template (Hodges et al., 2009; Bintu et al., 2012;
Fitz et al., 2016), what we term here ‘accuracy’. Here we surmount both limitations, obtaining highresolution, high-accuracy topographic and transcriptional maps of WT and modified nucleosomes.
By registering the dynamics of Pol II as a function of its position along the nucleosome, these maps
provide a means to interrogate how variant and epigenetically modified histones affect the dynamics
of transcription through the nucleosome.

Results
Single-molecule unzipping of nucleosomal DNA maps the topographyof the nucleosome barrier
To experimentally recapitulate the underlying physical process of barrier crossing, that is nucleosomal DNA unwinding, we mimicked the effect of Pol II passage through the nucleosome using
mechanical force. To this end, we adapted a previously described single-molecule DNA unzipping
assay (Hall et al., 2009; Rudnizky et al., 2016) in which the two strands of the nucleosomal DNA
are held in two optical traps, resulting in a Y-shaped configuration (Figure 1A). We engineered thestem ahead of the fork to consist of two consecutive segments of ‘601’ nucleosome positioning
sequence (NPS) (Lowary and Widom, 1998), and a short hairpin loop at the end to prevent tether
breaking once all double-stranded DNA (dsDNA) is converted into single-stranded DNA (ssDNA)(Figure 1A). During each experiment, we move the two traps apart at a constant speed of 20 nm/s.
When the force reaches ~17 pN, the dsDNA at the stem begins to unzip. When the stem segment
does not contain a nucleosome, the DNA unzips following a series of closely spaced transitions

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Figure 1. Dual-trap Optical Tweezers Single-molecule Unzipping Assay Unwinds Nucleosomal DNA and Maps Histone-DNA Interactions. (A) Geometryof the single-molecule unzipping assay. Dashed arrows denote directions of trap movement (20 nm/s) during unzipping (red arrow) or rezipping (blackarrow). 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
Figure 1 continued on next page

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Figure 1 continued
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 ofthe 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 grayhistograms 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 gray dashed lines, with the peak positions (inbp) 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 1—figure supplement 1 for representative unzipping traces and analysis.
DOI: https://doi.org/10.7554/eLife.48281.002
The following figure supplement is available for figure 1:
Figure supplement 1. Unzipping Traces of Single Human WT, H2A.Z, M3_M7, uH2B Nucleosomes and Tetrasomes.
DOI: https://doi.org/10.7554/eLife.48281.003

occurring in a narrow range of forces between 17–20 pN, dictated by the sequence of the template.
Once all dsDNA has been fully converted into ssDNA, the force increases sharply at the hairpin end(Figure 1B). The highly reproducible force-extension signatures from the two consecutive NPS
regions allow us to align traces from different unzipping experiments by placing the force and dwelltime of the opening junction at each base pair into register (Figure 1B and Figure 1—figure supplement 1). Upon force relaxation (rezipping), the pattern of closely spaced transitions is reproduced in
the inverse sense to that observed in the pulling direction (Figure 1B, Video 1).
Next, we repeated the experiment with the second NPS preassembled with a human WT nucleosome. The unzipping force-extension signature of the first NPS matches those obtained above, butthat of the nucleosome region deviates significantly due to histone-DNA contacts (Figure 1C and
Figure 1—figure supplement 1A, Video 2). Relaxation of the tether results in two identical, sequential rezipping signatures characteristic of the naked DNA in ~75% of the cases (Figure 1B and C),
indicating that full DNA unzipping led to complete histone removal. However, in 25% of the cases,
when we unzip the same molecule for the second time, the force reaches higher values than withbare DNA but lower than those observed the first time, likely reflecting residual histone-DNA interactions from nucleosomal relics. Since we have no knowledge of nucleosome integrity by the second
round of unzipping, we only analyzed the first round of unzipping data for each molecule. Because
we moved the trap at a constant speed, the dwell-time of the fork at each position reflects the local
histone-DNA interaction strength at that force. Indeed, in these constant pulling velocity experiments, the forces applied to histone-DNA contacts lying deeper in the structure depend on theVideo 1. Unzipping-rezipping of bare NPS DNA.
DOI: https://doi.org/10.7554/eLife.48281.004

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Video 2. Unzipping-rezipping of hWT nucleosome.
DOI: https://doi.org/10.7554/eLife.48281.007

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forces reached previously in undoing earlier contacts. This effect may lead to underestimation of themagnitude of later interactions. Accordingly, we also performed force-jump unzipping experimentson the same constructs in which we suddenly increased the force to 28 pN (this force was chosen tominimize the contribution of the dsDNA sequence to the dynamics of the fork) and held it constant
while monitoring the unzipping fork dwell-time at each position along the NPS (Figure 2A). In theseexperiments, the bare DNA construct unzips to the hairpin end instantaneously, while the fork dwellsin the WT nucleosome primarily at 25, 31, and 35 bp into the nucleosome (Figure 2B). The forceweighted residence time histograms (see Methods for details) of the unzipping fork along the entire
NPS obtained from these two types of experiments are similar and provide a quantitative descriptionof the barrier to nucleosomal DNA unzipping with single bp resolution that we term the nucleosome
topography map (Figures 1E and 2B).
The topography maps reveal that the unzipping fork encounters substantial resistance at around17, 26, 31, 41, 52, 61 and 69 bp into the nucleosome, which correspond to regions of proximal
dimer and tetramer interaction with the first half of nucleosomal DNA (Figure 1E). Interestingly, theresistance diminishes significantly after the dyad, suggesting that unzipping the first half of thenucleosome destabilizes the histone-DNA interactions of the second half. As previously observed,
major histone-DNA interactions first occur ~55 bp from the dyad and exhibit 5 or 10 bp periodicity
as the unzipping fork progresses (Hall et al., 2009; Rudnizky et al., 2016). This observation probably reflects the strong histone-DNA contacts along the DNA minor groove every 10 bp observed in
the crystal structure of the nucleosome (Luger et al., 1997). Compared to a previous study(Hall et al., 2009), we noticed a shorter residence time near the nucleosome dyad, which we attribute to differences in pulling geometry, buffer conditions, and/or histone source.
Unzipping of tetrasomes (H3/H4 tetramer assembled on NPS) revealed a substantially diminishedbarrier compared to the octamer, with unzipping fork dwelling events mostly restricted to locations
near the dyad, and much lower maximum force reached during the unzipping process (Figure 1E,
Figure 1—figure supplement 1B). These data indicate that the H2A/H2B dimer not only interacts
locally with the DNA but it also affects the strength of the H3/H4 tetramer-DNA interaction near the
dyad to orchestrate the overall nucleosome stability. As loss or exchange of H2A/H2B dimers has
been implicated in important biological processes such as DNA replication (Ramachandran andHenikoff, 2015), transcription (Ramachandran et al., 2017; Kireeva et al., 2002), repair(Ransom et al., 2010), and DNA supercoiling (Sheinin et al., 2013); these findings highlight thepotential role of non-local histone-DNA interactions in those processes.

H2A.Z and uH2B alter orthogonal parameters of the nucleosome
topography map
Human H2A.Z and uH2B nucleosomes show altered topography maps when compared to their WT
counterparts (Figures 1E and 2B). However, the relative magnitude and distribution of the peaks
are differently affected by these two modifications. Specifically, uH2B nucleosomes stabilize thedimer region (16 and 25 bp peaks) with minor effects on the tetramer region (Figures 1E and2B), suggesting that the attachment of ubiquitin to H2B enhances the barrier height locally. The
peaks after the dyad are less pronounced and correspond to regions where nucleosomal DNA interacts with the distal dimer. H2A.Z nucleosomes show enhanced peaks at 41 and 52 bp, while exhibiting much lower heights at 25 and 31 bp (Figures 1E and 2B), indicating a global redistribution of
the barrier’s strength along its width. Overall, in H2A.Z nucleosomes the dwell-time peaks are more
broadly distributed throughout the first half of the barrier than in their WT or uH2B counterparts,
while maintaining their 5 to 10 bp periodicity.
To determine whether this redistribution of the barrier strength reflects features of the individual
H2A.Z nucleosomes or the superposition of barriers derived from a heterogeneous molecular population resulting from an enhanced lateral mobility of these nucleosomes, as has been previously suggested (Rudnizky et al., 2016), we counted the number of transitions (rips) per unzipping trace inthe nucleosome region. Indeed, the H2A.Z-containing nucleosome has on average one more transition per trace than its WT counterpart (Figure 1—figure supplement 1F) but displays a similar standard deviation. This effect is also evident in constant force unzipping experiments on H2A.Z
nucleosomes, in which more dwell-time peaks are observed along the NPS (Figure 2B). To check
whether H2A.Z nucleosomes are more mobile compared to WT nucleosomes, we repeatedly
unzipped-rezipped single WT or H2A.Z nucleosomes up to the proximal dimer region (maximum

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Figure 2. 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 thesecond 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 constantforce 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 2—figure supplement 1 on assembly cooperativity of H2A.Z nucleosomes.
DOI: https://doi.org/10.7554/eLife.48281.005
The following figure supplement is available for figure 2:
Figure supplement 1. H2A.Z Nucleosomes Assemble More Cooperatively than WT nucleosomes.
DOI: https://doi.org/10.7554/eLife.48281.006

force reached to ~25–30 pN). If H2A.Z induces lateral mobility of the nucleosome, the position of initial force rise in the nucleosome region would shift between each partial unzipping-rezipping round,and should be quite evident in our finely registered traces. Surprisingly, in contrast to the report by
Rudnizky et al. (2016), we observed no lateral mobility with either WT or H2A.Z nucleosomes, as
indicated by the highly reversible and overlapping unzipping signatures in the proximal dimer region(Figure 1—figure supplement 1G and H). Together, these results indicate that the effect of the
H2A.Z histone variant in our experiments is not to increase the heterogeneity of the nucleosome
population but to significantly redistribute the strength of the barrier, effectively broadening it. Thisconclusion is also supported by the homogeneous migration of H2A.Z nucleosomes in native gels.
(Figure 1—figure supplement 1I).
The distal dimer interaction peaks for H2A.Z nucleosomes are visibly diminished relative to thoseof WT and uH2B nucleosomes (Figure 1E). Interestingly, we observed an increased cooperativity
during the assembly of H2A.Z nucleosomes. As the ratio of octamer to DNA is increased during
nucleosome reconstitution, we consistently observed significantly less hexasome formation with
H2A.Z than with H2A (Figure 2—figure supplement 1A). It is possible that the global decrease of
DNA interaction with the distal dimer observed with H2A.Z nucleosomes could also reflect a morecooperative disassembly during unzipping. To pinpoint what regions within H2A.Z are responsiblefor its assembly cooperativity, we generated a series of sequence swap mutants between H2A and
H2A.Z (Figure 2—figure supplement 1B) (Clarkson et al., 1999). Swapping the sequences of theChen et al. eLife 2019;8:e48281. DOI: https://doi.org/10.7554/eLife.48281

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M3 or the M7 region in H2A.Z with the corresponding sequences in H2A promotes the appearanceof hexasomes, indicating decreased cooperativity in assembly (Figure 2—figure supplement 1A).
Consistently, the topography map of M3_M7 nucleosomes (an M3 and M7 combined swap mutant)showed intermediate topographical features between H2A.Z and WT nucleosome, with the distal
dimer interaction peaks (at 109 and 122 bp) becoming more pronounced than those of H2A.Z nucleosomes (Figure 1E), consistent with the idea that cooperativity in disassembly correlates with thisdistal interaction. Structurally the M3 region corresponds to the ‘loop 1’ that mediates H2A.Z-H2A.Z
interactions within the octamer, and the M7 region corresponds to the ‘docking domain’ that mediates H2A.Z interactions with H3-H4 (Zlatanova and Thakar, 2008). These regions play importantroles in the stability of the histone octamer. Thus, unique physical properties of the H2A.Z octamer
likely account for the broadened barrier distribution we observed during unzipping.

