In-vitro mapping of nucleosome positions at base-pair resolution using ortho-phenanthroline

Abstract

The positions of nucleosomes along genomic DNA play a role in defining patterns of gene expression and chromatin organization. Determination of nucleosome positions in vivo and in vitro, as revealed by the locations of histones on DNA, have provided insight into mechanisms of nucleosome sliding, spacing, assembly, and disassembly. Here, we describe methods for the in vitro determination of histone-DNA contacts at base-pair (bp) resolution. The protocol involves the labeling of histones with ortho-phenanthroline (OP), site-specific cleavage of nucleosomal DNA, and processing and analysis of the resulting DNA fragments. This methodology provides an efficient and high-resolution means for studying kinetics and behaviors of enzymes that alter nucleosome structure and/or positioning, and can be used to identify preferred distributions of nucleosomes on natural DNA sequences.

Basic Protocol 1: Cysteine-specific chemical modification of folded histones with ortho-phenanthroline (OP)

Basic Protocol 2: Nucleosome sliding assay adapted for OP mapping of histone-DNA contacts

Basic Protocol 3: OP-mediated cleavage, processing, and analysis of DNA fragments using a sequencing gel

Support Protocol 1: Preparation of dideoxy sequencing ladders

Support Protocol 2: Preparation and running of a denaturing DNA sequencing gel

INTRODUCTION:

Nucleosomes are the basic unit of chromatin, consisting of ~146 base pairs (bp) of DNA wrapped around a central histone core. This DNA wrapping around histones limits the accessibility of DNA-interacting proteins —such as transcription factors (TFs)— to the underlying DNA and, thus, such nucleosomal organization has an inherently repressive nature (Lorch, LaPointe & Kornberg, 1987; Han & Grunstein, 1988; Kornberg & Lorch, 1999). Nucleosomes package the majority of genomic DNA, and their presence is tightly coupled to transcriptional regulation (Li, Carey & Workman, 2007). Due to their ability to block access of TFs, nucleosomes appear to regulate transcriptional activation through changes in their positioning on DNA (Nocetti & Whitehouse, 2016). In most genes, nucleosomes are depleted from active and poised promoters, with a strongly positioned nucleosome at or adjacent to the transcriptional start site (Jiang & Pugh, 2009). In gene bodies, nucleosomes are organized into regularly spaced arrays, which block inappropriate transcriptional initiation (Jiang & Pugh, 2009; Smolle & Workman, 2013; Prajapati, Ocampo & Clark, 2020).

Nucleosomes are physically reorganized by several families of chromatin remodeling factors, which can remove, reassemble, and reposition nucleosomes in various contexts (Prajapati, Ocampo & Clark, 2020; Gkikopoulos et al., 2011; Krietenstein et al., 2016). The positioning of nucleosomes and the action of chromatin remodelers are closely tied to transcriptional programs and, thus, are of fundamental importance for cell identity and pluripotency, and are known to be dysregulated in many human diseases, such as cancers (Kadoch & Crabtree, 2015; Michael et al., 2020; Tao et al., 2017).

In genome-wide studies, a common methodology for determining nucleosome positioning is MNase-seq (Wal & Pugh, 2012; Zhang & Pugh, 2011). Using micrococcal nuclease (MNase), this technique digests accessible DNA between nucleosomes, releasing mononucleosomes from bulk chromatin. Since the histone core largely protects DNA from digestion, MNase treatment produces a protection pattern that reflects the positioning of nucleosomes. With high-throughput sequencing, the locations of nucleosome-sized DNA fragments are identified and mapped back to the genome. MNase-seq has given great insights into the patterns of nucleosome positioning genome-wide, demonstrating a remarkable phasing of nucleosomes in gene bodies relative to transcription start sites (Jiang & Pugh, 2009; Gkikopoulos et al., 2011).

A limitation of MNase-seq, however, is that protection from enzymatic digestion can also be achieved by non-histone proteins, which has raised controversy about the identity of factors protecting some DNA fragments, particularly in nucleosome-depleted regions (Brahma & Henikoff, 2019). Since MNase maps nucleosomes solely by DNA protection, another limitation is that it is impossible to determine the precise positioning of histones when a footprint smaller than the expected ~146 bp is generated. Canonical nucleosomes consist of histones H2A, H2B, H3, and H4, which form an octamer from two copies of a central H3/H4 heterodimer (forming the (H3/H4)₂ tetramer) and two copies of flanking H2A/H2B heterodimers (Luger et al., 1997; McGinty & Tan, 2015). Since canonical nucleosomes have pseudo two-fold symmetry, the central nucleosome dyad is considered to be the center of the protected ~146 bp DNA fragment. However, histone-DNA interactions are continually disrupted, particularly during transcription, DNA repair, and DNA recombination, where canonical histones can be swapped out with histone variants (Henikoff & Ahmad, 2005; Talbert & Henikoff, 2014; Venkatesh & Workman, 2015; Kassem et al., 2020). Even without full disruption, nucleosomes can transiently unwrap from one or both sides (Li et al., 2005). Such DNA unwrapping allows for additional MNase cut sites, yielding smaller footprints (Henikoff et al., 2011). Additionally, one of the H2A/H2B dimers can be lost from the nucleosome, resulting in a hexasome, which protects only ~100 bp of DNA. For these shorter fragments, MNase-seq cannot accurately determine the position of the H3/H4 tetramer (Henikoff et al., 2011; Ramachandran, Ahmad & Henikoff, 2017).

An alternative method for determining nucleosome positioning is through site-specific DNA cleavage. DNA can be cleaved site-specifically on the nucleosome either through photo-crosslinking (Kassabov & Bartholomew, 2004) or local production of hydroxyl radicals (Flaus et al., 1996; Flaus & Richmond, 1999). In both cases, a cysteine residue, introduced at or adjacent to the DNA binding surface of the histone core, is used to target a reactive group. For photo cross-linking, the cysteine can be labeled with 4-azidophenacyl bromide (APB), and the resulting cross-links can be resolved into DNA nicks. For the local production of hydroxyl radicals, the cysteine can be labeled with derivatives of either EDTA or ortho-phenanthroline (OP). DNA cleavage by these methods is remarkably specific and reports on the location of the cysteine on the nucleosome. For in vivo experiments, DNA cleavage sites (usually made via OP) can be mapped with high-throughput sequencing, whereas for in vitro experiments, cleavage sites can be directly visualized on sequencing gels. Since the location of the labeled cysteine is known from the nucleosome structure (Luger et al., 1997), the cleavage site defines the translational and rotational phasing of DNA with respect to the histone core. DNA cleavage by both photo-crosslinking and site-directed hydroxyl radicals is highly localized, allowing for nucleosome positioning to be determined at base-pair (bp) resolution.

Although photo-crosslinking is a powerful technique for mapping local movements of DNA on the nucleosome (Winger et al., 2018), a disadvantage compared to OP-based mapping is that it requires about an order of magnitude more material. With OP, DNA cleavage arises from the production of hydroxyl radicals from the oxidation of Cu⁺ by H₂O₂ (Marshall et al., 1981). Unlike photo-crosslinking, which can only undergo a single reaction per site, hydroxyl radicals can be continuously generated by reducing agents that regenerate Cu⁺ (Marshall et al., 1981).

As mentioned above, an advantage of site-directed cleavage over MNase is that the precise location of the nucleosome can be determined, whether partially unwrapped or as a sub-nucleosomal assembly. Even for fully wrapped nucleosomes, cleavage by MNase typically produces a distribution of sites where DNA exits from the nucleosome, even for a single nucleosome position. MNase cleavage is, therefore, not preferred for high-resolution in vitro experiments where the goal is to map, for instance, how nucleosome positions change in response to chromatin remodeling enzymes. For in vitro experiments, a popular nucleosome positioning sequence is the Widom 601, which forms nucleosomes with a single reproducible position of DNA on the histone core (Lowary & Widom, 1998). With site-directed cleavage, one can determine how a single 601 positioned nucleosome redistributes in response to a chromatin remodeler (Ghassabi Kondalaji & Bowman, 2022), or determine preferred nucleosome positions on a natural DNA sequence.

In this article, we describe methods for mapping nucleosome positions in vitro, using OP-based DNA cleavage. A flowchart of the core procedures is shown in Figure 1. Basic Protocol 1 describes how to use OP to modify folded histones containing single cysteines. These labeled histones are then used to assemble nucleosomes. Basic Protocol 2 provides an example procedure for repositioning OP-labeled nucleosomes, using the chromatin remodeler Chd1. Basic Protocol 3 describes how to perform DNA cleavage reactions on OP-labeled nucleosomes. The fragments are then extracted, resolved on a denaturing sequencing gel, and compared with a dideoxy sequencing ladder to determine the histone-DNA contact sites. Support Protocols 1 and 2 explain how to generate the dideoxy sequencing ladders and to separate DNA fragments on urea denaturing gels, respectively.

Figure 1. Flowchart for determining histone-DNA contacts using ortho-phenanthroline (OP).

Protocols described in this article are highlighted in non-gray colors, whereas steps and reagents described elsewhere are shown in gray. Basic Protocol 1 (yellow) describes how to label folded histones with ortho-phenanthroline (OP), which can then be used for generating nucleosomes. Basic Protocol 2 (pink) gives an optional example assay for altering nucleosome positions using the Chd1 remodeler. Basic Protocol 3 (green) describes how to create and process site-specifically cleaved DNA using OP-labeled nucleosomes, with the products analyzed by urea denaturing gel (Support Protocols 1 and 2, blue and orange).