Observation of multiple nucleosomal states at the proximal dimer
regionOne unique feature from the nucleosome unzipping traces is the presence of fast, reversible unzipping transitions within the proximal dimer region—spanning the first 40 bp of the NPS—that manifest as ‘hopping’ of force and extension in the unzipping experiments (Figure 3—figure
supplement 1A). Hopping in this region is nucleosome-specific, as it is not observed during unzipping of bare NPS DNA (Figure 3—figure supplement 1B). The reversible nature of this transition
contrasts with the irreversible transitions observed deeper in the nucleosome, and may indicate a
rapid, small scale unzipping that is not large enough to disrupt the structure of the octamer. With
nucleosome unzipping occurring at approximately 20–30 pN (Figure 1C–D), to observe hopping at
our trap separation rate of 20 nm/s (with trap spring constant of approximately 0.3 pN/nm) requires
the system to transition between states on timescales of approximately 2 s or less. To better capture
these hopping dynamics, we fixed the trap distance such that the unzipping fork remained localized
within the proximal dimer region and monitored the fluctuations of the force and extension (passivemode experiment). Within an empirically determined trap distance range, we obtained equilibrium
extension hopping traces (Figure 3A), which show transitions on timescales of ~1 s, aligning with thenecessary rate of transitions between states. At each fixed trap separation, we determined the number of unzipped bp to obtain a probability distribution for the length of unzipped DNA (Figure 3B).
For both bare and nucleosomal DNA, these distributions show consistent peaks, as expected on aheterogeneous energy landscape where the system populates discrete energy wells separated bytransition barriers. We note that certain trap separations allow the system to sample multiple wells,
giving rise to a multi-modal distribution (Figure 3B, bottom panel), analogous to the hopping
observed in the constant pulling rate unzipping curves. Relative to bare DNA, WT nucleosomes display an additional peak in the distribution of unzipped bps, at approximately 28 bp after the start ofthe second NPS where the most significant contacts between DNA and the H2A-H2B dimer occur(Figure 3C). This peak implies the existence of a barrier to further unzipping that arises from bindingof the DNA to histones, and its position is consistent with the dwell time peak observed in the unzipping traces (Figure 1E, peak at 26 bp).
Assuming that the observed distributions (Figure 3B and C) correspond to equilibrium Boltzmann
statistics, we extracted the energy associated with unzipping of each bp in the proximal dimer
region of the nucleosome (Figure 3D, details in Methods). The presence of a strong interactionenergy peak at 32 bp into the WT nucleosome and the corresponding energy well preceding thisnew barrier position account for the appearance of the new preferred state in the distribution ofunzipped bps (Figure 3C, peak at 28 bp). Furthermore, the low barrier to rezipping from this state(* in Figure 3D) implies that the dynamics between the two states in the proximal dimer region of
the nucleosome (at approximately 18 bp and 28 bp) should be quite rapid, in keeping with the hopping behavior observed in the unzipping curves (Figure 3—figure supplement 1A). Subtracting theunzipping energy of bare DNA from that of the nucleosome, we obtained the additional energy
associated with each nucleosome type, providing a measure of the interaction energy between the
DNA and the octamer throughout the first quarter of the NPS (Figure 3D, inset). WT, H2A.Z, anduH2B nucleosomes all show strong DNA binding to the proximal dimer region, with a large peak in
the interaction energy centered at approximately 35 bp.

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Figure 3. 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 indicatesincreasing trap separation (purple to red), corresponding to clusters in Figure 3—figure supplement 1F. Gray 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 3—figure
supplement 1F. Each curve is from a different trap separation, matching colors in A and Figure 3—figure supplement 1F. Distributions are shown forboth bare DNA (top) and WT nucleosome (bottom). Vertical black dotted line indicates the start of the second NPS. Vertical gray 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 3—figure supplement 1F for force-extension data. (C) Zoomed-in view of the black dashed box in (B). Peakpositions are labeled in bp. (D) DNA unzipping energy computed by assuming the unzipped bp distributions from data in Figure 3—figure
supplement 1F (including distributions in B) correspond to equilibrium Boltzmann statistics. Inset DE 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 3—figure supplement 1
on hopping traces and analysis of energy landscape from equilibrium data.
DOI: https://doi.org/10.7554/eLife.48281.008
The following figure supplement is available for figure 3:
Figure supplement 1. Hopping of the Unzipping Fork Near the Proximal Dimer Region of the Nucleosome.
DOI: https://doi.org/10.7554/eLife.48281.009

A ‘molecular ruler’ gauges the positions of an elongating pol II withnear Base-pair accuracy
Having established the topography of the nucleosomal barrier via mechanical force-induced DNA
unwinding, we next set out to determine how this topography manifests in the dynamics of Pol II
during transcription through the nucleosome. We used a high-resolution dual-trap optical tweezers
instrument together with an improved nucleosomal transcription assay (Figure 4). To accurately
gauge the positions of Pol II on the template, we placed a ‘molecular ruler’ in front of the nucleosome (Figure 4A). The ‘molecular ruler’ consists of eight tandem repeats of an artificially designed
64 bp DNA that has a single well-defined, sequence-encoded pause site when transcribed by Pol II

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Figure 4. A ‘Molecular Ruler’ Gauges the Positions of an Elongating Pol II with Near-Basepair Accuracy. (A) Experimental design of an improved singlemolecule nucleosomal transcription assay. A single biotinylated Pol II (purple molecular structure) is tethered between two optical traps. Pol IItranscription is measured as increases in distance between the two beads at 10 pN constant force. The inset box shows the composition of thetemplate, which is constructed by ligating Pol II stalled complex (cyan), the molecular ruler (green), NPS DNA (or nucleosome, yellow-gray), and a shortinter-strand crosslinked DNA (for stalling Pol II, red). The ‘molecular ruler’ consists of eight tandem repeats of a 64 bp DNA (green), each harboring asingle 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 gray dashed lines are the predicted pause sites, whereas the short green lines mark the actual pausesof 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), 6A5T for PolII-SHL( 1)). Pol II, histones, template DNA, non-template DNA are colored in gray, green, red and blue,
respectively. See also Figure 4—figure supplement 1 on detailed characterization of the ‘molecular ruler’.
DOI: https://doi.org/10.7554/eLife.48281.010
The following figure supplement is available for figure 4:
Figure supplement 1. Biochemical and Single-molecule Characterization of the ‘Molecular Ruler’.
DOI: https://doi.org/10.7554/eLife.48281.011

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in bulk (Figure 4—figure supplement 1A, dashed rectangle) and in single-molecule assays(Figure 4B and Figure 4—figure supplement 1B–D). The repeating pausing patterns of Pol II withinthe ‘molecular ruler’ generated a periodicity of 21.1 ± 0.3 nm (Figure 4—figure supplement 1B),
corresponding to the length of 64 bp DNA under experimental force and buffer conditions. This
periodicity serves to align all transcription traces (Gabizon et al., 2018; Herbert et al., 2006) and it
also enables the accurate conversion of nanometer distances to basepairs of transcribed DNA (Figure 4—figure supplement 1D).
We used a bubble initiation method to assemble a stalled biotinylated yeast Pol II elongation
complex (Hodges et al., 2009; Bintu et al., 2012) that was ligated downstream to a 2 kb spacer
DNA and upstream to the ‘molecular ruler’, followed by a single nucleosome (Figure 4A). The ‘601’
NPS was used to ensure both precise nucleosome positioning and accurate assignment of Pol II positions as it crosses the barrier. Pol II transcription was restarted by supplying a saturating concentration of NTPs (0.5 mM). A Pol II stall site consisting of a short inter-strand cross-linked DNA segment
was placed downstream of the NPS (Figure 4A). In these assays, we used force-feedback to maintaina constant 10 pN assisting force throughout the transcription trajectory so that the increase of the
distance between the beads serves as an accurate measure of how far Pol II has transcribed. We findthis tethering geometry to be superior to prior designs (Hodges et al., 2009; Bintu et al., 2012)
because it isolates Pol II from the beads surfaces, thus mitigating photo-damage (Landry et al.,2009). A representative real-time trajectory of Pol II transcribing through the ‘molecular ruler’ followed by bare NPS DNA is shown in Videos 3 and 4.

Real-time, High-resolution Dynamics of Single Pol II Enzymes
Transcribing Through Single Nucleosomes
We first obtained traces of Pol II transcribing through bare NPS DNA (Figure 5A and B, black trace).
Utilizing the pausing patterns obtained with the ‘molecular ruler’ (Figure 4B, inset), we adapted arecently described algorithm (Gabizon et al., 2018) to align the traces such that the positions of Pol
II along the entire NPS are under registry (Figure 4B and Figure 4—figure supplement 1D). The
results show that Pol II has a median crossing time within the NPS of 11 s (Figure 6—figure supplement 1A), a pause free velocity of 28.9 ± 3.0 nt-s 1 (Figure 6—figure supplement 1H), and displays
very few backtracking events (0.17 per trace) (Supplementary file 2).
Next, we replaced the bare NPS DNA with an assembled Xenopus WT (xWT) nucleosome on thesame template (Videos 5 and 6). As expected, Pol II exhibits a dramatic slow-down, with a mediancrossing time of 129 s (Figure 6—figure supplement 1B), a pause-free velocity of 11.4 ± 4.1 nt-s 1(Figure 6—figure supplement 1H), and frequent pausing and backtracking within the NPS region(Figure 4C). 59% of Pol II succeeded in crossing the barrier (Figure 6—figure supplement 1F),
which is signaled by its reaching the stall site at
the end of the template (Figure 4B). Using the‘molecular ruler’ to precisely locate Pol II on thetemplate, and after subtracting the enzyme’s
footprint (16 bp) (Shao and Zeitlinger, 2017),
we obtained a median dwell-time histogram ofthe leading edge of Pol II along the entire NPS
with ±3 bp accuracy (depicted as linear and polarplots in Figure 6A). This dwell-time histogram,which we refer to as a transcriptional map of thenucleosome, illustrates at high resolution andaccuracy the height and width of the nucleosome barrier to the elongating Pol II. It complements the topographic map described above,translating it into a ‘functional map’. Our measurements represent a nearly 20-fold resolution Video 3. Pol II transcription through bare NPS DNA.
and accuracy improvement on previous attempts The horizontal gray dashed lines indicate predicted
to obtain a transcriptional map of the nucleoso- pause sites in the molecular ruler, the three horizontal
mal barrier, since those experiments could only black dashed lines represent NPS entry, dyad, and NPS
resolve roughly three barrier regions of ~50 bp exit, respectively. This applies to all other videos.
each, corresponding to entry, central and exit DOI: https://doi.org/10.7554/eLife.48281.012

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zones (Hodges et al., 2009; Bintu et al., 2012; Fitz et al., 2016).
Several features of Pol II barrier crossing dynamics emerge from the nucleosome transcriptionalmap (Figure 6A). First, the effect of the barrier begins immediately after the leading edge of Pol II
touches the NPS (3 bp peak); the strength of the barrier is largest at 28 bp and 10–20 bp before thedyad, and is negligible after crossing this pseudo-symmetry axis. Second, we identified a region
between 28–64 bp into the NPS where Pol II enters long-lived pauses and backtracks frequently(Figure 4C). These pauses are consistent among different molecules and exhibit a ~ 10 bp periodicity (28, 38, 48, 57, and 64 bp into the NPS). Notably, this region coincides with the region of maximum resistance in the single-molecule unzipping assay (Figures 1E and 2B), implying that thetranscriptional barrier encountered by Pol II while crossing the nucleosome reflects, to a first approximation, the barrier mapped by the unzipping assay. Third, some molecules were permanently
arrested in this region (Figure 5A,B, gray trace), but those that managed to cross it typically succeeded in reaching the stall site shortly thereafter (Figure 5A). Thus, we speculate that this region(28–64 bp) may play an important regulatory role for barrier crossing by the enzyme. The observed
asymmetry of the transcriptional map between both sides of the dyad axis (Figure 6A) may reflect a
substantial weakening of the histone-DNA interactions in the presence of the bulky resident enzyme
halfway across the barrier. However, the transcriptional map asymmetry is similarly observed in the
topographic map (Figure 1E), even though the bulkiness of the enzyme does not play a role in thoseexperiments. It is also possible that the barrier asymmetry reflects changes in the nucleosome integrity by the invading polymerase (Kireeva et al., 2002) or the propagating unzipping fork.
Xenopus histones are traditionally the most-widely used in nucleosome studies because they are
well behaved in recombinant form. Since we employed recombinant human histones in the unzipping assays, it was of interest to compare the Pol II transcriptional maps of Xenopus nucleosomeswith those obtained with their human counterparts utilizing the same 601 NPS. As seen in
Figure 6A, the maps are quite similar except that human nucleosomes confer a significantly higher
barrier to transcription (see Figure 5A–B, Videos 7 and 8 for representative traces) than those of
Xenopus in the proximal dimer region (Figure 6A, the 28 bp peak in orange and red panels).