Taken together, the methods presented here should enable a user to study nucleosome positioning in vitro at high resolution, allowing for a deeper understanding of factors that alter histone-DNA contacts, and how DNA sequence influences preferred nucleosome positions.

STRATEGIC PLANNING:

The OP-labeling reaction takes place via alkylation of the sulfhydryl group of a cysteine residue. Thus, a histone variant containing a cysteine close to DNA must be purified and properly folded into histone dimers, tetramers, or octamers (Luger, Rechsteiner & Richmond, 1999) before conducting the experiments described in Basic Protocol 1, for histone labeling. After generating an OP-labeled nucleosome, it may be of interest to some users to carry out experiments that change the position of the histone core on DNA. Basic Protocol 2 provides an example procedure for repositioning an OP-labeled nucleosome using a chromatin remodeler. For this procedure, the chromatin remodeler must be purified beforehand. This nucleosome sliding protocol is not required for nucleosome mapping with OP, and may be bypassed if one only wishes to determine the position(s) of the histone core on DNA after nucleosome assembly/reconstitution.

Historically, cysteines introduced at either H2B(T87C) or H4(S47C) have been used for OP-guided histone mapping in both in vitro and in vivo experiments (Brogaard et al., 2012; Henikoff et al., 2014; Fei et al., 2015; Ramachandran & Henikoff, 2016). More recently, OP modification of H3(Q85C) in yeast has been used to determine the position of nucleosomes across the genome (Chereji et al., 2018) (Figure 2). Since these cleavage sites are on different histones and, thus, report on different locations around the nucleosome, selection of the site for modification by OP depends on the goal and nature of the experiment. OP labeling at H4(S47C) is ideal for reporting on the position of the central H3/H4 tetramer in vitro, as this position cleaves DNA adjacent to the two-fold (dyad) axis of the nucleosome. For in vivo mapping of yeast nucleosomes, H3(Q85) has been shown to be superior, as cleavage at this site releases a 51 bp fragment symmetrically spanning the central dyad that more robustly reports on nucleosome positions by high-throughput sequencing (Chereji et al., 2018). H2B(T87C) can also be used to determine the dyad locations and, thus, the positioning of nucleosomes; however, since this labeling site is located on the H2A/H2B dimer, it will not be present on both sides of hexasomes, which lack one H2A/H2B dimer (Ghassabi Kondalaji & Bowman, 2022). This can be nevertheless advantageous, particularly for in vitro experiments where specific interactions between DNA and an H2A/H2B are being studied. If the wild-type histone possesses a cysteine residue (e.g., Xenopus laevis H3C110), its mutation to alanine (e.g., H3(C110A)) is recommended.

Figure 2. OP-directed cleavage patterns on the nucleosome.

Schematic representation of DNA cleavage sites for nucleosomes assembled from histones labeled with OP at three positions: (a) H4(S47C), (b) H2B(T87C), and (c) H3(Q85C) (numbering for vertebrate canonical histones). The numbers show the distance in bp from the nucleosome dyad (position zero), with positive numbers indicating cleavage sites 3’ relative to the dyad, and negative numbers indicating cleavage sites 5’ relative to the dyad. Each OP moiety locally cleaves both strands of the DNA duplex, resulting in two distinct cleavages for each DNA strand.

Prior to assembling nucleosomes, DNA and histones must be separately prepared. DNA can be generated by large-scale PCR and then purified, as described elsewhere (Nodelman et al., 2020; Dyer et al., 2004). After separate purification of each of the four histones, histones are combined and folded together into octamers (H2A, H2B, H3 and H4), tetramers (H3/H4 only), and/or dimers (H2A/H2B) (Dyer et al., 2004) (see Figure 1, in gray). OP labeling (Basic Protocol 1) is carried out on the folded histones containing the target cysteine (see Reagents and Solutions). In general, nucleosomes can be assembled by combining DNA with the octamer in a 1:1 ratio, or DNA with H2A/H2B dimer plus H3/H4 tetramer in a 1:2:1 ratio (Dyer et al., 2004). We have found that nucleosome reconstitutions using only histone octamers often have a significant level of hexasome contamination, which lack one H2A/H2B dimer (Levendosky et al., 2016). Therefore, we typically add excess dimer to enrich for full nucleosomes. Thus, if the OP modification is on the H2A/H2B dimer, we recommend labeling both histone octamers and dimers, combining them in a 3.5:1 molar ratio of OP-labeled octamer to its matching OP-labeled dimer. OP-labeled nucleosomes can then be purified similarly to unlabeled nucleosomes (Nodelman et al., 2020; Dyer et al., 2004) and stored indefinitely at −80°C.

Selection of the DNA sequence depends on the target experiment. When nucleosomes are reconstituted with well-positioning DNA sequences via salt gradient dialysis, OP-guided DNA cleavage reactions (Basic Protocol 3) generate distinct DNA fragments that are easily identifiable on a denaturing sequencing gel (Support Protocol 2). Thus, if the target experiment does not require a specific DNA sequence (e.g., natural DNA sequences), a recommended DNA sequence for assembling nucleosomes from labeled folded histones is the Widom 601 nucleosome positioning sequence (Lowary & Widom, 1998). In addition to sequence, and depending on the target experiment, one should also consider whether DNA extensions (called linker or flanking DNA) beyond the nucleosome core should be included. For instance, to study nucleosome repositioning by some remodeling enzymes such as Chd1 (Basic Protocol 2), the presence of flanking DNA on at least one side of the nucleosome is essential (Stockdale et al., 2006; Nodelman et al., 2021).

To be able to detect the DNA fragments generated from OP-guided cleavage reactions (Basic Protocol 3) on a denaturing sequencing gel (Support Protocol 2), the DNA used for nucleosome assembly must also be labeled. Although radioactivity can be used, we prefer using DNA fluorescently labeled on 5’ ends. Fluorescent labels are easily incorporated using 5’-labeled oligos during PCR, and unlike radiolabeled DNA, both strands can be simultaneously followed by using two different fluorophores (such as FAM and Cy3 or Cy5), as long as an appropriate fluorescence scanner is available, such as a Typhoon (Cytiva, formerly GE). Further, fluorescently-labeled nucleosomes can be stored up to several years at −80°C without significant loss of signal, unlike radiolabeled DNA. In addition to generating the DNA needed for nucleosome reconstitution, fluorescently labeled oligos are also used for preparing dideoxy sequencing reactions (Support Protocol 1), needed as ladders for interpreting DNA cleavage sites at bp resolution.

As mentioned earlier, OP-labeled nucleosomes are used for a target experiment that requires histone mapping at bp resolution. This target experiment can be as simple as confirming the positioning of the histone core on Widom 601 sequence, to more complicated experiments such as nucleosome sliding by chromatin remodelers (see Basic Protocol 2). No matter what the target experiment is, it must be designed in a way to generate sample(s) with 0.5–2 pmol OP-labeled nucleosome(s) as substrates for site-specific DNA cleavage reactions (Basic Protocol 3). Additionally, the total volume of the OP-labeled nucleosome samples before the start of the DNA cleavage reaction in Basic Protocol 3 must be ≤ 60 μL. The buffers and solution components for the target experiment must be compatible with the cleavage reaction. For instance, the buffer must be free from (or below a threshold value of) divalent cations that can form coordinated complexes with OP and lack components (like EDTA) that chelate copper.

BASIC PROTOCOL 1: Cysteine-specific chemical modification of folded histones with ortho-phenanthroline (OP)

This protocol describes the chemical modification of folded histone core proteins with ortho-phenanthroline (OP). The modification takes place through an alkylation reaction of a cysteine residue with an OP derivative containing an iodoacetyl moiety. As mentioned in Strategic Planning, two sites commonly used for determining histone core positioning in vitro are H2B(T87C) and H4(S47C).

The procedure starts by determining the desired scale of the OP-labeling reaction. Various amounts of histones, ranging from 1.5 to 90 nmol, can be successfully labeled by this protocol. The amount of folded histones to label depends on the number of the reactions to be performed in the target experiment (e.g., nucleosome sliding in Basic Protocol 2). However, since the labeled products can be stored indefinitely at −80°C and used in future experiments, it is often desirable to aim for larger-scale reactions, as they are more efficient. Additionally, multiple OP-labeling reactions on dimer and octamer of H2B(T87C), and tetramer and octamer of H4(S47C) can be performed simultaneously with this protocol.

It is recommended to start the reaction in the evening, allotting at least 2 hours for labeling before proceeding with an overnight incubation of the reaction mixture. Labeling buffer and post-labeling dialysis solution should be prepared not earlier than the day of performing the labeling reaction. However, to increase efficiency, OP solutions can be made in advance and stored at room temperature.

Note: Iodoacetyl OP derivative and substituted OP moiety are light-sensitive. Minimize the exposure of OP-labeled histones and assembled nucleosome to ambient or high-intensity light as much as possible.

Materials:

• Purified folded histones (see Reagents and Solutions)

• Labeling buffer (see Reagents and Solutions)

• 5 mM TCEP solution (see Reagents and Solutions)

• 4.5 mM OP solution (see Reagents and Solutions)

• 2-Mercaptoethanol (βME; Sigma-Aldrich, cat. no. M6250, ≥99.0% purity, 14.3 M)

• Ice

• Post-labeling dialysis buffer (prepare two 2-L solutions, see Reagents and Solutions)

• Ultrapure water (e.g., Milli-Q water)

• Sterile-filtered (0.22 μm) glycerol (Sigma-Aldrich, cat. no. G7893, ≥99.5% purity, 100% (v/v))

• Liquid nitrogen

• Lamp with low intensity light bulb (e.g., incandescent 15 W light bulb), for the dark room

• Tube rotator (e.g., Globe Scientific, cat. no. 1141T92)

• Aluminum foil

• 15-mL conical tubes

• 1.5-mL sterilized microcentrifuge tubes (e.g., Fisher, cat. no. 05-408-129)

• 2.0-mL sterilized microcentrifuge tubes (e.g., Fisher, cat. no. 05-408-138)

• Microdialysis chambers (e.g., Fisher mini dialysis device, cat. no. PI88400; or prepared in-house from 1.5-mL sterilized microcentrifuge tubes as described in (Nodelman et al., 2020; Vary, Fazzio & Tsukiyama, 2004))

• Ventilation (fume) hood

• Forceps

• Razor blades

• Bunsen burner

• Scissors

• 50-mL conical tubes

• 3.5-kDa molecular weight cutoff (MWCO) dialysis tubing (e.g., Spectra/Por 3.5 kDa MWCO, cat. no. 132724)

• Plastic dialysis clips

• Float buoy

• Glass beakers (2 × 2-L)

• Magnetic stir bars and a stir plate

• Parafilm

• Kimwipes

• 10-KDa spin concentrators (e.g., Amicon Ultra-0.5 ml, cat. no. UFC501096 with 0.5-mL filtration capacity, Amicon Ultra-4, cat. no. UFC801024 with 4-mL filtration capacity or Amicon Ultra-15, cat. no. UFC901024)

• Benchtop centrifuge at 4°C or microcentrifuge (e.g., Eppendorf 5424) in a cold room

• 0.6-mL sterilized microcentrifuge tubes (e.g., Fisher, cat. no. 05-408-120)

• −80°C freezer

Protocol steps

Determine the quantity and volumes required for the labeling reaction

• 1
  Calculate the volume of purified folded histone stocks to be labeled:

  $$\text{Volume of histone core needed }\left(\mu \text{L}\right)=\frac{\text{\# of nanomoles of histone core to label }\left(\text{nmol}\right){\text{ × 10}}^{\text{3}}\left(\raisebox{1ex}{$\text{nmol}$}\!\left/ \!\raisebox{-1ex}{$\mu \text{mol}$}\right.\right)}{\text{Concentration of purified histone core stock }\left(\mu \text{M}\right)}$$

  For example, if the concentration of histone octamer stock solution is 300 μM, and 15 nmol of histone octamer is to be labelled, then 50 μL of stock solution should be used.

• 2
  Calculate the initial volume of labeling reaction. This volume ensures that the final concentration of folded histones in the labeling mixture is 10 μM, and is used in the next step to determine the volume of the labeling buffer require for the reaction.

  $$\text{Initial volume of labeling reaction }\left(\mu \text{L}\right)\text{= }\frac{\text{\# of nanomoles of histone core to label }\left(\text{nmol}\right){\text{× 10}}^{\text{3}}\left(\raisebox{1ex}{$\text{nmol}$}\!\left/ \!\raisebox{-1ex}{$\mu \text{mol}$}\right.\right)}{\text{10 }\left(\mu \text{M}\right)}$$

  For example, to label 15 nmol of histone core octamer, the initial reaction volume is 1500 μL.

• 3
  Calculate the amount of labeling buffer required for the labeling reaction.

  $$\text{Volume of labeling buffer needed for labeling mixture }\left(\mu \text{L}\right)\text{ =Initial volume of labeling reaction }\left(\mu \text{L, see step 2}\right)\text{-Volume of histone core added }\left(\mu \text{L, see step 1}\right)$$

  For example, if the concentration of octamer stock solution is 300 μM, and 15 nmol of histone octamer is to be labelled, then 50 μL of octamer stock solution should be added to 1450 μL of labeling buffer.

• 4
  Calculate the volume of a 5 mM TCEP solution, pH 7.5, needed for the reaction (to be used in step 12), such that the labeling mixture has 10 nmol TCEP for every nmol of added folded histone.

  $$\text{Volume of 5 mM pH-balanced TCEP solution }\left(\mu \text{L}\right)\text{= }\frac{\text{10 ×}\left(\text{\# of nanomoles of histone core to label }\left(\text{nmol}\right)\right){\text{×10}}^{\text{3}}\left(\raisebox{1ex}{$\text{nmol}$}\!\left/ \!\raisebox{-1ex}{$\mu \text{mol}$}\right.\right)}{\text{5 }\left(\text{mM}\right){\text{ ×10}}^{\text{3}}\left(\raisebox{1ex}{$\mu \text{M}$}\!\left/ \!\raisebox{-1ex}{$\text{mM}$}\right.\right)}$$

  For example, if 15 nmol of histone octamer is to be labelled, then 30 μL of 5 mM TCEP solution (pH-balanced) would be needed.

• 5
  Calculate the volume of the 4.5 mM OP solution to be added to the labeling reaction (to be used in step 13). A 30-fold excess of OP solution relative to number of moles of folded histone allows for efficient labeling.

  $$\text{Volume of 4}.\text{5 mM OP solution }\left(\mu \text{L}\right)\text{= }\frac{\text{30 ×}\left(\text{\# of nanomoles of histone core to label }\left(\text{nmol}\right)\right){\text{ ×10}}^{\text{3}}\left(\raisebox{1ex}{$\text{nmol}$}\!\left/ \!\raisebox{-1ex}{$\mu \text{mol}$}\right.\right)}{\text{4}.\text{5 }\left(\text{mM}\right){\text{ ×10}}^{\text{3}}\left(\raisebox{1ex}{$\mu \text{M}$}\!\left/ \!\raisebox{-1ex}{$\text{mM}$}\right.\right)}$$

  For example, if 15 nmol of histone octamer is to be labelled, then 100 μL of 4.5 mM OP solution should be used for the labeling reaction.

• 6
  Determine the total volume of the histone-labeling mixture (prior to βME addition).

  $$\text{Total volume of histone labeling mixture}\left(\mu \text{L}\right)\text{= Summation of volumes calculated in steps 2, 4, and 5}$$

  For example, if 15 nmol of histone octamer is to be labelled, then the total volume of labeling mixture after addition of 5 mM TCEP solution (pH-balanced) and 4.5 mM OP solution, would be 1630 μL.

• 7Calculate the volume of neat βME liquid (14.3 M) required to quench the labeling reaction. The final concentration of βME should be within 40–50 mM range.

  For example, if 15 nmol of histone octamer is to be labelled, then 4.6–5.7 μL of concentrated βME (14.3 M) would be sufficient to quench the labeling reaction.

• 8
  Calculate the total volume of quenched histone labeling reaction.

  $$\text{Total volume of quenched reaction }\left(\mu \text{L}\right)\text{=Summation of volumes calculated in steps 6 and 7}$$

Perform the labeling reaction on folded histones

• 9Place the OP solution, a lamp with a low intensity light bulb, a tube rotator, and aluminum foil in the dark room.
• 10Pipette the calculated volume of labeling buffer (determined in step 3) into a room-temperature (RT) microcentrifuge or 15-mL conical tube, keeping the tube at RT.

  Use a sterilized 1.5-mL or 2-mL microcentrifuge tube or a 15-mL conical tube, depending on the calculated total volume of the reaction (see step 8). The tube should be not more than half-filled, to allow the solution to efficiently be mixed on a tube rotator (see step 14).

• 11Thaw the snap-frozen folded histone stock on ice. Pipette the calculated volume of folded histone stock (determined in step 1) into the labeling buffer at RT. Mix thoroughly by gently pipetting up and down.
• 12Add the required volume of a 5 mM TCEP solution (pH-balanced), calculated in step 4, to the labeling mixture. Mix thoroughly and keep the reaction at RT for 10 minutes.
• 13Transfer the reaction to the dark room and add the volume of 4.5 mM OP solution calculated in step 5 to initiate the labeling reaction.
• 14Cover the reaction tube with aluminum foil and place it in a tube rotator. Allow the reaction to proceed at RT for 2 hours.

  In the meantime, prepare for dialysis. If the volume calculated in step 8 is ≤ 1.5 mL, prepare microdialysis chambers from 1.5-mL microcentrifuge tubes (Nodelman et al., 2020; Vary, Fazzio & Tsukiyama, 2004), as described below. Dialysis chambers will be used on day 2 of the labeling reaction.

  Each microdialysis chamber made from the top of a 1.5-mL microcentrifuge tube can hold up to 150 μL. Attempting to dialyze more than 150 μL per chamber may result in sample loss. Prepare enough chambers for the total volume of quenched labeling reaction.

  To prepare the microdialysis chambers, first lay sterilized 1.5-mL microcentrifuge tubes horizontally on a heat-resistant surface inside the hood. Using forceps, heat a razor blade over the flame of a Bunsen burner until it begins to turn red. While holding the bottom of a microcentrifuge tube, use the hot razor blade held by the forceps to immediately cut the tube ~5 mm below the opening, which will create a ring. With scissors, cut off the lids of each tube. Using cool forceps, briefly (one or two seconds only) hold each piece over the flame to melt and smooth the sharp edges. Do not hold over flame for longer periods to avoid burning the plastic.

  Store the microdialysis chamber (caps and rings) in a clean 50-mL conical tube to avoid contamination. This step can be done within few minutes but if the forceps become too hot, turn off the flame and allow the forceps to cool down, then continue.

  If the volume of the reaction is >1.5 mL, use a dialysis bag instead, which can be prepared on day 2 of histone labeling, according to steps 18–19. See below.

  Alternatively, commercially-available dialysis devices can be used for a broad volume range (e.g., 100 μL to 30 ml with Slide-A-Lyzer Cassettes, Thermo Scientific), which do not require assembly and can be directly used on day 2 of histone labeling.