Dynamic interplay between pol II and the nucleosome during barriercrossing
Interestingly, we observed extensive two state transition dynamics while Pol II is paused at certain
sites (frequently at 28 and 63 bp) (Figure 4C, Video 6). While Pol II hopping at 28 bp coincides withhopping of the unzipping fork near this region and probably reflects sampling of alternative nucleosomal states ahead of the enzyme, hopping at a much deeper location into the nucleosome (63 bp)
may have a more complex origin. We hypothesize that these latter dynamics may be due to local Pol
II-histone interactions or re-wrapping of the nucleosome in front of a backtracked enzyme. Indeed,
these hopping dynamics occur exclusively after Pol II enters a deeply backtracked state (Figure 5—
figure supplement 1).
We also investigated whether Pol II remains functionally competent after the crossing. The pausefree velocity of Pol II after exiting the nucleosome resumed to 70% (28.6 ± 0.8 nt-s 1) of its value
before the crossing (41.1 ± 1.0 nt-s 1). This observation seems to indicate that while the enzyme
remains functionally competent, its dynamic state has been affected by the encounter (Figure 6—
figure supplement 1H). It remains unknown what changes in the enzyme are responsible for thisslowing down and if they are reversible. On the other hand, we probed the integrity of the nucleosome by pulling away the two beads after Pol II crossed the barrier. If the nucleosome survived thetraversal by Pol II, it would now lie between the two tethering points, that is the upstream DNA handle and the polymerase. Rarely (<5%) these pulling curves displayed the force-extension signaturenormally associated with the presence of a nucleosome, suggesting that in situ reassembly of thenucleosome following Pol II traversal was inefficient under our assay conditions. A similar low efficiency of nucleosome reassembly was observed from transcription assays in bulk in the absence offactors added in trans such as FACT (facilitates chromatin transcription) (Hsieh et al., 2013).

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H2A.Z enhances the width anduH2B the height of thetranscriptional barrier
Next, we investigated the effects of human H2A.
Z and uH2B on Pol II transcription dynamics. Our
unzipping assay revealed that H2A.Z and uH2B
have distinct effects on the nucleosome barriers(Figures 1E and 2B). Under the same buffer andforce conditions, Pol II alone was capable ofcrossing either H2A.Z (Figure 5C and D, Videos 9
and 10) or uH2B (Figure 5E and F, Videos 11
and 12) nucleosomes. While the crossing probabilities of Pol II through H2A.Z and WT nucleosomes are similar (58% and 59%, respectively)
Video 4. Pol II transcription through bare NPS DNA,
(Figure 6—figure supplement 1F), the distribuNPS zoom.
tions of pause sites within the NPS are markedly
DOI: https://doi.org/10.7554/eLife.48281.013
different (Figure 6A) in that H2A.Z is seen tocause a global redistribution of Pol II pause sites
along the entire NPS. Such scattered distribution(3, 28, 36, 59, 66, 87, 101, 115, 125 and 138 bp) differs significantly from that of WT nucleosome (3,28, 57, 64 bp) where most pauses occur before the dyad. This spreading of the barriers for H2A.Z
nucleosomes is a further indication that the force applied to the upstream DNA (which is the same inboth experiments) is not the dominant factor responsible for the asymmetry of the WT barrier acrossthe dyad, but that the actual histone-DNA interactions are. Furthermore, these differences areunlikely to stem from H2A.Z nucleosomes being mis-positioned on the starting template, because
the first pause site at ~3 bp, where the leading edge of Pol II begins to interact with the nucleosome,
is observed in every molecular trajectory obtained with H2A.Z nucleosomes. Instead, we attributethem to broadened distributions of DNA-histone interactions as seen in the topography map for
H2A.Z nucleosomes (Figures 1E and 2B, also see discussion on energetic profiles of H2A.Z nucleosome in next section). Clearly, H2A.Z strongly modulates the width of the nucleosomal barrier totranscription.
In contrast, Pol II transcription through human uH2B nucleosomes has a slightly higher passage
probability (76%) than through hWT nucleosomes (Figure 6—figure supplement 1F), although at
the expense of longer crossing times (Figure 6—figure supplement 1E). The pause site distribution
also resembles that of the hWT or the xWT nucleosomes (Figure 6A), however, the median dwelltime of Pol II at pause sites near the dyad is more than double that for the WT nucleosome, suggesting that uH2B enhanced the height of the nucleosomal barrier to Pol II (Figure 6A). The overallpause-free velocities of Pol II transcription through H2A.Z (11.4 ± 7.1 nt-s 1) and uH2B nucleosomes(12.9 ± 3.4 nt-s 1) are lower than that through hWT (18.2 ± 7.1 nt-s 1) nucleosomes (Figure 6—figure supplement 1H). Consequently, the median crossing times of Pol II through H2A.Z (262 s) anduH2B (304 s) nucleosomes are longer than that through hWT nucleosomes (230 s) (Figure 6—figure
supplement 1C–E). Note, however, that pause-free velocity contributes negligibly to crossing time,as the latter is dominated by long pauses. Including pausing, translocation and backtracking, Pol II
takes longer to cross uH2B or H2A.Z than hWT nucleosomes (Figure 6B).
Similar to when traversing hWT nucleosomes, Pol II backtracks frequently during transcription
through H2A.Z and uH2B nucleosomes (Figure 5—figure supplement 1A,C and E). The average
number of backtracks and backtrack depths are similar, but backtrack durations are longer when Pol
II transcribes through H2A.Z nucleosomes than through WT and uH2B counterparts(Supplementary file 2), which may again reflect the broader extent of histone-DNA interactions in
H2A.Z nucleosomes. Failure to recover from some backtracks seems to contribute to Pol II arrests,
the positions of which were more scattered for Pol II transcribing through uH2B and H2A.Z than
through WT nucleosomes (Figure 6—figure supplement 1G). Interestingly, backtracked Pol II frequently enters long-lived pauses, some of which are also accompanied by frequent two-state transition dynamics (hopping behavior) (Figure 5—figure supplement 1D and F). During some of thelong-lived pauses associated with crossing of uH2B nucleosomes, we also observed three-state

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Figure 5. 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 gray dotted lines are the pause sites within the ‘molecularruler’. 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
Figure 5 continued on next page

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Figure 5 continued
blue, green, red and cyan are examples of successful nucleosome crossing, while the trace in gray 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 forclarity. 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 highresolution 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 5—figure supplement 1 on
backtracking and hopping dynamics.
DOI: https://doi.org/10.7554/eLife.48281.014
The following figure supplement is available for figure 5:
Figure supplement 1. Long-lived Pauses of Pol II in the Nucleosome are Associated with Backtracking and Hopping Dynamics.
DOI: https://doi.org/10.7554/eLife.48281.015

hopping behavior (Figure 5—figure supplement 1F) and large hopping transition events(Figure 5F, green trace) of Pol II. As this behavior is rarely observed in WT and H2A.Z traces (Figure 5), we speculate that these dynamics of Pol II reflect the presence of the bulky ubiquitin attachment. Collectively, these data reveal that H2A.Z mainly enhances the width and uH2B mainly
enhances the height of nucleosomal barrier to transcription. Consistent with the previously reported
higher stability of uH2B nucleosomes (Batta et al., 2011; Chandrasekharan et al., 2009), they posean overall higher barrier magnitude––especially in the region near the dyad––than their WT counterparts to the passage of polymerase. It is also worth noting that for WT and H2A.Z, but not for uH2B
nucleosomes, the transcriptional map replicates the corresponding topography map, suggestingthat there could be uH2B-Pol II specific interactions that are not present in the unzipping assay.
Our findings provide direct evidence that H2A.Z or uH2B by themselves affect the crossing
dynamics of Pol II; and despite their differential effects on nucleosome topography, they both represent stronger barriers than WT nucleosomes for Pol II.

A mechanical model for pol II transcription through the nucleosome
We use a simplified mechanical model to calculate the expected polymerase dwell times along
nucleosomal DNA given a profile of DNA-octamer interaction energies. We build on a previously
published Pol II model that includes a mechanical DNA linkage between Pol II and the nucleosome,in addition to nucleosome-Pol II steric interactions (Figure 7A) (Koslover and Spakowitz, 2012).
The model assumes that Pol II is unable to actively separate the DNA from the surface of theoctamer. Instead, the enzyme behaves as a ratchet that makes progress by rectifying the unwrapping fluctuations of the nucleosomal DNA. The enzyme can also backtrack and diffuse forward to reengage the 3’-end of the nascent transcript with its active site. The extended model incorporates
varying binding energies for the DNA along thenucleosome.
Polymerase progress along the nucleosomal
DNA is modeled as a series of transcription
steps and backtracking excursions (Figure 7B),
adapting the model of Dangkulwanich et al.
(2013) to include interactions with the nucleosome. The individual polymerase steps are
assumed to occur on an energy landscape thatencompasses both the elastic energy of deforming the unwrapped DNA linker between Pol II
and the nucleosome core particle, and the interaction energy between the wrapped DNA andthe core particle (Figure 7C). For a given Pol II
position, the amount of unwrapped DNA isassumed to fluctuate rapidly around an energy
minimum that balances these two contributions. Video 5. Pol II transcription through xWT nucleosome.
As Pol II steps forward, the linker shortens and DOI: https://doi.org/10.7554/eLife.48281.018

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the elastic energy increases, with a longer linker
entailing a smaller energy increase and thus amore rapid rate constant for polymerase forward
motion. The interaction energy profile between
the DNA and the histone octamer ahead of Pol II
determines the ensemble of linker lengths and,thus, indirectly controls the average time spent
by Pol II at each DNA base pair. This modelallows us to calculate the expected dwell time ofthe polymerase at each position.
Accordingly, we use the DNA-octamer interaction energies extracted from equilibrium
unzipping data (Figure 3D) to calculate theexpected mean dwell times of polymerase in theVideo 6. Pol II transcription through xWT nucleosome,proximal nucleosomal region. The dwell time
NPS zoom.
peaks resulting from these equilibrium interacDOI: https://doi.org/10.7554/eLife.48281.019
tion energies approximately correspond topeaks in the experimental dwell time profiles(Figure 7—figure supplement 1). The DNAoctamer interaction energies cannot account for the first dwell time peak (the peak at ~3 bp in
Figure 6A) corresponding to the initial encounter between Pol II and the nucleosome. We hypothesize that this peak is the result of additional interactions between Pol II and the histone proteinsrather than arising from the difficulty of peeling DNA from the nucleosome.
Because equilibrium interaction energy data was available only for the initial section of the nucleosome (Figure 3), Pol II dwell times further into the nucleosomal sequence cannot be predictedfrom the data available. Instead, we solve the inverse problem: given the measured Pol II mean dwell
times (Figure 7D), we fit the DNA-nucleosome interaction energies (Figure 7E, details in Methods)
required to generate this dwell time profile. The dependence of dwell times on the nucleosome
binding energy in this model is non-local—time spent at a particular bp depends on the energy
required to unwrap a segment of DNA ahead of the polymerase. Consequently, peaks in the dwell
time profile arise from interactions that involve both large energy values and span a substantial
length of DNA (extended regions of strong binding). For example, the peaks at 29 bp in the dwell
time profiles for hWT, uH2B, and H2A.Z nucleosomes (Figure 7D), correspond to large peaks in theinteraction energy at 32 bp, consistent with the interaction energy profiles obtained from equilibrium
DNA unzipping data (Figure 3D). A two-peaked region of strong binding at roughly 43 bp in the
H2A.Z and uH2B nucleosomes gives rise to corresponding double peaks in the dwell time profiles
41 bp into the nucleosomes, with substantially longer pausing times for H2A.Z nucleosomes in thisregion. An additional broad region of strong binding is seen just before the dyad axis, 62 bp in hWT
and uH2B nucleosomes, resulting in the observed Pol II pausing peaks 59 bp into the nucleosome.
Interestingly, according to our mechanical model, the predicted interaction energies necessary to
generate the observed dwell-time profiles are similar in magnitude for all three nucleosomal types.
However, wider peaks of strong binding give rise to increased pause durations in the uH2B transcriptional profile, while a distribution of many narrower peaks accounts for the increased number of
pausing sites in the H2A.Z profile (Figure 7D,E).

Discussion
For the last 20 years the crystal structure of the nucleosome (Luger et al., 1997) has guided ourview of the packaging unit of the genome and suggested its role as a regulator of gene expression.
As a mechanical and energetic barrier, the nucleosome gates the accessibility of genomic DNA, constituting a fundamental regulatory mechanism for all DNA-templated processes including replication,transcription, repair, recombination, and chromatin remodeling. Epigenetic modifications and histone variants are known to modulate all of these processes. The question of whether this modulation
results from the recruitment of trans-acting factors, or responds to changes in the intrinsic propertiesof the barrier, or both, has not previously been addressed.