• 15After the 2-hour reaction period (step 14), transfer the tube and the tube rotator to the cold room (4 °C) and continue mixing the reaction overnight. Following the overnight labeling reaction, transfer the reaction tube to the dark room and keep the sample on ice.

  In all subsequent steps of this protocol, avoid exposing the OP-labeled folded histone to ambient light whenever possible.

• 16Quench the labeling reaction by adding βME to a final concentration of 40–50 mM, as calculated in step 7. Mix by gently pipetting up and down, then store on ice for 15–60 min.

  After quenching, incubation with βME should be less than one hour. Proceed with steps 17 to 20 within this 1-hour period.

• 17Prepare the dialysis membrane by cutting the appropriate length of 3.5-kDa molecular weight cutoff dialysis tubing.

  Wear clean gloves when handling the membrane.

  If the total volume of the quenched reaction is more than ~1.5 mL, cut enough membrane for setting up a standard dialysis bag (dialysis tube, plastic clips, and a float buoy).

  If microdialysis chambers are being used (see step 14), cut enough of the membrane to make one ~3 × 3 cm square for each dialysis cap.

• 18Soak the membrane in a beaker containing ≥ 50 mL of ultrapure water to remove preservatives. Allow the membrane to soak at 4 °C for ~15 minutes.
• 19Rinse the membrane (inside and outside) with ultrapure water.

  If a standard dialysis bag is being used (see step 14), rinse a float buoy and plastic dialysis clips with ultrapure water.

  Check the membrane for possible leakage by holding the bottom of the tube tightly between two fingers and filling it up with ultrapure water. If the water level decreases within a few seconds, cut a new piece of dialysis membrane, and repeat steps 17–19.

  In case of using microdialysis setup, cut the dialysis membrane open into a single-layered flat sheet, then cut out a ~3 × 3 cm square for each cap.

  Do not allow the dialysis membrane(s) to dry out. Keep the rinsed membrane(s) in ultrapure water at all times.

• 20Just before starting the dialysis, and in the cold room, add 700 μL of 14.3 M βME to 2 liters of prechilled post-labeling dialysis buffer, in a 2-L beaker. Allow the solution to mix on a stir plate and then transfer the 2-L beaker to the dark room. Do not allow the solution to warm up.
• 21After the labeling mixture has incubated with the βME quench for ~1 hr (step 16), transfer the mixture to a prepared dialysis bag or the microdialysis chambers in a darkroom using a pipette. Immediately after filling and sealing, place the dialysis bag/chambers into the 2-L beaker with the cold post-labeling dialysis buffer containing βME.

  To setup microdialysis chambers, start by placing the microdialysis caps on a clean surface. Pipette up to 150 μL of quenched labeling mixture into the cap. Avoid making bubbles. Gently take out membrane squares from ultrapure water and remove excess water by holding the top of the square with one gloved hand while gently pulling it between two fingers of the other hand. Lay the center of the square on the microdialysis cap and do not change its position afterward, to avoid sample loss. Place the microdialysis ring over the membrane gently but firmly, pushing the ring down onto the cap. The ring holds the dialysis membrane against the opening in the cap, completing the chamber. Once secured, trim the membrane if the cap is not aligned on the center of square and the membrane is too large in one side. However, avoid trimming it down too much as the membrane outside the ring helps the chamber float on the surface during dialysis.

  When transferring the microdialysis chamber into the post-labeling dialysis buffer, make sure that the dialysis buffer comes in contact with the central surface of the dialysis membrane (that is, not blocked by a bubble), to ensure efficient buffer exchange.

  Repeat for the rest of the caps by preparing four chambers at a time and placing them on ice until all four chambers are made. Place the microdialysis chamber on ice with the cap facing upward, to avoid any contacts between the membrane and ice. After making each set (four chambers), trim their membranes and transfer them to dialysis buffer. Depending on the total volume of quenched reaction, the last cap may contain less than 150 μL.

  If multiple different labeling reactions are being performed simultaneously, be sure to mark the microdialysis caps (small-scale reaction) or plastic dialysis clips and float buoy (large scale reactions).

• 22Cover the 2-L beaker containing the dialysis bag/chambers with parafilm and wrap the entire beaker with aluminum foil. Using a stir plate, mix the solution in the cold room (4 °C) overnight.

  The microdialysis chambers should face down toward the solution for efficient dialysis. Use a low setting on the stir plate, to avoid flipping the chambers.

  If there are too many chambers (≥ 12), they will not all float freely on the surface of the dialysis buffer. In this case, either use a beaker with a larger opening for the dialysis or increase the mixing rate. However, care must be taken to avoid violent contact between the chambers and to prevent their contact with the stir bar at the bottom of the beaker, which can result in sample loss by dissociating the microdialysis chambers.

• 23The following day, add 700 μL of 14.3 M βME to the second 2-L batch of unused and pre-chilled post-labeling dialysis buffer and mix briefly in the cold room.
• 24Move both 2 L beakers to the dark room – the 1^st 2-L post-labeling dialysis buffer containing the dialysis bag/chambers, and the 2^nd 2-L post-labeling dialysis buffer with freshly added βME.
• 25Transfer the dialysis bag/chambers from the 1^st dialysis solution to the 2^nd post-labeling buffer. Wrap the beaker with the dialysis bags/chambers with aluminum foil and move back to the cold room.
• 26Continue dialysis in the cold room for at least 3 additional hours.
• 27Before the end of the dialysis, pre-chill microcentrifuge tubes or conical tubes that will accommodate the volume of the dialyzed reaction. At the end of the dialysis, transfer the beaker to the dark room. Use a pipette to transfer the sample from the dialysis bag/chambers into a pre-chilled microcentrifuge tube or conical tube. Cover the tube with aluminum foil and keep it on ice for further processing.

  To take out the sample from the microdialysis chamber, first place the caps, membrane-side up, on a clean surface. Gently wick away the excess buffer with a Kimwipe, but do not touch the membrane. After dabbing away the dialysis solution, slightly tilt the chamber and, using a P200 (set higher than 150 μL), gently pierce the membrane inside the ring and at the bottom. Make sure to pipette out all the sample by sweeping the P200 tip around the interior walls of the bottom cap and collecting all of the solution.

Prepare the OP-labeled folded histone for long-term storage

• 28
  Using a pre-chilled spin concentrator, concentrate the OP-labeled sample to a final concentration between 60–500 μM using a cold room microcentrifuge or a refrigerated centrifuge pre-chilled and run at 4 °C. The concentration of the OP-labeled folded histone is calculated using the equation below:

  $$\text{Estimated labeled histone core concentration }\left(\mu \text{M}\right)\text{= }\frac{\text{initial \# of nanomoles of folded histone used for labeling reaction }\left(\text{step1, nmol}\right){\text{× 10}}^{\text{3}}\left(\raisebox{1ex}{$\text{pmol}$}\!\left/ \!\raisebox{-1ex}{$\text{nmol}$}\right.\right)}{\text{volume of concentrated labeled sample }\left(\mu \text{L}\right)}$$

  Make sure the sample has minimal exposure to ambient light during this step.

  During the concentrating process, a small amount of the sample can pass through the 10-kDa molecular weight cutoff filter at very high speeds. To avoid sample loss, be sure that the centrifuge speeds are less than the maximum g force recommended for the concentrators.

  The size of the 10-kDa molecular weight cutoff spin concentrator (see Materials) to use to concentrate the folded histone is dependent on the initial and final volume of the sample. For initial sample volumes of less than 2 mL, use Amicon Ultra-0.5, which gives a minimum final volume of ~15 μL. For initial volumes between 2–15 mL, use Amicon Ultra-4, which gives a final volume of ≥ 50 μL. For volumes above 15 mL, the preferred concentrator is Amicon Ultra-15, which gives a final volume of ≥ 200 μL.

• 29After concentration, transfer the retained sample from the spin concentrator to a pre-chilled 1.5-mL microcentrifuge tube.

  To minimize sample loss, rinse the spin concentrator twice with 10–40 μL of either fresh buffer or the flow-through.

• 30Bring up the sample to at least 10% (v/v) glycerol using 100% (v/v) sterile-filtered (0.22 μm) glycerol.

  To pipette glycerol, either use a wide-bore tip or use a sterilized razor blade to cut ~1 cm off the end of a P200 pipette tip. Then, gently and slowly pipette glycerol and add into the labeled folded histone sample. Mix by pipetting up and down until the solution is homogeneous.

  When pipetting glycerol, and to minimize the retention of glycerol on the outside of the tip, allow the tip to barely touch the surface of the liquid and avoid dipping the tip inside the 100% glycerol.

• 31
  Calculate the theoretical concentration of labeled sample after glycerol addition using the equation below:

  $$\text{Theoretical sample concentration after glycerol addition }\left(\mu \text{M}\right)\text{= }\frac{\text{initial \# of nanomoles of folded histone }\left(\text{step 1; nmol}\right){\text{× 10}}^{\text{3}}\left(\raisebox{1ex}{$\text{pmol}$}\!\left/ \!\raisebox{-1ex}{$\text{nmol}$}\right.\right)}{\text{volume of concentrated labeled sample }\left(\mu \text{L}\right)\text{+ Volume of 100\% glycerol}\left(\mu \text{L}\right)}$$
• 32Pre-chill a sufficient number of 0.6- or 1.5-mL microcentrifuge tubes to distribute the entire sample in 10-μL aliquots. After aliquoting, immediately snap-freeze tubes in liquid nitrogen and store at −80 °C.

  The frozen OP-labeled folded histone can be stored at −80°C for a few years.