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Figure 6. 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 widthis 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
Figure 6 continued on next page

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Figure 6 continued
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 theentire 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 6—figure supplement 1 on statistics of the crossing time, crossing probability, pause-free velocity
and arrest position.
DOI: https://doi.org/10.7554/eLife.48281.016
The following figure supplement is available for figure 6:
Figure supplement 1. Crossing Time, Crossing Probability and Pause-free Velocity of Pol II during Transcription through NPS DNA or Nucleosomes.
DOI: https://doi.org/10.7554/eLife.48281.017

Low-resolution single molecule assays showed that it is possible to follow molecules of Pol II as
they cross the nucleosomal barrier (Hodges et al., 2009; Bintu et al., 2012). However, these studies
only yielded gross features of the barrier and failed to provide the crucial spatial-dependent dynamics of the crossing that are required to rationalize the effect of nucleosome modifications at themolecular level.
Very recently, cryo-EM structures of Pol II-nucleosome complexes have provided snapshots of Pol
II paused at major histone-DNA contacts and suggested sites of interaction with other factors(Farnung et al., 2018; Kujirai et al., 2018; Ehara et al., 2019). Missing from these structures isinformation about the dynamics of barrier crossing by the enzyme: what are the time windows available for in-trans interactions with these discrete sites, how are these related to the local energetic
magnitude of the barrier, and how are they modulated by epigenetic modifications and histone variants. Using a ‘molecular ruler’, we have been able to locate individual Pol II molecules along thetemplate with high precision and to extract their molecular trajectories as they transcribe throughnucleosomes at near bp resolution and accuracy. These trajectories unveil unprecedented details on
the general dynamics (translocating, pausing, hopping and backtracking) as well as the residence
times of the enzyme at every position as it progresses through the nucleosome, providing insights
into how gene expression is regulated spatially and temporally at a single nucleosome level.
Our results reveal that the proximal dimer region of the nucleosome (~28 bp) in the transcription
direction is a major physical barrier for Pol II and may serve as an important regulatory checkpoint
for gene expression. In this region, Pol II frequently enters long-lived pauses; this result is consistent
with the observation of a major Pol II pause site at the superhelical location SHL( 5) reported
recently (Kujirai et al., 2018). Interestingly, pausing at this location is accompanied by extensive
hopping dynamics, likely reflecting unwrapping/rewrapping of the nucleosomal DNA around theoctamer and/or structural rearrangements of the nucleosome. Indeed, partially unwrapped nucleosomal intermediates have been detected in vitro by time-resolved small angle X-ray scattering(Chen et al., 2014), by cryo-EM (Bilokapic et al.,
2018)
and
in
vivo
by
MNase-seq(Ramachandran et al., 2017). The location ofthese structural intermediates coincides with thedifferent nucleosomal hopping states observed
as the unzipping fork reaches the proximal
dimer, reinforcing the interpretation that local
DNA/histone interactions determine the dynamics of Pol II in this region and its ultimate progress beyond it.
Traditionally, the dyad has been viewed as
the strongest histone-DNA contact point andtherefore as the highest barrier position in WT
nucleosomes. Unambiguously assigning Pol II’s
residence time with bp resolution has allowed us
to define a transcriptional map of the barrier,which indicates that the proximal dimer region Video 7. Pol II transcription through hWT nucleosome.
and not the dyad represents the highest barrier DOI: https://doi.org/10.7554/eLife.48281.020

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Video 8. Pol II transcription through hWT nucleosome,
NPS zoom.
DOI: https://doi.org/10.7554/eLife.48281.021

Video 9. Pol II transcription through H2A.Z nucleosome.
DOI: https://doi.org/10.7554/eLife.48281.022

to an elongating Pol II. This observation is consistent with the unzipping experiments that also
reveal the proximal dimer region as mechanically the most stable. We posit here that the change indynamics of the polymerase, progressing slowly in this region, provides a crucial time window toallow for other facilitative or inhibitory factors to bind and further modulate the strength of the barrier to the transcribing enzyme. For instance, FACT may bind to the nucleosome and remove one
histone dimer ahead of the Pol II and reassemble the nucleosome after Pol II traversal(Belotserkovskaya et al., 2003; Mason and Struhl, 2003; Chen et al., 2018). These early regulatory
steps as Pol II invades the nucleosome not only gate gene expression but also permit the regulationof chromatin integrity and of epigenetic modifications. As Pol II progresses further into the nucleosome, the strength of the barrier appears to be dynamically modified either through nucleosome
destabilization, the steric bulkiness of the enzyme, or both. Beyond the dyad, there is practically nobarrier in WT nucleosomes, again in agreement with a recent cryo-EM structural report(Kujirai et al., 2018).
Consistent with these observations, modifications that play important regulatory roles such as
H2A.Z and uH2B, mainly affect the proximal dimer region, although their effects are not circumscribed to this location. Pol II transcription through nucleosomes bearing H2A.Z or uH2B reveal thatthese modifications strongly increase the strength of the barrier, but do so distinctively: H2A.Z
increases the width of the barrier whereas uH2B increases its height. Significantly, the topographic
map of the WT and H2A.Z barriers before the dyad, as determined here by force-induced nucleosomal DNA unwinding, closely parallels the transcriptional map of Pol II, indicating that to a first
approximation, the ability of the enzyme to crossthe barriers in this region is dictated by the energetic requirements of disrupting DNA-histoneinteractions.
In vitro, H2A.Z has been observed to either
enhance or decrease nucleosome stability
depending on the assays used (Zlatanova andThakar, 2008; Bönisch and Hake, 2012). Our
improved optical tweezers experiments offer
unprecedented resolution that captures a more
complex picture in which H2A.Z redistributes thestrength of the barrier across and beyond thedyad, effectively increasing its width. Accordingly, the physical barriers across the H2A.Z
nucleosome are lower, yet more globally distrib- Video 10. Pol II transcription through H2A.Z
uted. The precise origin of this broader distribu- nucleosome, NPS zoom.
tion is not known. A previous study suggested DOI: https://doi.org/10.7554/eLife.48281.023

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Video 11. Pol II transcription through uH2B nucleosome. Video 12. Pol II transcription through uH2B
nucleosome, NPS zoom.
DOI: https://doi.org/10.7554/eLife.48281.024
DOI: https://doi.org/10.7554/eLife.48281.025

that H2A.Z nucleosomes are more mobile compared to their WT counterparts, although theextent and cause of the mobility remain unclear (Rudnizky et al., 2016); however, we do not
observe this enhanced mobility in our experiments. Like a previous study (Park et al., 2004), ourdata support the idea that the H2A.Z octamer is more stable than its WT counterpart within the
nucleosome. As a result, H2A.Z hexasomes are barely observed during nucleosome assembly andwe find that the M3 and M7 regions within H2A.Z are important for conferring such increased
octamer stability. We speculate that increased octamer stability strengthens the overall nucleosomal
barrier as reflected in the increased crossing time of Pol II through such nucleosomes.
The effect of H2A.Z on the nucleosomal barrier can be seen as that of re-distributing the strengthof the barrier from height to width. Interestingly, the Arrhenius dependence of barrier crossing time
predicts that the time to cross a barrier of height n x h is proportional to the nth-power of the timeto cross a barrier of height h. In contrast the time to cross n sequential barriers of height h is proportional to n times the time to cross each one of the barriers. Thus, based on these considerations
alone, we would expect that H2.A.Z would decrease the crossing time of Pol II, not increase it, as
observed. Therefore, factors other than barrier crossing time, but favored by the presence of thebarrier (e.g. backtracking, pausing, etc), are the ones that dominate the overall crossing time in thecase of H2A.Z nucleosomes. By dividing the height of the barrier into several smaller ones, H2A.Z
nucleosomes provides the enzyme more opportunities at different locations to pause, backtrack andpossibly interact with regulatory factors acting in trans such as chaperones and chromatin remodelers. In vivo, the effects of H2A.Z on transcription are complex and somewhat species-dependent.
The strong barrier posed by H2A.Z nucleosomes may explain its role in poising quiescent genes for
activation in yeast (Santisteban et al., 2011) and its prevalence in +1 nucleosomes across eukaryotic
genomes (Weber et al., 2014). In contrast, the observation that H2A.Z facilitates transcription in
multi-cellular organisms (Weber et al., 2014) is more likely due to recruitment of trans-acting
factors.
Using homogeneous, chemically-defined recombinant nucleosomes, we also demonstrated thatuH2B strengthens histone-DNA interactions at the dimer region and increases the overall barrier
strength to Pol II. Interestingly, while uH2B occurs at the dimer region, its effect on Pol II transcription propagates to other regions of the nucleosome including the region preceding the dyad. Thus,
the effects of epigenetic modifications are not merely local but may extend further into the barrier.
Uncovering such position-dependent nucleosome properties and dynamics has been possible by thehigh resolution and accuracy achieved in our single-molecule assays.
In vivo, H2B ubiquitination is highly dynamic and both the addition and removal of ubiquitin are
required for optimal transcription (Wyce et al., 2007). Like H2A.Z, it is not known whether these
phenotypes are due to altered nucleosome stability or to impaired or facilitated recruitment oftrans-acting factors. Nevertheless, higher levels of H2A.Z and uH2B are observed in transcriptionallysilent gene promoters in yeast (Batta et al., 2011; Zhang et al., 2005). Preventing H2B ubiquitination in yeast led to increased Pol II occupancy and transcription from quiescent promoters

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Figure 7. 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) Modelof 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 fromthe most recent main pathway state. Pol II steps forward one base pair with rate k0 or can enter a backtracked pathway by stepping backward one basepair at rate kb1. From backtracked positions, Pol II can move forward a base pair with rate kfn or can backtrack another base pair at rate kbn. Moving
forward from the first backtracked state returns Pol II to the main pathway. (C) Energy landscape of nucleosome-Pol II interaction, for constant DNAnucleosome interaction energies of 1kBT 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 areunfavorable 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 inFigure 6. (D) Dwell time profiles for human WT, H2A.Z, and uH2B nucleosomes. Solid black lines are experimental mean dwell times and colored
Figure 7 continued on next page

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Figure 7 continued
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 7—figure supplement 1 for fitting of nucleosome energy profiles basedon Pol II dwell times.
DOI: https://doi.org/10.7554/eLife.48281.026
The following figure supplement is available for figure 7:
Figure supplement 1. Fitting Nucleosome Energy Profiles Based on Pol II Dwell Times.
DOI: https://doi.org/10.7554/eLife.48281.027(Batta et al., 2011). These observations are consistent with our data that H2A.Z and uH2B provide
orthogonal and selective means to enhance the transcription barrier thus contributing to the maintenance of the transcriptional ground state and to gene silencing. Our results show that while thesemodifications may also act indirectly through their actions on the binding of trans-acting factors,
they exert a direct and significant effect on transcription dynamics by Pol II.
We have developed a unified mechanical model that uses the experimentally determined spaceresolved residence times of the enzyme at each position on the nucleosome to determine the energetics of the barrier. This model is quite general and should prove useful in predicting the behavior
of Pol II through alternative barriers, and in understanding the mechanics of barrier crossing forother molecular motors.
The dynamics of Pol II transcription through the nucleosome in vivo are affected by numerous
other factors such as higher-order chromatin folding, DNA topology, and transcription regulators
including histone chaperones, elongation factors, and chromatin remodelers. Integrating one ormore of these elements in single-molecule assays such as the one presented here provides an interesting avenue for future work to fully elucidate the features and principles underlying this biologically crucial and biophysically complex molecular encounter. Because epigenetic modifications arepotent regulators of eukaryotic gene expression, these results shed new light on the mechanistic link
between modifications enriched on the +1 nucleosome and the barrier to transcription. More
broadly, the real-time characterization of the dynamics of Pol II molecules traversing through nucleosomes at the highest resolution and accuracy reported so far, and the resulting nucleosome transcriptional map, constitute important steps towards uncovering the physical mechanisms
underpinning the regulation of eukaryotic gene expression.

Materials and methods
Key resources table
Reagent type(species) or
resource

Additional
information

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Antibody

NeutrAvidin(deglycosylated
native avidin
from egg whites)

Thermo Fisher

Cat# PI31000

powder dissolved
in PBS, 0.5 mM

Antibody

Anti-Digoxigeninfrom sheep

Sigma-AldrichCat#
11333089001,
RRID:AB_514496

powder dissolved
in PBS, 0.2 mg/ml

StrainDH5a competent cells

Fisher Scientific

Cat# 18-265-017

StrainBL21(DE3)
competent cells

NEB

Cat# c2527H

StrainAgilent SURE 2
Supercompetent Cells

Fisher Scientific

Cat# 200152

ChemicalcompounddNTP set
100 mM Solutions

Fisher Scientific

Cat# R0181

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ChemicalcompoundNTP set100 mM Solutions

Fisher Scientific

Cat# R0481

Chemicalcompound3’-Deoxyadenosine5’-Triphosphate

TriLink Biotechnologies

Cat# N3001

Chemicalcompound3’-Deoxyguanosine5’-Triphosphate

TriLink Biotechnologies

Cat# N3002

Chemicalcompound3’-Deoxycytidine5’-Triphosphate

TriLink Biotechnologies

Cat# N3003

Chemicalcompound3’-Deoxyuridine5’-Triphosphate

TriLink Biotechnologies

Cat# N3005

ChemicalcompoundTrioxsalen

Sigma-Aldirch

Cat# T6137

Chemicalcompound[a 32P]-ATP

Perkin Elmer

Cat# BLU003H250UC

Commercial
assay or kit

T4 DNA ligase

NEB

Cat# 0202L

Commercial
assay or kit

E. coli DNA ligase

NEB

Cat# 0205L

Commercial
assay or kit

Phusion high-fidelity
DNA polymerase

NEB

Cat# M0530S

Commercial
assay or kit

DraIII-HF

NEB

Cat# R3510S

Commercial
assay or kit

BsaI-HF

NEB

Cat# R3535L

Commercial
assay or kit

BglI

NEB

Cat# R0143S

Commercial
assay or kit

EagI-HF

NEB

Cat# R3505S

Commercial
assay or kit

SapI

NEB

Cat# R0569S

Sequencebased reagent

Lambda DNA

NEB

Cat# N3011S

Recombinant
DNA reagent

Primers formaking constructs
and DNA templatesThis paper

IDT custom order

Recombinant
DNA reagent

pGEM-3z/601

AddgeneCat# 26656

Recombinant
DNA reagent

pGM3z-8repeat-2BsaI

This paper

N/A

Software,algorithm

LabVIEW VIs

Comstock et al. (2011)

RRID:SCR_014325

Software,algorithm

Matlab scripts fordata analysis

This paper

RRID:SCR_001622

Software,algorithm

Python scripts fordata analysis

This paper

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ImageJ

NIH (open source)

RRID:SCR_003070

Other

Pierce Streptavidin
Magnetic Beads

Thermo Fisher

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BD 1 mL Insulin
Syringe with Slip Tip

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information

Supplementary file 1

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Dual-trap time-sharedhigh resolution optical tweezers(Comstock et al., 2011)

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1 mm carboxylated
polystyrene beads

Bang Laboratories, Inc.

Cat# SVP-10–5

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1.26 mm streptavidin
polystyrene beads

Spherotech

Cat# PC04001-PC04N

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Ni-NTA Agarose

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Millipore Sigma

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Amicon Ultra-0.5
Centrifugal Filter Unit, 3K

Millipore Sigma

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Zyppy Plasmid
Maxiprep Kit

Zymo Research

Cat# D4028

Other

Typhoon imager

GE Healthcare

TRIO Variable Mode

Additional
information

General materials
All DNA modifying enzymes were purchased from New England Biolabs (NEB). Oligonucleotides
were purchased from Integrated DNA Technology (IDT). Nucleotide triphosphates were purchasedfrom Thermo Fisher Scientific, and standard salts and buffer components were purchased from
Sigma Aldrich. Cloning and DNA template construction follows standard molecular biology techniques unless otherwise noted. The sequences of all oligos used are listed in Supplementary file 1.

DNA constructs for single-molecule unzipping experiments
DNA arms of the Y structure are prepared by standard PCR reactions using lambda DNA as the template. The left (with BsaI site) and right arm (with biotin) of the Y was amplified using oligos ZC01ZC02 and ZC03-ZC04, respectively. The left arm was digested with BsaI, and annealed with the rightarm to form the Y. The length of the left (after ligation) and right arm dsDNA are 937 and 911 bp,
respectively.
The first (for alignment) and second (for loading) NPS were amplified with ZC05-ZC06 and ZC07ZC08 respectively from the pGM3z-601 plasmid. The first NPS fragment was digested with BsaI, andthe second NPS fragment was digested with BsaI/DraIII. The first NPS was ligated with the Y to form
Y-alignment. The second NPS was ligated with the end hairpin to form NPS-hp. The end hairpin was
pre-folded by heating oligo ZC09˚C to 98˚C for 2 min, followed by a slow decrease of temperature
to 25˚C at 1 ˚C/min. Y-alignment were purified using agarose gel. NPS-hp was purified using native
PAGE followed by electro-elution and anion exchange chromatography with HiTrap-Q column.

Beads preparation
To couple oligonucleotides to polystyrene beads, ZC10 was hybridized to ZC11 to generate a double stranded oligo containing a phosphorylated 5’-CGGT overhang. Annealing was performed byheating a 1:1 mixture of the oligos in water (0.25 mM each) to 95˚C for 10 min, followed by coolingto room temperature. This resulted in the following oligo duplex:
5’ NH2-TTAATTCATTGCGTTCTGTACACG 3’
3’ TTAAGTAACGCAAGACATGTGCTGGC-phos 5’

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1 mm diameter carboxylated polystyrene beads were coupled to the prepared double-stranded
duplex as follows: 10 mL of 10% (W/V) beads were washed four times with 200 mL coupling buffer(0.1 M MES, pH = 4.7, 150 mM NaCl, 5% DMSO), and dispersed in 20 mL coupling buffer. All centrifugations took place for 5 min at 4500 g. 10 mL of 20 mM double stranded oligo and 6 mL of 2 M 1ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) were added, followed by vigorous shaking for 2
hr at room temperature. At this point another 10 mL of 2M EDC were added, followed by overnightshaking at room temperature. The remaining active EDC was then quenched by adding 2.5 mL of 1
M glycine, and the beads were washed 5 times with storage buffer (Tris 20 mM pH = 8, 1 mM
EDTA, 0.05% Tween 20, 5 mM NaN3) with 3 min of sonication between washes. The beads (1 mm
oligo beads) were finally dispersed at a concentration of 1% (w/v) and stored at 4˚C.
The beads were passivated by diluting 6-fold in TE (20 mM Tris, pH 8.0, 1 mM EDTA) and addition of b-casein to a final concentration of 1 mg/ml. The beads were vortexed for 10 min, washedonce with TE, dispersed to a concentration of 0.2% (w/v) in TE and stored at 4˚C until theexperiment.

Histone octamer assembly and purification
Recombinant human histones H2A, H2B, H3.3 and H4 were purchased from the Protein Expression
and Purification (PEP) Facility at Colorado State University. H2A.Z, H2A/H2A.Z swap mutants and all
Xenopus laevis histones were expressed in E. coli, purified and reconstituted into octamers according to standard protocols (Dyer, 2003). uH2B was prepared by crosslinking ubiquitin (G76C) andH2B (K120C) as previously described (Long et al., 2014).

Nucleosome reconstitution on NPS-hp
Purified human histone octamers or tetramers were stored in 10 mM Tris, pH 7.6, 1.6 M NaCl, 1 mM
EDTA, 1 mM DTT, 20% glycerol at 80˚C. The reconstitution of nucleosome was performed using asalt dilution method as described (Dyer, 2003). Briefly, NPS-hp DNA and histone octamers or tetramers were mixed in different molar ratios ranging from 1:0.8 to 1:1.4 and initial salt concentrationof the mixture (10 mL) was brought to 2 M NaCl. These reactions were incubated at 30˚C in a PCR
machine and the following amounts of dilution buffer (10 mM Tris, pH 7.6, 1 mM EDTA, 1 mM DTT,
0.1 mg/mL BSA) were added every 15 min: 3.3, 6.7 5, 3.6, 4,7, 6.7, 10, 30, 20, 100 mL. The reaction
products were analyzed by native polyacrylamide gel electrophoresis (4%, acrylamide:bisacrylamide
ratio 37.5:1) with 0.5  TBE plus 5 mM MgCl2 on ice. The reaction that gave no aggregates and minimal amounts of free DNA was chosen for further concentration using Amicon Ultra centrifugation filters with Ultracel 100K membrane. Concentrated nucleosomes were supplemented with 0.02 %NP40 and stored at 4˚C.

Optical tweezers assay for single-molecule unzipping
Unzipping oligo beads were prepared by ligating 5’-CGGT 1 mm polystyrene oligo beads with
Y-alignment DNA, NPS-hp nucleosome (or NPS-hp DNA) using E. coli DNA ligase. The reaction was
carried out at 16˚C for 2–3 hr. The ligated beads were diluted with TB50 buffer (20 mM Tris, pH 8.0,
50 mM KCl, 5 mM MgCl2, 1 mM DTT, 10 mM NaN3, 0.1 mg/mL BSA) to a final bead density of
0.00006% (w/v) and loaded into the tweezing chamber, which was filled with TB50 buffer. Tweezing
chamber was pretreated with 5% Pluronic F-127 and 1 mg/mL BSA followed by washing with TB50
buffer prior to each experiment. The 1.26 mm SA beads were diluted directed with TB50 buffer tothe same bead density as that of oligo beads.
Optical tweezers experiments were performed in a custom-made dual-trap optical tweezers
instrument modified from the design in Comstock et al (Whitley et al., 2017; Comstock et al.,2011). In this configuration, a 1064 nm laser is passed through an acousto-optic deflector, with thelaser alternating in position between the two traps every 5 ms. The position of the beads relative tothe traps was measured using back focal plane interferometry (Bustamante et al., 2009). Single
tethers were formed in situ inside the chamber by trapping an oligo bead in one trap and an SA
bead in the second trap, and bringing in close proximity the two traps to allow the biotin on theright arm of the Y to interact with streptavidin (Streptavidin bead).
The pulling experiments used to obtain the average dwell times in Figures 1E and 2B are nonequilibrium in nature, reflecting an average over many out-of-equilibrium encounters with eachChen et al. eLife 2019;8:e48281. DOI: https://doi.org/10.7554/eLife.48281

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nucleosome type, and are not direct indicators of equilibrium interaction strength alone. In particular, dwell times to rupture in a given trace will depend on the forces reached during a prior rupture
event. Complementary probes of DNA-histone interaction strength (see below) verify that many ofthe peaks in Figures 1E and 2B are due to strong binding interactions.
Due to the nonequilibrium nature of the experiments used to obtain the residence times in
Figures 1E and 2B, we caution against the use of such ‘pulling’ residence times alone to assessinteraction strength with high resolution. Data from these pulling experiments is most useful whenassessed in conjunction with data from other experimental techniques, such as the equilibrium hopping data in Figure 3, and the transcriptional residence times of Figures 6 and 7.

Unzipping at a constant trap separation speed
Once a single tether was confirmed, the trap distance was reset to a value at which the tether force
was ~0.4 pN. Unzipping was initiated by moving the two traps apart at a constant speed of 20 nm/s
for a total of 875 nm. Under this condition, this pulling speed corresponds to a loading rate of 1.8–
2.5 pN/s. Rezipping was conducted at the end of unzipping by bringing the two traps together tothe initial trap position at the same speed. The tether was broken manually by increasing the trap
distance and calibration was performed as previously described (Berg-Sørensen and Flyvbjerg,
2004). Data was acquired at 800 Hz.

Unzipping at 28 pN constant force
To unzip the construct at constant force, the tether was initially held at ~10 pN and force feedback
was turned on to maintain the tether at a constant force of 28 pN. The position and distance
between the two beads ware recorded at 800 Hz until the construct was fully unzipped. The force
feedback was turned off and the tether was relaxed to ~0.5 pN. For the purpose of aligning thetraces and accurately converting nanometer distance to basepairs unzipped, an unzipped trace using
constant trap separation speed (as described above) was further obtained from the same tether.

Partial unzipping up to the proximal dimer region to test nucleosome
mobility
Unzipping was performed at constant trap separation speed of 50 nm/s up to the proximal dimer
region where the force starts to rise above the baseline of bare DNA construct, but does not reach
30 pN. The partial unzipping was followed by rezipping to the initial trap position. Typically, thisresults in a trap movement of ~620 nm. After repeating the unzipping-rezipping cycle for 5–10 times,
a final unzipping that unzips all the way to the hairpin end (trap movement of 875 nm) was performed to disrupt the whole nucleosome. A bare DNA trace was also collected immediately following this final unzipping.

Unzipping at constant trap distances to record hopping traces near theproximal dimer interaction region
To capture hopping of the unzipping fork near the proximal dimer interaction region of the nucleosome, the trap was manually moved apart at a small distance increment of 7.1 nm. At each discrete
trap position (passive mode), the distance between the two beads was recorded at 2.5 kHz for 10–
300 s. Initially, only fast hopping events characteristic of dsDNA unwinding were present. Once theunzipping fork arrived at the proximal dimer interaction of the nucleosome, additional slow transition
events, due to histones binding with and dissociating from dsDNA or ssDNA, could be seen. Recording was terminated when the force reached ~23 pN. The tether was then relaxed to ~0.5 pN. For the
purpose of aligning the traces and accurately converting nanometer distance to basepairs unzipped,
an unzipped trace using constant trap separation speed (as described above) was further obtained
from the same tether.