BASIC PROTOCOL 2: Nucleosome sliding assay adapted for OP mapping of histone-DNA contacts

This protocol describes how to reposition nucleosomes with a chromatin remodeling enzyme, using nucleosomes made from OP-labeled histones generated in Basic Protocol 1. This sliding protocol serves as an example, showing how an experiment should be planned for further analysis by OP-guided DNA cleavage, which is described in Basic Protocol 3. We illustrate the procedure using the Saccharomyces cerevisiae Chd1 remodeler. Similar procedures have been used for nucleosome sliding with other remodelers such as S. cerevisiae RSC, human SNF2h, and Drosophila ACF (Ghassabi Kondalaji & Bowman, 2022; Levendosky & Bowman, 2019)

Nucleosomes used for this protocol should be reconstituted via salt gradient dialysis (Nodelman et al., 2020; Dyer et al., 2004) using purified DNA and OP-labeled histone octamers (Basic Protocol 1). The DNA fragments used here contain the 145-bp Widom 601 sequence and are prepared via large-scale PCR reactions (See Strategic Planning). Both DNA strands are typically labeled at the 5’-end with a fluorophore, such as FAM, Cy3 or Cy5. A fluorescent dye is required on at least one DNA strand to determine the sites of DNA cleavage (Basic Protocol 3), which corresponds to the position of the histone core during nucleosome sliding. Using one fluorophore on each DNA strand has the advantage of tracking both DNA strands simultaneously.

For a given set of reaction conditions, OP-labeled nucleosomes are first preincubated with the remodeler in the absence of ATP. This is called the “main mix”, and nucleosome sliding is initiated with the addition of ATP. Aliquots containing 0.8 pmol OP-labeled nucleosomes are taken at different timepoints after ATP addition and added to a quenching solution to stop the sliding reaction by eliminating the available ATP. This sampling continues until the sliding reaction has equilibrated, where nucleosome positions no longer change over time. Prior to ATP addition, aliquots are removed to later identify the histone core position at timepoint zero (t = 0). Nucleosome positions can be monitored at high-resolution with site-specific DNA cleavage analysis (Basic Protocol 3).

EDTA has long been used as an effective quencher of ATP-dependent reactions, as it sequesters Mg²⁺, which is essential for sliding reactions. As an alternative, sliding reaction can be quenched through addition of a stop DNA such as salmon sperm DNA prior to DNA-cleavage reaction. However, stop DNA increases the background and, thus, decreases the signal-to-noise-ratio in OP-mediated DNA cleavage patterns (Ghassabi Kondalaji & Bowman, 2022). To bypass the need for EDTA and salmon sperm DNA in conventional sliding assays, nucleosome sliding reactions here are stopped by the rapid enzymatic consumption of ATP by hexokinase (Ghassabi Kondalaji & Bowman, 2022). This ATP-depletion method is an example that shows how pre-cleavage reactions must be adapted for OP-mediated DNA cleavage. Additional considerations are provided in the Commentary section.

Although OP is light sensitive, this and following protocols can be done in ambient light without affecting the results in a noticeable way. This protocol should be immediately followed by Basic Protocol 3. Thus, one should prepare to carry out Basic Protocol 3 before initiating nucleosome sliding or other experiments that may alter nucleosome positioning.

Materials:

• OP-labeled nucleosomes (see Basic Protocol 1 for OP-labeled histones, and Reagents and Solutions for purified nucleosomes)

• Purified Chd1 remodeler (see Reagents and Solutions)

• 500 mM ATP stock solution (see Reagents and Solutions)

• Sliding buffer (see Reagents and Solutions)

• Ice

• Quenching buffer (see Reagents and Solutions)

• Aluminum foil

• Sterilized 1.5-mL microcentrifuge tubes (e.g., Fisher, cat. no. 05-408-129)

• Microcentrifuge tube rack(s)

• Digital timer (e.g., Fisherbrand, cat. no. 06-662-5)

Protocol steps

Determine the quantity and volumes required for a single main mix

Steps 1–5 determine the amount of materials required for a single main mix reaction, from which several sliding timepoints can be sampled.

• 1
  Calculate the amount of an 800-nM stock of purified OP-labeled nucleosome required to prepare the main mix (which will be used in step 11). The final nucleosome concentration should be 40 nM.

  $$\text{Volume of 800 nM nucleosome stock for main mix }\left(\mu \text{L}\right)\text{= }\frac{\left(\text{\# of sliding timepoints+1}\right)\text{× 0}.\text{8 }\left(\text{pmol}\right){\text{×10}}^{\text{3}}\left(\frac{\text{pmol}}{\text{nmol}}\right)}{\text{800 }\left(\text{nM}\right)}$$

  Each aliquot removed from the sliding reaction at a specific timepoint should contain 0.8 pmol of OP-labeled nucleosome. With 800 nM concentration, that corresponds to 1 μL of stock nucleosome per reaction. We typically measure 3 to 7 timepoints.

  This calculation is set up so that the main mix should have enough volume for 1 extra timepoint (# of sliding timepoints + 1). This extra volume is essential to ensure that, after removal of all previous time points, there is sufficient volume remaining for the last timepoint.

  The stock concentration of 800 nM is a typical value used to prepare sliding mixtures. We normally obtain nucleosome stock concentrations above 1 uM, and dilute those down to 800 nM. If the available stock concentration is less than the suggested 800 nM, adjust the equation accordingly.

• 2
  Calculate the amount of 4 μM Chd1 stock required for the main mix (step 12). Final concentrations of Chd1 in sliding reactions should be 200 nM.

  $$\text{Volume of 4 μM Chd1 stock in main mix }\left(\mu \text{L}\right)\text{ = }\frac{\left(\text{\# of sliding timepoints+1}\right)\text{× 200 }\left(\text{nM}\right)\text{ ×20 }\mu \text{L}}{\text{4 }\left(\mu \text{M}\right){\text{ × 10}}^{\text{3}}\left(\raisebox{1ex}{$\text{nM}$}\!\left/ \!\raisebox{-1ex}{$\mu \text{M}$}\right.\right)}$$

  Since 4 μM to 200 nM is a 20-fold dilution, and the total reaction is 20 μL, each reaction should utilize 1 μL of the 4 μM Chd1 stock.

• 3
  Determine the volume of 1.0 mM ATP solution to be added to the main mix (step 16). The final concentration of ATP in these sliding reactions is 50 μM.

  $$\text{Volume of 1 mM ATP stock to initiate sliding reaction }\left(\mu \text{L}\right)\text{ = }\frac{\left(\text{\# of sliding timepoints}\right)\text{× 50 }\left(\mu \text{M}\right)\text{ ×20 μL}}{\text{1}.\text{0 }\left(\text{mM}\right){\text{ × 10}}^{\text{3}}\left(\raisebox{1ex}{$\mu \text{M}$}\!\left/ \!\raisebox{-1ex}{$\text{mM}$}\right.\right)}$$

  Here, the final ATP concentration (50 μM) is limiting, which slows down the sliding reaction. For saturating ATP concentrations, which would give a maximum sliding rate, use a final concentration of 1 mM instead.

• 4
  Calculate the volume of sliding buffer required (step 10).

  $$\text{Volume of added sliding buffer }\left(\mu \text{L}\right)\text{=}\left(\left(\text{\#of sliding timepoints}\right)\text{×20 }\left(\mu \text{L}\right)\right)-\text{total volume of nucleosome, remodeler, ATP }\left(\mu \text{L}\right)\text{+ 19 }\left(\mu \text{L}\right)$$

  The additional 19 μL corresponds to the aliquot taken at time zero, before addition of ATP.

Prepare the solutions and materials for the nucleosome sliding reactions

• 5Prepare fresh sliding buffer (see Reagents and Solutions) and keep on ice.

  1.5 mL of sliding buffer is usually sufficient for preparing a total of 36 sliding timepoints (different sliding mixtures, several timepoints) and for diluting the concentrated solutions of nucleosome, remodeler, and ATP.

• 6Freshly prepare 800 μL of quenching buffer (see Reagents and Solutions) and keep it on ice.

  This volume is enough to quench a total of 36 timepoints (all sliding mixtures and sampled timepoints).

• 7Prepare a 1.0 mM ATP stock solution from a 500 mM ATP stock solution (step 3), using sliding buffer, and keep it on ice.

  The total volume of 1.0 mM ATP stock solution must be enough for all sliding mixtures with 50 μM ATP.

• 8Prepare a diluted OP-labeled nucleosome stock solution with a final concentration of 800 nM. Cover it with aluminum foil and keep on ice.

  To prepare the 800 nM nucleosome stock, serially dilute the concentrated OP-labeled nucleosome stock with sliding buffer.

  Any left-over concentrated OP-labeled stock can be used for future experiments within a week. Cover the left-over stock with aluminum foil and store on ice in the cold room.

• 9Make a 4-μM Chd1 stock solution by diluting concentrated Chd1 stock solution with sliding buffer. Keep the diluted solution on ice.

Perform the nucleosome sliding reactions

• 10Add sliding buffer to a 1.5-mL microcentrifuge tube designated for sliding reaction according to the volume calculated in step 4.
• 11Add the volume of 800 nM nucleosome solution calculated in step 1. Keep the tube at RT in a microcentrifuge tube rack.
• 12Pipette the required volume of 4 μM Chd1 (calculated in step 2) into the sliding mixture. Gently but thoroughly mix by pipetting. Incubate for ~5 minutes.
• 13Meanwhile, add 20-μL aliquots of quenching buffer to new 1.5-mL microfuge tubes. Prepare enough tubes so that there is one tube per timepoint during the sliding reaction. Mark the tubes accordingly (e.g., t = 0, 15 seconds, etc.).