Construction of the 8  repeat ‘molecular ruler’ plasmid
The plasmid that contains a single repeat sequence (pGEM3z-1repeat) was first cloned by modifying a pGEM3z-T7A1 plasmid (Gabizon et al., 2018). The construction of the plasmid with eight tandem repeat sequences (pGEM3z-8repeat) was carried out by following a published protocol(Wu et al., 2016) using BglI, DraIII and EagI restriction sites. This method allows doubling of theChen et al. eLife 2019;8:e48281. DOI: https://doi.org/10.7554/eLife.48281

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repeat number following each cycle of cloning. To ease isolation and purification of the 8  repeat
DNA for future ligation steps, we removed the internal BsaI site in pGEM3z-8repeat and introduced two BsaI sites flanking the 8  repeat region by using an overlap PCR strategy. Briefly, twofragments outside the 8  repeat region of the plasmid were amplified using oligos ZC12-ZC13 andZC14-ZC15, respectively, and assembled into one fragment using ZC12-ZC15. The assembled fragment, which is devoid of the internal BsaI site, was digested with SapI/EagI and ligated with SapI/
EagI digested 8  repeat fragment from pGM3z-8repeat. The resulting plasmid pGM3z-8repeat2BsaI contains the 8  repeat sequence flanking by two BsaI sites, which are included in oligos
ZC12 and ZC15. All plasmids containing repeat sequences were transformed and grew in SURE2
competent cells at 30˚C. Large amounts of pGM3z-8repeat-2BsaI plasmids were purified from150 mL of SURE2 cells using Zyppy Plasmid Maxiprep Kit.

DNA templates for pol II nucleosomal transcription assay
The 8  repeat DNA with proper overhangs were digested from pGM3z-8repeat-2BsaI using
BsaI-HF and purified using 8% native PAGE.
The crosslinked DNA (XLink) used to stall Pol II at the end of the template was prepared byannealing ZC16 and ZC17. The annealed oligos were diluted to 1 mM in TE with 20% DMSO and 50
mM trioxsalen, irradiated by 340 nm UV light for 15 min. Extra trioxsalen (10 mM more) was added
and the oligos were irradiated for another 15 min. This procedure was repeated to ensure complete
crosslinking. The crosslinked oligos were bound to 1 mL HiTrap Q column (GE Healthcare), washedwith 4 mL TE buffer, then 4 ml TE buffer + 250 mM NaCl, eluted with TE + 1 M NaCl, and desaltedusing Amicon Ultra centrifugation filters with Ultracel 3K membrane. The sequences of the crosslinked oligos are:5’ phos-GGTGTACAGAACGCAATGAATT 3’
3’ GGACCACATGTCTTGCGTTACTTAA 5’
NPS DNA (308 bp) that contains the 147 bp ‘601’ NPS was amplified from a pGMZ-3z/601 plasmid using oligos ZC18 and ZC19. The NPS DNA was digested with BsaI/DraIII, purified using HiTrap
Q column, and ligated to the crosslinked oligo. The ligation product (NPS-Xlink) was purified using
8% native PAGE.
The 2 kb upstream spacer DNA and 1.5 kb biotin handle DNA were amplified from lambda DNA
using oligos ZC20-ZC21, ZC22-ZC23, respectively. PCR products were digested with BsaI and purified using 1% agarose gel. Both the 2 kb spacer and 1.5 kb biotin handle DNA were ligated to 5’CGGT 1 mm polystyrene oligo beads overnight at 16˚C using T4 DNA ligase (NEB). The ligatedbeads were first washed with TE + 0.5 M KCl + 20 mg/mL b-casein, then washed twice with TE + 20
mg/mL b-casein and resuspended in TE + 20 mg/mL b-casein to a concentration of 0.02% (w/v) for
1.5 kb biotin handle, and 0.2% (w/v) for 2 kb spacer DNA. The ligated beads were stored at 4˚C until
experiments.

Assembly of yeast pol II stalled complex
Biotinylated yeast Pol II holoenzyme was expressed, purified and biotinylated as previously
described (Kireeva et al., 2003) and was a generous gift of Prof. Craig Kaplan. The stalled Pol II
elongation complex was prepared by a bubble initiation method followed by uridine triphosphate(UTP) starvation (Hodges et al., 2009). The sequences for the template DNA strand (TDS), non-template DNA strand (NDS) and short RNA (RNA9) are:
NDS:
5’AGGTCTCAGAAGACGCCCGAACAACAGACACAAACACCACGGCCGGCGAGCCAGACACGACCAATTATCTATGTAACTTGCCATATTCAGGATTAT 3’RNA9:5’ GACGCCCGA 3’TDS:
3’TCCAGAGTCTTCTGCGGGCTTGTTGTCTGTGTTTGTGGTGCCGGCCGCTCGGTCTGTGCTGG
TTAATAGATACATTGAACGGTATAAGTCCTAATAGTCA-phos 5’
To assemble the stalled complex, TDS was incubated with RNA9, heated to 45˚C and cooled
down to 20˚C at 1 ˚C/min to form the TDS/RNA9 hybrid. Pol II was added to the hybrid and incubated at room temperature (RT) for 10 min, followed by NDS addition and incubation at 37˚C for 15

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min. Transcription was initiated by adding ATP/GTP/CTP to a final concentration of 10 mM each andthe reaction was incubated at RT for 10 min. If Pol II succeeded in restarting, it will be stalled at thefirst A site on TDS (bolded and underlined in the sequence above) due to absence of UTP. The relocation of Pol II to the stall site will also expose a BsaI site (underlined above) shielded initially by Pol
II and only those complexes in which Pol II succeeded in restarting can be digested and further
ligated to the 2 kb upstream spacer. The stalled complex was digested with BsaI-HF at 37˚C for 15
min, aliquoted, and stored at 80˚C until usage.

Nucleosome reconstitution on NPS-Xlink template
Xenopus WT (xWT) nucleosome was reconstituted by salt-dialysis using NPS-Xlink DNA with Xenopus laevis recombinant histone octamer. Human WT (hWT), uH2B and H2A.Z nucleosomes werereconstituted similarly to those used in the single-molecule unzipping assay, except that NPS-Xlink
DNA was used. The efficiency of nucleosome reconstitution was assessed by 4% native PAGE. In
case where a significant amount of free DNA was present, the nucleosome was further purified bysucrose gradient ultracentrifugation. The nucleosomes were concentrated, supplemented with 0.02
% NP40 and stored at 4˚C.

Optical tweezers assay of pol II transcription through the nucleosome
Transcription was performed in TB50 buffer. NTPs concentration was 0.5 mM each of ATP, CTP,
GTP, UTP. The 1.5 kb biotins beads were pre-incubated with 0.5 mM neutravidin for 10 min at room
temperature and diluted with TB50. Pol II sample beads were prepared by ligating the 1 mm 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 hr. 0.02% of NP40 was also
included in the ligation reaction. The overhangs of the various components were optimized suchthat the ligation occurs at desired orders. The sample beads were diluted with TB50 + 0.02% NP40.
The full sequence of the assembled transcription template was available at the end of the
document.
To perform the experiment, we first captured a 1.5 kb biotin bead in one trap followed by a Pol II
sample bead in the other trap. The two beads were rubbed against each other until a tether isformed. If the tether has expected length, the pair of beads was moved to the experimental position, which is close to the outlet of the NTPs channel. Force feedback was turned on to maintain aconstant force of 10 pN and the NTPs channel was opened to start transcription. Data acquisition
was started right after force feedback was turned on and terminated once the polymerase reached
the end or arrested for more than 300 s without dynamics. To probe the fate of transcribed nucleosomes, force feedback was turned off and the trap distance was reset to a value that gives less than
1 pN force on the tether. The two beads were pulled away from each other by increasing trap distance at a constant speed of 20 nm/s, until the force reaches above 40 pN. From these pulling
curves, we rarely detected rips normally associated with nucleosome unwrapping. Trap distance wasfurther increased to break the tether and the beads were calibrated. All transcription data wasrecorded at 800 Hz.

In vitro pol II transcription on the 1  repeat template
The 1  repeat DNA template was amplified from pGM3z-1repeat plasmid using oligos ZC24ZC25, digested with BsaI-HF and purified by agarose gel extraction. To determine the main pausesite in the 64 bp repeat sequence, Pol II stalled complex was radioactively labeled with [a-32P]-ATP
during initial pulsing. The stalled complex was loaded on streptavidin-coated magnetic beads. The
beads were washed with TB130 (20 mM Tris, pH = 8.0, 130 mM KCl, 5 mM MgCl2, 10 mM DTT, 20
mg/mL BSA) and ligated to the 1  repeat DNA template using T4 DNA ligase for 1 hr at RT. Transcription was chased by adding 40 mM NTPs mix (ATP, UTP, CTP, GTP, final concentration of 40 mMeach) to the stalled complex beads, and terminated by adding 2  urea stop buffer (8 M urea, 50
mM EDTA) at 10, 20, 40, 60, 120, 300, 600 and 900 s. In parallel, transcription was chased by adding
40 mM NTPs mix together with 50 mM of each type of 3’-deoxynucleotide RNA chain terminators(3’dATP, 3’dCTP, 3’dGTP, 3’dUTP, TriLink Biotechnologies). The reactions were allowed to proceed
at room temperature for 10 min before terminated by adding the 2  urea stop buffer. Samples
were extracted with Phenol: Chloroform: Isoamyl alcohol (1:0.9:0.01), precipitated with ethanol andChen et al. eLife 2019;8:e48281. DOI: https://doi.org/10.7554/eLife.48281

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dissolved in 2  formamide sample buffer (95% formamide, 5 mM EDTA, pH 8.0, with bromophenol
blue and xylene cyanol). RNA was resolved on 12% denaturing PAGE, dried and exposed to a phosphorimager screen. Images were captured on the Typhoon imager (GE Healthcare) and processedby ImageJ.

Mechanical model for pol II transcription through the nucleosome
Pol II dynamics
Our model for Pol II dynamics is illustrated in Figure 7B. In this model, Pol II takes forward main
pathway steps by one base pair at a rate k0 or can enter a backtracked pathway by stepping backone base pair at rate kb1. Once backtracked, Pol II takes steps one base pair forward at rate kfn andsteps one base pair backward at rate kbn. Stepping forward from the first backtracked state returns
Pol II to the main pathway.
Transition rates depend on force f, with main pathway and backtracking step rates given by
k0 ðf Þ ¼ k00 ed0 ‘f =ðkB TÞ ;0 d0 ‘f =ðkB TÞ
0
kfn ¼ kfneand kbn ¼ kbn
e ð1 dfb Þ‘f =ðkB TÞ

k00, kfn0, and kbn0 are the zero force rate constants. d0 and dfb are splitting factors, representing
the transition state location. ‘ = 0.34 nm is the step size, the length of one DNA base pair.
kBT = 4.11pNnm is the thermal energy at room temperature.
Our model is adapted from the Pol II dynamics model and parameterization of
Dangkulwanich et al. (2013). Dangkulwanich models Pol II forward stepping as three stages, withthe first two reversible, and the third effectively irreversible. Our experimental condition of highnucleotide concentration leads to a nearly instantaneous second transition, and we combine the tworemaining transitions into a single irreversible transition with rate k0. The zero-force forward rates of
the two remaining stages in Dangkulwanich are 88 s 1 and 35 s 1, combined into k00 = 25 s 1. The
rate of initial backstepping, kb1, is only from the first of the three main pathway states in Dangkulwanich. Accordingly, we weight this zero-force initial backtracking rate, 6.9 s 1, by the probability of
being in the main pathway state eligible for backtracking, kb10 = (35/66)6.9 s 1. Backtracking is
restricted to a maximum of three base pairs, such that kbn = 0 for n  4. The remaining parameters
are kfn0 = 1.3 s 1 for all n, kbn0 = 1.3 s 1 for 1  n  3, d 0 = 0.64, and dfb = 0.5, taken directly from
Dangkulwanich.