  Do not use smaller (e.g., 0.6-mL) microcentrifuge tubes. Further processing of quenched mixtures necessitates the use of 1.5-mL microfuge tubes.

• 14After the 5-minute incubation of remodeler and nucleosome (step 12), transfer 19 μL of the main mix to the quenching tube designated for timepoint zero (t = 0).
• 15Set a P20 pipette to 20 μL and prepare a P200 for mixing the sliding mixture.

  The P20 pipette will be used to remove aliquots from the sliding mixture at the designated time points. The P200 pipette should be set at ~50% of the main mix volume, and will be used to mix the solution as soon as ATP has been added.

• 16Start the reaction by adding ATP solution to the main mix. Pipette the required volume of ATP (calculated in step 3) into the main mix and simultaneously start the timer.
• 17After ATP addition, immediately mix the sliding mixture using the P200 pipette.
• 18At the first timepoint, remove 20 μL of the sliding mixture and add it to the designated quenching tube. Using the same pipette tip, quickly mix the quenched mixture gently but thoroughly.
• 19Repeat step 19 for subsequent timepoints.

  During longer waiting periods between time points (e.g., between 2 minutes- and 16 minutes timepoints), prepare the solutions required for subsequent processing of the quenched tubes according to steps 1–4 in Basic Protocol 3.

• 20Process the quenched mixtures according to Basic Protocol 3.

  After quenching the sliding reactions, initiating the OP-mediated cleavage reactions (steps 5–10 in Basic Protocol 3) can be delayed up to ~1 hr without any detriment. In cases where the time course extends longer than 1 hr, begin OP-mediated cleavage reactions on earlier time points, before taking later time points.

BASIC PROTOCOL 3: OP-mediated cleavage, processing, and analysis of DNA fragments using a sequencing gel

In this protocol, OP-guided DNA cleavage reactions are used to identify DNA-histone contacts at bp resolution and map the position of the histone core on DNA. These reactions are performed on OP-labeled nucleosome samples obtained from the target experiment (e.g., quenched sliding reaction in Basic Protocol 2). The process involves continuous in-situ generation of highly reactive hydroxyl radicals via repetitive cycles of redox reactions. The cycle is formed by coupling Cu²⁺ reduction by a thiol to Cu⁺/H₂O₂ oxidoreduction that returns the copper to its initial oxidation state (Cu²⁺) for the next cycle (Figure 3). Chelating copper cations to the OP moiety on the folded histone (e.g., H2B(T87C)) directs generated hydroxyl radicals to cleave the DNA backbone at a specific location (Figure 2). The reaction is quenched by addition of a copper-chelating reagent with higher affinity than OP, which renders copper cations unavailable for the OP moiety. The resulting DNA fragments are subsequently extracted, resolved on a denaturing sequencing gel (Support Protocol 2), and compared with a dideoxy sequencing ladder –prepared via Sanger (chain-termination) reaction (Support Protocol 1)– to determine the histone-DNA contact sites.

Figure 3. The generation of hydroxyl radicals through repetitive cycles of redox reactions.

Shown is a schematic of the copper redox cycles that result in DNA cleavage. The substituted ortho-phenanthroline (OP) moiety is a strong chelator of copper cations, which localizes a series of redox reactions (highlighted in pink and turquoise) in the vicinity of a unique cysteine on the nucleosome core particle. These reactions are initiated by addition of a thiol (RSH; e.g., 3-mercaptopropionic acid), which reduces the OP-chelated cupric (Cu²⁺) to cuprous (Cu⁺) cations. The presence of H₂O₂ returns the copper to its initial oxidation state while also producing reactive hydroxyl radicals. Coupling the two redox reactions (pink and turquoise) consumes the thiol and H₂O₂ while recycling the copper, resulting in continued generation of hydroxyl radicals, which can cleave the DNA backbone within a 4 Å distance of the OP moiety.

Materials:

• Cleavage buffer (see Reagents and Solutions)

• 0.8 pmol OP-labeled nucleosome samples (e.g. quenched sliding mixtures obtained from Basic Protocol 2).

• 2X Mapping buffer (see Reagents and Solutions)

• 3-Mercaptopropionic acid (MPA; Sigma-Aldrich, cat. no. M5801, ≥99%, 11.5 M)

  CAUTION: MPA is corrosive and very toxic. Wear protective gloves, a lab coat, and eye protection while handling as a neat liquid (11.5 M). Avoid inhalation and work under a ventilated hood while handling the neat liquid. Store the purchased bottle vertical and sealed at RT.

• H₂O₂ (EMD Millipore, cat. no. HX0635).

• 25 mM Neocuproine (see Reagents and Solutions)

• Post-cleavage buffer (see Reagents and Solutions)

• 25:24:1 (v/v/v) phenol:chloroform:isoamyl alcohol, saturated with 10 mM Tris. HCl, pH 8.0, 1 mM EDTA (Sigma-Aldrich, cat. no. P3803)

  CAUTION: 25:24:1 (v/v/v) phenol:chloroform:isoamyl alcohol is toxic, corrosive, and potentially carcinogenic. Work under a well-ventilated hood and avoid inhaling fumes from the solution. Wear personal protective equipment including gloves, eye goggles, and a lab coat. Discard gloves, microcentrifuge tubes, and pipette tips in a waste container designated for phenol.

• 24:1 (v/v) chloroform:isoamyl alcohol (Sigma-Aldrich, cat. no. C0549-1PT)

• 3 M NaOAC/HOAC, pH 5.2 solution (see Reagents and Solutions)

• 20 mg·mL⁻¹ Glycogen (see Reagents and Solutions)

• 100% ice-cold ethanol (Pharmco, 200 proof, cat. no. 111000200)

• Ice

• 75% (v/v) iced-cold ethanol

• Dideoxy sequencing ladders (prepared according to Support Protocol 1)

• Small-fragment formamide loading dye (see Reagents and Solutions)

  CAUTION: Formamide is toxic, suspected to be carcinogenic and may damage the unborn child. When working with formamide, wear gloves, eye protection, face shield and a protective lab coat. Avoid inhalation by working under a ventilated hood and prevent skin contact.

• Urea Denaturing DNA sequencing gel sandwich (prepared according to Support Protocol 2)

• Sterilized 1.5-mL microcentrifuge tubes (e.g., Fisher, cat. no. 05-408-129)

• Ventilation (fume) hood

• Aluminum foil

• Heat block

• 15-mL conical tubes

• Digital timer

• Vortexer

• Microcentrifuge (for use at both RT and 4°C; e.g., Eppendorf 5424 in a cold room)

Protocol steps

Day 1 - Perform OP-mediated DNA cleavage reactions

Solutions in steps 1–4 should be made fresh before starting the DNA cleavage reactions

• 1Prepare 1.0 mL of cleavage buffer in a 1.5-mL microcentrifuge tube and keep it at RT.

  1.0 mL is enough to process up to 49 samples with 0.8 pmol OP-labeled nucleosomes in each mixture (e.g. quenched sliding mixtures obtained from Basic Protocol 2).

• 2Prepare 1.0 mL of 2X mapping buffer in a 1.5-mL microcentrifuge tube and keep it at RT.

  1.0 mL is enough to process up to 49 samples with 0.8 pmol OP-labeled nucleosomes in each mixture (e.g. quenched sliding mixtures obtained from Basic Protocol 2).

• 3Prepare a 60 mM MPA solution by mixing 380.6 μL of ultrapure water (e.g., Milli-Q) with 2.0 μL of MPA (11.5 M). Vortex and keep at RT.

  Prepare 383 μL of the solution, which is enough to process 36 samples with 0.8 pmol OP-labeled nucleosomes in each mixture (e.g. quenched sliding mixtures obtained from Basic Protocol 2).

• 4Prepare a fresh 1.0-mL aliquot of 60 mM H₂O₂ by mixing 973 μL of ultrapure water (e.g., Milli-Q) and 6.0 μL of 30% (v/v) H₂O_2. Vortex briefly and keep it at RT away from direct light.

  H₂O₂ is light sensitive. Do not expose solutions containing H₂O₂ to direct light.

• 5Add cleavage buffer to each 0.8 pmol OP-labeled nucleosome sample, to achieve a final volume of 60 μL, and keep at RT.
• 6Add 20.0 μL of 2X mapping buffer to each sample.
• 7Set a P200 (tip attached) to ~70 μL, and P20 (tip attached) to 20.0 μL. Open the lid of the 15-mL conical tube containing the 25 mM neocuproine solution.
• 8Using a P10, take 10.0 μL of 60 mM MPA solution and place it on the interior walls of each tube containing the quenched reactions with 0.8 pmol OP-labeled nucleosomes. Tilt the tube slightly to prevent the droplet from falling into the quenched reaction. Immediately take 10.0 μL of 60 mM H₂O₂ and add it to the OP-nucleosome mixture while simultaneously dragging the MPA droplet from the tube walls into the same mixture. Quickly start the timer.
• 9Immediately begin mixing the solution using the pre-set P200 pipette.

  The total timescale of the cleavage reaction is 30 seconds. In this step, continue mixing the reaction for ~20 seconds (10 seconds before quenching the cleavage reaction).

• 10In the ~10 seconds before the end of the cleavage reaction (i.e. 20 seconds after starting the reaction), take 20.0 μL of 25 mM neocuproine and when the timer reaches 30 seconds, add it to quench the cleavage reaction.
• 11After performing DNA cleavage reactions on all the samples, add 120 μL of post-cleavage buffer to each tube and briefly vortex.
• 12Using a heat block, heat the cleavage mixtures at 70°C for 20-30 minutes and then allow solutions to cool down at RT.

Extract, process, and run the DNA fragments on denaturing sequencing gel

• 13Add 240 μL of 25:24:1 (v/v/v) phenol:chloroform:isoamyl alcohol (saturated with 10 mM Tris, pH 8.0, 1 mM EDTA) to each cleavage mixture. Vortex for ~5 seconds.

  Before vortexing the solutions, make sure the lids are completely closed, to avoid spillage.

• 14In a microcentrifuge, spin the tubes at maximum speed for 2 minutes at RT.
• 15Transfer the top layer (containing DNA fragments) of each sample to a separate, fresh 1.5-mL microcentrifuge tube and label accordingly. Discard the bottom layer.

  Avoid pipetting the white cloudy materials (typically protein), and only take the top layer, trying to obtain as much of top layer as possible, to minimize sample loss. Pipetting a volume of ~210 μL without taking the white materials is sufficient for this step.

  The permanent markers used to label microcentrifuge tubes are not resistant to most organic solutions/solvents, including 25:24:1 (v/v/v) phenol:chloroform:isoamyl alcohol or ethanol (used later). Therefore, care must be taken to avoid accidentally erasing the labels with residual organic solutions/solvents in the following steps. We recommend labeling both the lids and sides of the tubes.

• 16Add an equal amount (~210 μL) of 25:24:1 (v/v/v) phenol:chloroform:isoamyl alcohol, pH 8.0 to each of the collected top-layer solutions.
• 17Vortex and spin as above (step 14).
• 18Repeat step 15. Use fresh microcentrifuge tubes.
• 19Add an equal amount of 24:1 (v/v) chloroform:isoamyl alcohol to the solution-phase top layer (step 18) to remove residual phenol. Vortex and spin as above (step 14).
• 20Collect the top layer in new sets of 1.5-mL microcentrifuge tubes.
• 21Add 3 M NaOAC/HOAC, pH 5.2 to a final concentration of ~300 mM (i.e. 1/10 dilution).

  The purpose of this step is to increase the final concentration of the salt (monovalent cations such as K⁺ and Na⁺) to 300 mM. Do not attempt to increase the salt concentration by adding NaCl, as normally done for DNA precipitation from solutions containing SDS, which is present in post-cleavage buffer. NaCl has low solubility in 75% ethanol; thus, it is difficult to remove in preparing DNA for loading on the sequencing gel.

• 22Add 20 mg·mL⁻¹ glycogen to a final concentration of 0.5 mg·mL⁻¹.

  To achieve this, add 20 mg·mL⁻¹ glycogen in a 1 to 40 volumetric ratio. For example, after an 18 μL addition of 3 M NaOAC/HOAC to 180 μL of DNA-containing solution, add ~5 μL of 20 mg·mL⁻¹ glycogen.

• 23Add 100% of ice-cold ethanol to a final concentration of 75% (v/v).

  This can be achieved by adding three times the volume of 100% ethanol relative to the glycogen-added solution in the previous step. For instance, after a 5-μL addition of 20 mg·mL⁻¹ glycogen in the example in step 22, the total volume of the solution is 203 μL, and adding 609 μL of 100% ice-cold ethanol results in a final ethanol concentration of 75%, which is required for DNA precipitation by ethanol.

• 24Store the tubes at −20°C (or −80°C) overnight (12 h) to precipitate DNA.

  If the user decides to perform the day 1 and day 2 steps in this protocol in a single day (e.g., having very few nucleosome samples to be mapped), the DNA sample obtained here should be precipitated for at least 1 hour at −80 °C before continuing.

Day 2 – finish preparing DNA and analyze samples on a sequencing gel

• 25The next day (or when ready to continue), spin the tubes using a pre-chilled microcentrifuge at maximum speed for 30 minutes at 4 °C.

  Orient the hinge of all tubes up during the spin to identify the location of the pellet after centrifugation.

• 26Using a P1000, discard the supernatant by gently pipetting from the side of the tube that is opposite the precipitated DNA.

  Place the tubes on ice and do not allow them to warm up.

  DNA pellets less than 10 μg are normally not visible at this stage. The amount of extracted DNA here is typically less than 10 μg, however, in some cases, a very small white pellet may be observed.

  Whether the DNA pellet is visible or not, do not directly pipet from the vicinity of the precipitation site.

• 27Add 750 μL of ice-cold 75 % (v/v) ethanol.
• 28Spin the tubes at maximum speed at 4°C for 10–15 minutes.
• 29Discard the supernatant as in step 26.
• 30Pulse-spin at 4 °C. Then, using a P20, remove the left-over supernatant without touching the pellet or the side of the tube expected to have the adhered DNA precipitate.
• 31Air-dry the tubes for 1.5–2 hours at RT.

  Do not attempt to speed the evaporation by heating the microcentrifuge tubes. If needed, the tubes can be placed in a vacuum (e.g., speedvac centrifugal evaporator) to accelerate water solvent removal.

  Do not process the tubes until completely dry.

• 32During this 1.5–2 hour time period, prepare dideoxy sequencing ladders according to the steps described in Support Protocol 1.
• 33After 1.5–2 hours and once dried, resuspend each pellet by adding 4.0 μL of small-fragment formamide loading dye. Gently pipette the solution up and down a few times on the side of the tube where the DNA pellet should be. Store the tubes at −20 °C until ready to load on the sequencing gel.
• 34At the end of the day 1 of DNA cleavage and histone mapping, or at least 6 hours before ready to run the sequencing gel, prepare the denaturing DNA sequencing gel sandwich according to steps 1–10 in Support Protocol 2.
• 35Once the sequencing gel is polymerized (on the morning of day 2 or at least 6 hours after pouring the sequencing gel), follow steps 11–42 in Support Protocol 2 to separate and visualize the cleaved DNA fragments on the denaturing sequencing gel.

  See a discussion of sample data in Understanding Results.

SUPPORT PROTOCOL 1. Preparation of dideoxy sequencing ladders

Dideoxy sequencing ladders provide markers for determining the size and cleavage sites of DNA fragments at single nucleotide resolution. For each nucleotide (A, C, G, and T), a ladder is generated through primer extension, where occasional inclusion of the specific dideoxynucleotide (ddNTP; e.g., ddATP) terminates nucleotide addition. As a result, each Sanger mixture contains truncated single-stranded DNA fragments that all terminate with one of the four ddNTPs. These ladders are then separated on a urea denaturing sequencing gel alongside the cleaved DNA sample of interest, to identify where cleavage occurred.

When generating the dideoxy sequencing ladder, one should use the same template DNA and the primer as those used for the PCR reaction that amplified the nucleosomal DNA. As mentioned earlier, for histone mapping experiments, we often label the nucleosomal DNA by using primers with 5’ fluorophores (e.g., FAM and Cy3). Each dideoxy ladder is prepared with a single primer, therefore, tracking cleavage sites on both strands requires separate primer extension reactions with each primer.

It is also worth mentioning that 5.0-μL Sanger Reactions are required for running a single sequencing gel. If the same dideoxy ladder will be used in future sequencing gels, the Sanger reactions can be scaled up for efficiency, as the generated products can be stored indefinitely at −20°C.

Materials:

• Fluorescently-labeled DNA oligos (e.g., from IDT; primers used to prepare the nucleosomal DNA via PCR prior to nucleosome assembly, see Strategic Planning)

• DNA plasmid (same as the one used as PCR template to amplify nucleosomal DNA prior to nucleosome assembly, see Strategic Planning)

• Sequencing kit (e.g., Applied Biosystems sequenase dye primer manual cycle sequencing kit, 50 reactions, cat. no. 792601KT; including concentrated thermo reaction buffer, thermo sequenase DNA polymerase with thermostable inorganic pyrophosphatase, ddNTP, and dNTP termination mixtures)

• Small-fragment formamide loading dye (see Reagents and Solutions)

  CAUTION: Formamide is toxic, suspected to be carcinogenic, and may damage the unborn child. When working with formamide, wear gloves, eye protection, face shield, and a protective lab coat. Avoid inhalation by working under a ventilated hood and prevent skin contact.

• 0.2-mL sterilized PCR tubes

• Thermocycler (e.g., Bio-Rad T100)

• Sterilized 1.5-mL microcentrifuge tubes (e.g., Fisher, cat. no. 05-408-129)

• Heat block

Protocol steps

1. Set up a PCR master mix using reagents from the sequencing kit, and the fluorescent DNA primers and DNA template (plasmid DNA) used to amplify the DNA construct for nucleosome reconstitution. One master mix should be set up for each labeled primer. Use the following table as a guide, which is given in the manual for the Applied Biosystems sequenase kit:

   Reagents for the PCR master mix	single reaction*
   100 ng·μL−1 plasmid DNA	13.8 μL
   10x Thermo reaction buffer	2.2 μL
   1 μM fluorescently-labeled primer	1.0 μL
   Thermo sequenase DNA polymerase with thermostable inorganic pyrophosphatase	1.0 μL
   Final volume	18 μL

   ^*

   a single master mix gives enough material for three sequencing gels

2. For each master mix, label four PCR tubes: A, C, G, T. To each tube, add 1.0 μL of the appropriate ddNTP, and then add 4.0 μL of the PCR master mix.