Nucleosome effect on polymerase kinetics
The model above describes transitions of the polymerase on DNA, but does not incorporate theeffect of the nucleosome, which is expected to hinder forward stepping. We adapt a previous model(Koslover and Spakowitz, 2012) to describe the DNA polymerase-nucleosome system on a twodimensional energy landscape (Ej,w). The first dimension (j) is the position of the polymerase and thesecond (w) is the number of DNA base pairs unwrapped from the nucleosome. This energy landscape incorporates the mechanics of the DNA, polymerase, and nucleosome interaction (namely,
steric exclusion between polymerase and nucleosome and bending of the unwrapped DNA) as
described in the section below.
For a given length of unwrapped DNA, there is a change in energy associated with the polymerase stepping forward,
DEj;w ¼ Ejþ1;w

Ej;w ;

which modulates the rate of that step according to
k0j;w ðf Þ ¼ k00 exp½d0 ð‘f

DEj;w Þ=ðkB TÞ:This assumes that the step forward involves a transition state at fractional position d 0 and that theenergy landscape is linear between positions j and j+1.
We assume that the wrapping and unwrapping of DNA from the nucleosome is much faster thanthe polymerase stepping kinetics. In this case, the system is equilibrated along the w dimension, andChen et al. eLife 2019;8:e48281. DOI: https://doi.org/10.7554/eLife.48281

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the overall stepping rate for the polymerase can be described as a weighted average over all the
stepping rates: d0 ‘f  P e Ejw e d0 DEjw
ðj;effÞ
wP
k0¼ k00 e kB T
Ejw
we
An analogous calculation is done for the forward and backward stepping rates in the backtracked
state:
 P E ð1 d ÞDE
ð1 dfb Þ ‘fj 1;w
fb
e jw eðj;effÞ
0
w P
kbn
¼ kbn
e kB TEjw
ew
P Ejw d DE
dfb ‘f ee fb j;w
ðj;effÞ
0 kB T
wP
kfn
¼ kfn
e
Ejw
w

e

(all the energies in the above are expressed in units of kBT). Overall, the presence of the nucleosome modifies the polymerase kinetics by making it much slower to step forward if doing so would
require a substantial increase in energy associated with bending of the linker DNA ahead of thepolymerase.

Energy landscape for polymerase–nucleosome system
The free energy Ejw is defined by the location of the polymerase at basepair j (relative to the start of
the nucleosome) and the number of DNA base pairs unwrapped from the nucleosome, w.
ðLÞEjw ¼ EN þ Eint
The first term EN(L) is the conformation energy for the DNA linker N base pairs in length between
Pol II and the nucleosome (Koslover and Spakowitz, 2012). We use a highly simplified mechanicalmodel for this system, where the histone core of the nucleosome is treated as a steric sphere ofradius Rnuc = 3.2 nm and Pol II is treated as a steric sphere of radius Rpol = 7 nm. The DNA is modeled as a wormlike chain with 35.4 nm persistence length, that must stretch from the center of thepolymerase to positions along a spiral wrapped around the nucleosome (Figure 7A). For a given
length of DNA unwrapped ahead of the polymerase (‘N), the bending energy is calculated by optimizing the wormlike chain configuration subject to the constraint that the steric spheres for polymerase and nucleosome may not overlap. If very little DNA is unwrapped ahead of the polymerase, thelinker is short and must bend tightly to avoid steric overlap (leading to high energies). If more of the
DNA is unwrapped, the linker may not need to bend at all (EN(L)=0 for lengths above approximately
30 bp).
The second contribution Eint is the energy of DNA interaction with the nucleosome. This includes
unfavorable bending of the DNA around the nucleosome and favorable DNA-nucleosome binding
interactions. Ntot = 147 base pairs can bind to the nucleosome, and each can have a different interaction energy. For w DNA base pairs unwrapped from the nucleosome
Eint ¼

Ntot
X

fi ;

i¼wþ1

where ji is the interaction energy of i’th base pair with the nucleosome.

Determining dwell times and fittingWith the quantitative model of polymerase dynamics, we can determine mean dwell times. We analytically determine the mean time for the polymerase to reach the n+1’th state after first reaching
the n’th state (Koslover and Spakowitz, 2012).
This model assumes the binding/unbinding of the DNA ahead of the polymerase is always equilibrated as the polymerase steps backward and forward. This is a reasonable assumption, given the
rapid equilibration time for DNA unwrapping, but only up to the point when the DNA fully unwraps
from the nucleosome. Our model neglects the additional entropic contributions of DNA and polymerase separating completely in solution and cannot properly predict the dwell times at the very
end of the polymerase transcribing through the nucleosome.

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Using the lsqcurvefit routine in Matlab, we fit the DNA-nucleosome interaction energies ji tomatch the quantitative model mean dwell times to the experimental mean dwell times, smoothed bytaking the local average over a 3 bp span. As shown in Figure 7D, we only include experimental
mean dwell time where the polymerase is positioned within the nucleosomal binding sequence(j  0). Prior to these base pairs, we use a mean dwell time (k00) 1 = 0.04 s.

Data analysis
Single-molecule unzipping data analysis
Using the calibration data, we calculated the complete force-extension curves for each tether. The
analysis consisted of the following steps:
First, the relaxation of the fully unzipped construct (that is, after the complete unzipping of theconstruct and before rezipping of dsDNA has begun, corresponding to a force range of ~20–40 pN)
was fit to a model in which 1850 bp of dsDNA are described as a worm-like chain with a persistence
length of 35.4 nm, a stretch modulus of 1020 pN and a contour of 0.34 nm/bp (Bouchiat et al.,
1999; Bustamante et al., 1994), and 872 bases of ssDNA are described using an extensible freely
jointed chain with a contour length of 0.59 nm/base (Mills et al., 1999; Smith et al., 1992). The
dsDNA parameters were estimated by analyzing the pulling curves of 4.7 kb dsDNA molecules. The
other parameters (stretch modulus and Kuhn lengths for the ssDNA and an offset of the extension
to account for bead size variation) were fit, resulting in a Kuhn length of 1.45 ± 0.02 nm, a stretch
modulus of 975 ± 61 pN and an offset of 29 ± 2 nm (errors are 95% confidence intervals over all
traces, N = 234). These values are close to previously published values (Smith et al., 1996). Using
these parameters, we calculated the number of unzipped base pairs at all positions along the pulling
trace.
Second, we performed a minor adjustment on the extension to align the two NPS repeats on thebare DNA template. In principle, identical positions in the two NPS repeats should be 197 bp apartin distance, and they are expected to behave identically in the trace (same force-extension signatures). However, the calculated distance obtained initially is typically different from this value of 197
bp. At this point, we rescaled the data along the x-axis (number of unzipped base pairs) to maintain
436 unzipped base pairs at the end of the unzipping curve and a distance of 197 base pairs between
identical positions on the two NPS sequences. To find the correct scaling factor, we rescaled thedata using a range of scaling factors (from 170 to 197) using the following equation:
Nrescaled ¼ 436

197
 ð436factorNnotrescaled ÞFor each factor, we binned the data points in 0.5 bp window and calculated the force-weighted
residence histogram along the sequence. We then calculated the correlation between the histogramat positions along the first NPS and the histogram at positions along the second NPS:
XResð first copy iÞ  Resð first copy i þ 197Þ
Correlation ¼
first copy i

The factor giving the maximum correlation was selected, and the data was finally rescaled using
this factor. Using this approach, we generated a mean residence histogram of the first NPS from allbare DNA unzipping traces. The rescaling factors were typically between 180 to 190. The requirement for rescaling to satisfy the periodicity may result from bead size variations or deviations fromthe models used to describe the pulling traces.
Third, once an aligned mean residence histogram of the first NPS was obtained from unzipping
traces of bare DNA, a slightly modified operation was performed on unzipping traces of the nucleosome datasets. Again, the relaxation after complete unzipping was fit and the number of unzippedbase pairs were calculated, and again rescaled using a range of rescaling factors. This time, the correlation between the residence time histogram of the first NPS in the nucleosome traces and themean residence histogram of the first NPS obtained in the previous step was calculated and maximized. The rescaling factors for nucleosome data had the same range as for the bare DNA data.

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Residence time analysis of unzipping traces
After obtaining the fitting parameters for both dsDNA and ssDNA, bead-to-bead distances of theunzipping traces were converted to unzipped basepairs. The unzipped basepairs of the traces werethen aligned, scaled and normalized to the beginning of the second NPS by subtracting 248 bp (thesecond NPS begins at 249 bp of the Y stem region). For traces obtained at constant trap separation
speed (20 nm/s), a force weighted residence time (RT) between each bp was calculated by summing
the forces of all data points between two consecutive unzipped basepairs (Figure 1E). Therefore,
long residence time (i.e. more data points) while under higher force within a particular bp would
result in a high force-weighted RT in this analysis. The force-weighted RT accounts for force differences along the unzipping trace and serves as a proxy of the strength of histone-DNA interactions of
the nucleosome. For constant force unzipping traces, residence time at each bp was calculated bycounting intervening data points N. Because data frequency is 800 Hz, RT therefore equals to N/800(Figure 2C). RT histograms are plotted as mean values from all traces.

Analysis of the number of unzipping transitions in unzipping traces
A transition in the unzipping trace is defined as a peak in the residence time histogram that is above
a certain threshold. For each unzipping trace obtained at constant trap separation speed, we identified transition events by looking for maxima in the RT histogram and manually applying a thresholdto avoid too many transitions (rips) from just bare DNA. The chosen threshold cannot be too high,as the RT for H2A.Z unzipping traces generally have more peaks but lower amplitude for each peak.
This analysis (Figure 1—figure supplement 1F) revealed that on average, H2A.Z nucleosome unzipping traces have at least one more ripping transition than those of WT nucleosomes.

Analysis of the partial unzipping data to test mobility
The final unzipping trace or the bare DNA unzipping trace was used to fit the WLC model to obtain
the elasticity, offset and scaling parameters of a particular tether. These parameters are applied toprevious partial unzips from the same tether. All traces for a particular tether are plotted together
without any further alignments. Note, for both WT and H2A.Z nucleosomes, the initial force risealways occurs at the same position without lateral shifts. During the force rise at the proximal dimer
region, the unzipping fork randomly dwells at nearby locations (~5 bp away), consistent with nucleosome hopping in this region.

Analysis of hopping (equilibrium) data at constant trap positions
To explore steady-state behavior of DNA on the nucleosome, the trap separation was held fixedsuch that the DNA experiences wrapping and unwrapping fluctuations in the proximal dimer region,‘hopping’ on and off the nucleosome. A trace of force-extension pairs is measured at each trap separation (Figure 3—figure supplement 1F), followed by a final unzipping and relaxation trace at constant trap velocity. The following subsections describe our analysis methods for extracting from thisdata an underlying energy landscape for DNA base pairing energies and the energies of interaction
with the nucleosome.

Calculation of unzipping energies from force-extension traces
Because the pulling and extension curves for bare DNA overlap closely with no hysteresis(Figure 1B), we assume this process is at equilibrium. The energy associated with unzipping eachbasepair can then be computed from the work done by the pulling force during unzipping, with acorrection for the work required to extend the newly unzipped bases.
To start, we find the fractional extension of dsDNA worm-like chains and ssDNA freely jointed
chains at a given force, zds(F) and zss(F), respectively (Wang et al., 1997). The length of ssDNA Lss
between the two dsDNA handles of length Lds for each force-extension pair (F,s) is then given by
Lss ¼

s

2zds ðFÞLds
zss ðFÞ

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Nunzip ¼

Lss
;
2lss

where ‘ss is the ssDNA length per base pair, with a factor of two because twice the base pair lengthof ssDNA is obtained when unzipping one base pair of dsDNA. Each number of base pairs unzipped
Nunzip can now correspond to a specific force F, length of ssDNA Lss, and fractional extension of
ssDNA zss. The unzipping energy of each base pair of ssDNA is the overall work to extend the two
newly unzipped bases minus the work required to stretch those bases to the observed extension.
Z 2lss zss
DEunzip ðNunzip Þ ¼ 2Fzss lssFFJCðxÞdx :0

Alignment of force-extension data
Because the bead radius cannot be known precisely, individual data collection runs are shifted arbitrarily along the extension axis. We use the final complete pulling curve to account for this shift. The
pulling curves for each experimental run with bare DNA are first mutually aligned (Figure 3—figure
supplement 1C) and the average trace is used to calculate the absolute shift along the extension
axis.
Specifically, we calculate the unzipping energy for each basepair as described in the previous section. The two copies of the NPS give rise to duplicate features in the base-pair interaction energy
landscape, whose separation depends on the absolute values of the end-to-end extension input into
the calculation. We therefore shift the averaged force-extension curve along the x axis in such a waythat the duplicate energy features are separated by precisely 197 bp (Figure 3—figure supplement
1D). The same shift is assumed for the equilibrated hopping data obtained for each individual DNA
molecule prior to the corresponding pulling trace. No scaling of the x-axis is done in this analysis.
In our calculations we used dsDNA persistence length of 35.4 nm, dsDNA stretching modulus
1020 pN, 0.34 nm contour length per base pair, ssDNA segment length 1.03 nm, ssDNA stretchingmodulus 1000 pN, 0.59 nm contour length per base pair. The ssDNA parameters were obtained byfitting the final region of the averaged pulling curve for bare DNA traces, where the hairpin has
been completely unzipped. The calculated force-extension relation for a molecule with a 434 bp
unzippable region, terminated with a 4-base hairpin, and connected to two dsDNA handles (1848bp), given the fitted unzipping energies, is shown in Figure 3—figure supplement 1E.
Pulling traces with bound nucleosomes present are aligned to the averaged pulling trace for bare
DNA based on the force-extension curve features prior to reaching the second NPS (specifically,
extensions below 870 nm are used for alignment).

Extracting DNA-nucleosome interaction energies from equilibrated hoppingdata
For each trap separation, the number of base pairs unzipped (N) is obtained for each force-extension
pair, populating a distribution in N (Figure 3B). Assuming the system is in thermodynamic equilibrium, the probability for each number of base pairs unzipped is converted to a relative energy foreach number of base pairs unzipped. Subtracting the energy of DNA stretching and the energy forthe off-center beads in the optical traps gives the cumulative relative energy to unzip the given number of base pairs. The difference in this cumulative relative energy between consecutive base pairs isthe energy to unzip each base pair. The various fixed trap separations provide overlapping ranges
for the energy of unzipping for each basepair (Figure 3—figure supplement 1G), and the averagevalue from all trap separations that span a particular value of N is used for further analysis.
To find the energy of the DNA-nucleosome interactions, the unzipping energy for bare DNA (nonucleosome present) is subtracted from the unzipping energy for a system with a nucleosome present (WT, H2A.Z, and uH2B).
Given the extracted energies of unzipping bare DNA and peeling DNA off the nucleosome forthe region accessed by the equilibrium hopping data, we can calculate the predicted force extension
relation in an equilibrium pulling curve (Figure 3—figure supplement 1H). We note that theobserved forces in the nucleosome-bound region during the constant velocity pulling traces are substantially higher, emphasizing that these traces are obtained out of equilibrium.

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Nucleosome transcription data analysis
The alignment of the ‘molecular ruler’, data analysis on pausing, backtracking and residence time of
Pol II was performed essentially the same as recently described (Gabizon et al., 2018). Briefly, foreach trace the region expected to contain the repeats (8  64 bp) was aligned to find the physicallength of the repeat in nanometers, and the aligned traces were aligned between themselves and to
the known pause sites discovered by biochemical studies (described below). The pause site withineach repeat is located at the 59th nucleotide (T) of the 64 bp DNA and the periodicity of the physicallength of each repeat is found to be 21.1 nm at 10pN force. The position of the polymerase along
the nucleosome was obtained by extrapolating the position from the aligned repeat region. To plot
the transcribed distance (bp) of the leading edge of Pol II relative to NPS, we applied an offset of 16
bp to account for the footprint of Pol II.
The crossing time is calculated as the total duration of the leading edge of Pol II crossing theentire 147 bp NPS region. Only traces that reached the stall site are included in crossing time
analysis.
Example traces of Pol II hopping at certain regions (Figure 5—figure supplement 1) were analyzed with a classic Hidden Markov Model (HMM) by fitting to two (for hWT or H2A.Z) or three states(for uH2B).
Probability of arresting is calculated as the percentage of traces that entered NPS but did notreach the stall site, while probability of crossing is the percentage of traces that successfully reached
the stall site. Typically, we considered a trace that paused 300 s or longer without any associateddynamics to be arrested. For arrested traces, percentages of traces that arrested before or after the
dyad are also calculated based on their arrest position.
Pause-free velocities (bp/s) of Pol II before, inside and after NPS are estimated by calculating theinverse of the median residence time (s/bp) at distinct sites. To account for sequence bias, the threefastest sites (lowest median residence time) are chosen from each sampling range. For regions
before or after NPS, sites up to 100 bp away from the NPS region are sampled.

Full sequence of the unzipping template
The sequence below shows the stem of the Y structure that contains two consecutive pieces of NPS
DNA (bold and italic). The red sequence is the stem of the end hairpin (four bases of the loop notshown here). The 147 bp core ‘601’ NPS is underlined in each segment. The upstream DNA isbridged to the two arms of the Y. The DNA in between are ligation sites. The full length of ssDNA
after complete unzipping will be 872 bases (including extra bases from the loop of the hairpin).
(arms). . .TTTTGACTACTGACGCGGACATTCAGGAGATGGACCCTATACGCGGCCGCCCTGGAGAATCCCGGTGCCGAGGCCGCTCAATTGGTCGTAGACAGCTCTAGCACCGCTTAAACGCACGTACGCGCTGTCCCCCGCGTTTTAACCGCCAAGGGGATTACTCCCTAGTCTCCAGGCACGTGTCAGATATATACATCCTGTGCATGTATTGAACAGCGACCTTGCAACGATGGACCCTATACGCGGCCGCCCTGGAGAATCCCGGTGCCGAGGCCGCTCAATTGGTCGTAGACAGCTCTAGCACCGCTTAAACGCACGTACGCGCTGTCCCCCGCGTTTTAACCGCCAAGGGGATTACTCCCTAG
TCTCCAGGCACGTGTCAGATATATACATCCTGTGCATGTATTGAACAGCGACCTTG
CACCCTCCACTCTAGA

Full sequence of the Pol II transcription template
Pol II loading sequence (NDS/TDS), 8  repeat DNA, core ‘601’ NPS and crosslinked DNA (Xlink)
are in blue, bold, italic and red, respectively. The transcription starvation site (T in NDS) is bolded
and underlined. The NPS-Xlink DNA used for octamer loading is underlined.
AGGTCTCAGAAGACGCCCGAACAACAGACACAAACACCACGGCCGGCGAGCCAGACACGACCAATTATCTATGTAACTTGCCATATTCAGGATTATCAGTAGCGGAAGAGCGAGCTCGG
TACCCGATCCAGATCCCGAACGCCTATCTTAAAGTTTAAACATAAAGACCAGACCTAAAGACCAGACCTAAAGACACTACATAAAGACCAGACCTAAAGACGCCTTGTTGTTAGCCATAAAGTGATAACCTTTAATCATTGTCTTTATTAATACAACTtACTATAAGaAGAGACAACTTAAAGAGACTTAAAAGATTAATTTAAAATTTATCAAAAAGAGTATTGACTTAAAGTCTAACCTATAGGATACTTACAGCCATCGAGAGGGACACGGGGAAACACCACCAGCCTCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGAC

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TAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGTGGATCCGCCGGCCGCAACGATGGACCCTATACGCGGCCGCCCTGGAGAATCCCGG
TGCCGAGGCCGCTCAATTGGTCGTAGACAGCTCTAGCACCGCTTAAACGCACGTACGCGCTG
TCCCCCGCGTTTTAACCGCCAAGGGGATTACTCCCTAGTCTCCAGGCACGTGTCAGATATATACATCCTGTGCATGTATTGAACAGCGACCTTGCCGGTGCCAGTCGGATAGTGTTCCGAGCTCCCACTCTAGAGGATCCCCGGGTACCGAGCTCGAATTCGCCCTATAGTGAGTCGTATTACAATTCACTGGCCGTCGCACCCTGGTGTACAGAACGCAATGAATT

Full sequence of the 1  repeat template for in vitro transcription
The 1  64 bp repeat sequence is in bold. CAACGCCTCCCGGGCTCACCATCATCCTGACTAGTCTTTCAGGCGATGTGTGCTGGAAAGATCTTATGTCACCCCGTGGATCCGCCGGCCGTCATCACCA
TCATCCTGACTAGAGTCCTTGGCGAACCGGTGTTTGACGTCCAGGAATGTCAAATCCGTGGCGTGACCTATTCCGCACCGCTGCG

Data and code availability
Raw data will be made available from the Dryad Digital Repository: https://doi.org/10.5061/dryad.
8sb6h8n.
MATLAB scripts have been deposited in GitHub (Chen, 2019; copy archived at https://github.
com/elifesciences-publications/dataprocessDNAunzipping).

Acknowledgements
We thank Guillermo Chacaltana and Robert Sosa for technical assistance, and all members of Bustamante laboratory for critical discussion. This work was supported by Howard Hughes Medical Institute, NIH Grants R01GM032543 and R01GM071552 to CB, R01GM098401 to TY, R01GM097260 to
CK, the US Department of Energy Office of Basic Energy Sciences Nanomachine Program underContract DE-AC02-05CH11231 to CB, a grant from Alfred P Sloan Foundation to EFK (FG-2018–
10394), and a Burroughs Welcome Fund Collaborative Research Travel Grant to TY; CB is a Howard
Hughes Medical Institute Investigator.

Additional information
Funding
FunderGrant reference number

AuthorNational Institute of General
Medical Sciences

R01GM032543

Carlos Bustamante

National Institute of General
Medical Sciences

R01GM071552

Carlos Bustamante

National Institute of General
Medical Sciences

R01GM098401

Tingting Yao

National Institute of General
Medical Sciences

R01GM097260

Craig D Kaplan

Basic Energy Sciences

Nanomachine Program
under Contract DE-AC0205CH11231

Carlos Bustamante

Alfred P. Sloan Foundation

FG-2018-10394

Elena F Koslover

Burroughs Wellcome Fund

Collaborative Research
Travel Grant

Tingting Yao

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Howard Hughes Medical InstituteCarlos Bustamante

The funders had no role in study design, data collection and interpretation, or thedecision to submit the work for publication.
Author contributions
Zhijie Chen, Conceptualization, Resources, Data curation, Formal analysis, Investigation,
Methodology, Writing—original draft, Project administration, Writing—review and editing; Ronen
Gabizon, Conceptualization, Resources, Data curation, Formal analysis, Investigation, Methodology,
Writing—review and editing; Aidan I Brown, Software, Formal analysis, Investigation, Writing—review and editing, Analyzed unzipping data and developed the mechanical model; Antony Lee,
Software, Formal analysis, Writing—review and editing, Cloned the initial 1 x repeat template,
Developed the algorithm for analyzing transcription data; Aixin Song, Resources, Purified humanhistones and assembled human octamers; César Dı́az-Celis, Resources, Methodology, Assembled
Xenopus nucleosomes; Craig D Kaplan, Resources, Funding acquisition, Provided biotinylated yeast
Pol II; Elena F Koslover, Software, Formal analysis, Funding acquisition, Investigation, Writing—review and editing, Analyzed unzipping data and developed the mechanical model; Tingting Yao,
Conceptualization, Resources, Data curation, Supervision, Funding acquisition, Investigation,
Methodology, Project administration, Writing—review and editing; Carlos Bustamante,
Conceptualization, Supervision, Funding acquisition, Project administration, Writing—review andediting
Author ORCIDs
Zhijie Chen
https://orcid.org/0000-0003-1376-5750
Ronen Gabizon
https://orcid.org/0000-0002-3626-5073
Aidan I Brown
https://orcid.org/0000-0002-6600-8289
Antony Lee
https://orcid.org/0000-0003-2193-5369
Aixin Songhttps://orcid.org/0000-0002-4377-7528
César Dı́az-Celis
https://orcid.org/0000-0002-6659-8434
Craig D Kaplan
https://orcid.org/0000-0002-7518-695X
Elena F Koslover
https://orcid.org/0000-0003-4139-9209
Tingting Yao
https://orcid.org/0000-0003-4101-9691
Carlos Bustamante
https://orcid.org/0000-0002-2970-0073
Decision letter and Author response
Decision letter https://doi.org/10.7554/eLife.48281.034
Author response https://doi.org/10.7554/eLife.48281.035

Additional files
Supplementary files. Supplementary file 1. Oligos used in this study.
DOI: https://doi.org/10.7554/eLife.48281.028. Supplementary file 2. Pol II backtrack analysis.

DOI: https://doi.org/10.7554/eLife.48281.029. Transparent reporting form

DOI: https://doi.org/10.7554/eLife.48281.030

Data availability
Matlab scripts for processing unzipping curves and hopping data have been deposited in GitHub athttps://github.com/lenafabr/dataprocessDNAunzipping (copy archived athttps://github.com/elifesciences-publications/dataprocessDNAunzipping). Raw data are available from Dryad https://doi.org/
10.5061/dryad.8sb6h8n. Further information and requests for resources and reagents should bedirected to and will be fulfilled by the Lead Contact, Carlos J Bustamante (carlosb@berkeley.edu).

Chen et al. eLife 2019;8:e48281. DOI: https://doi.org/10.7554/eLife.48281

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Research article

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The following dataset was generated:
Author(s)Year Dataset title

Chen Z, Gabizon R, 2019 Data from: High-resolution andhigh-accuracy topographic andBrown A, Lee A,
transcriptional maps of theSong A, Diaz-Celis
nucleosome barrier
C, Kaplan C, Koslover E, Yao T,
Bustamante C

Dataset URL
https://doi.org/10.5061/
dryad.8sb6h8n

Database and
Identifier
Dryad Digital
Repository, 10.5061/
dryad.8sb6h8n

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