3. Mix each tube by gently pipetting up and down. Total volume of the solution in each PCR tube should be 5.0 μL.

4. Using a thermocycler, run the following PCR program:

   PCR step	Temperature (°C)	Time
   1	95.0	1 minutes
   2	95.0	30 seconds
   3	55.0	30 seconds
   4	72.0	1 minutes
   5, go to step 2, repeat 40 times
   6	72.0	10 minutes
   7, end the run	-	-

5. After completion of the thermocycler program, transfer the contents of each PCR tube into separate 1.5-mL microcentrifuge tubes (to fit them in a heat block, see below) and add an equal volume (5.0 μL) of small-fragment formamide loading dye. Label the tubes according to the primer and ddNTP (e.g., FAM, ddCTP) used to prepare each PCR reaction.

6. Using a heat block, heat the solutions at 70°C for 2 minutes.

7. Store tubes indefinitely at −20°C.

   These dideoxy ladders should be used to determine the sites of DNA cleavage for samples processed in Basic Protocol 3. The procedure for loading the dideoxy ladders on a sequencing gel is given in Support Protocol 2, step 27.

SUPPORT PROTOCOL 2. Preparation and running of a denaturing DNA sequencing gel

This protocol describes the steps for preparing a denaturing urea sequencing gel and resolving ≤200-bp DNA fragments. Separation relies on the differences in electrophoretic mobilities of negatively-charged DNA fragments, which due to denaturation by heat and urea, correspond to the sizes of the DNA fragments.

Since DNA fragments here will be visualized with a fluorescent dye (e.g., FAM or Cy3), it is important to prepare the sequencing gel using plates that are made of low-fluorescence glass. To reduce the background signal, it is also crucial to be extremely thorough with cleaning the gel plates (described below). Before attempting to use the sequencing gel to resolve DNA fragments extracted from OP-mediated DNA cleavage reactions (Basic Protocol 3), we recommend that novice users first practice preparing and running a sequencing gel with only the dideoxy sequencing ladders (from Support Protocol 1). For analysis of OP-cleaved DNA fragments, samples need to be prepared in advance in the form of dry pellets (see Basic Protocol 3) and resuspended in small-fragment formamide loading buffer (see Reagents and Solutions).

In many cases, it is convenient to polymerize sequencing gels overnight on the benchtop (~14 hours), and thus prepared at the end of day 1 of Basic Protocol 3. However, as little as six hours is sufficient for full polymerization, allowing days 1 and 2 to be merged into a single day, with preparation of the gel in the morning.

A critical factor in preparing the sequencing gel is preserving the temperature of the denaturing urea-acrylamide solution close to 4°C (above 0°C and below 10°C) at all times. This solution has high concentration of urea and thermal fluctuations can cause the urea to crystalize.

Materials

• 10% APS (see Reagents and Solutions), kept on ice

• Tetramethylethylenediamine (TEMED; Fisher, cat. no. BP15020)

• 70% (v/v) isopropanol

• High-vacuum grease (e.g., Dow Corning)

• Ultrapure water (e.g., Milli-Q water)

• 70% (v/v) ethanol in water

• 10% (v/v) acetic acid in water

• Deionized water (DI)

• 10X TBE (see Reagents and Solutions)

• Denaturing sequencing gel solution (see Reagents and Solutions), at 4 °C

  CAUTION: Acrylamide (within the denaturing sequencing gel solution) is neurotoxic and suspected to be carcinogenic. When working with acrylamide solutions, wear gloves, eye protection, and a protective lab coat.

• Dideoxy sequencing ladders (Support Protocol 1)

• DNA samples, site-specifically cleaved with OP (Basic Protocol 3)

• Paper towels or diaper pads

• Sequencing gel electrophoresis system (e.g., LabRepCo S2 apparatus with Surface temperature monitor, cat. no. 21105036; LabRepCo silicon spacer foam blocks, cat. no. 11678018; gel company Li-Cor 0.4 mm well forming comb, cat. no. CLW50-040 (50-well) or Thomas Scientific 0.4 mm well comb, cat. no. 4266L33 (40-well); Expedeon custom-made low-fluorescence long glass plate (4 mm × 33 cm × 42 cm); Expedeon custom-made low-fluorescence short glass plate (4 mm × 33 cm × 39.3 cm); gel company customized length (42 cm) Li-COR 0.4 mm mylar spacers)

• Kimwipes

• Gel sealing tape (preferably with ≥ 3.5 cm width, e.g., VWR, cat. no. MSPP-GT-72-15)

• Extra-large binder clips (e.g. Office Depot, cat. no. 308957)

• 100-mL glass beaker

• 100-mL graduated cylinder

• Bubble level (for pouring acrylamide gels)

• 60-ml plastic syringe with needle

• (Multicolor)fluorescence scanner (e.g., Typhoon 5 variable mode imager with multistage scanning tray, cat. no. 29187198 and 33 cm × 42 cm glass plate guides, cat. no. 29215514)

Protocol steps

Preparing for pouring the denaturant DNA sequencing gel

• 1Chill TEMED by placing the bottle on ice.
• 2
  Prepare the glass plates of the sequencing gel. First, place each gel plate gently on a clean padded surface (e.g. paper towels or an absorbent ‘diaper’ pad). Then, using Kimwipes, clean each plate using the following procedure:

  1. Remove all grease from both sides of the glass plate (from previous usage) using 70% isopropanol. Dry with Kimwipes.
  2. Wash both sides of the plate twice with ultrapure water, then dry with Kimwipes.
  3. Wash the inside surface (that will contact gel) with 70% EtOH. Dry with Kimwipes.
  4. Wash the inside surface twice with ultrapure water. Dry with Kimwipes.
  5. Wipe the inside surface with ~1 mL of 10% acetic acid. Dry with Kimwipes.
  6. Wash the inside surface twice with ultrapure water. Dry with Kimwipes.
  7. Using a Kimwipe dampened with ultrapure water, remove lint and dust particles from the interior surface of the plate.
     We recommend designating one side of each plate as the interior (the side contacting the gel) and the other side as the exterior (the outside surface of the glass gel sandwich). Before pouring the gel, it is more important to thoroughly clean the interior side; the exterior should be similarly cleaned just before scanning the gel.
     Although laborious, we find that a rigorous cleaning is essential for obtaining clean scans of the gel, as the fluorescence scanner is extremely sensitive. A thorough cleaning of both plates will take ~30 minutes. Avoid using paper towels to wipe the plates during the gel cleaning process (before pouring the sequencing gel solution, after gel polymerization and before scanning the gel). Rubbing the plates with dry paper towels creates static electricity that can interfere with electrophoresis.
• 3Assemble the two glass plates with 0.4 mm side spacers using sealing tape and binder clips (Figure 4). The two glass plates taped together is called the gel sandwich.

  Position the binder clips symmetrically to ensure even pressure on the gel.

Figure 4. Stepwise assembly of a denaturing DNA sequencing gel.

(a) With the long sequencing plate on paper towels, place the 0.4-mm side spacers along the inside long edges of the sequencing plate, with sealing foam blocks at the top of the long plate. (b) Keeping the spacers in place, carefully lay the short plate on top of the long plate, lining up the bottom corners. (c) Next, tape the two plates together. With the bottom edge of the plate sandwich hanging off the bench, gently apply sealing tape that wraps over both bottom corners. With gloves, smooth the tape along the plate edges, then lay the overhanging tape along the top and bottom of the gel sandwich. (d) Repeat the process for taping up the sides of the gel sandwich. (e) To the bottom of the sandwich, apply another piece of sealing tape that overhangs each corner by ~5–8 cm. (f) Place an additional piece of sealing tape at the top of the gel sandwich, by the foam blocks. (g) Using 8 extra-large binder clips, clamp the edges of the glass plates onto the spacers. Do not place binder clips on glass plates where no spacer is present, as it will result in an uneven gel and distorted image. (h) Create a small incline by resting the glass plates on a binder clip wrapped in a paper towel. (i) Pour the denaturing gel solution as described in Support Protocol 2, inserting the comb and laying nearly flat on the wrapped binder clip for polymerization. (j) After the gel is polymerized, remove the tape, wipe the gel sandwich and assemble it on an S2 apparatus sealed with high-vacuum grease along the silicon gasket. Secure the clamps against the gel sandwich by firmly tightening all screw knobs.

Pouring the denaturing sequencing gel

• 4Place a 100-mL beaker on ice to chill for ≤ 5 minutes. Meanwhile, have the 0.4 mm 40-well comb ready near the gel setup.
• 5Remove the 100-mL pre-chilled beaker from ice and place it on the benchtop. Immediately, using a 100-mL graduated cylinder, pour 80 mL of the denaturing sequencing solution (previously at 4 °C) into the beaker. Immediately proceed to the next step to minimize warming up the solution.

  Placing the denaturing sequencing gel solution on ice increases the possibility of urea crystallization.

• 6Using a P1000, add 290 μL of pre-chilled 10% APS solution and gently but quickly swirl the solution. Using a P200 pipette, immediately add 29 μL of TEMED and similarly swirl to mix. Avoid making bubbles.

  The rate of gel polymerization is affected by the amounts of APS and TEMED, and the ambient temperature of the room. Higher amounts of APS and TEMED and higher temperatures accelerate polymerization.

  If the gel polymerizes while pouring, disassemble the gel sandwich and reclean the interior sides of the plates before reassembling the gel sandwich.

• 7Immediately pour the solution inside the sandwiched gel plates by holding the plates with one hand at ~50–70° relative to the benchtop, while continuously pouring the solution with the other hand.

  If the urea starts to crystallize (to a very minor extent; very few observable crystals) while pouring the gel, gently rock the gel sandwich back and forth to dissolve the crystals. Keep the gel solution from overflowing while rocking.

• 8Pour approximately half of the solution into the gel sandwich, then hold the sandwich nearly perpendicular relative to the benchtop and inspect for bubbles. If there are any bubbles, vigorously tap the surface of the short plate while rotating the plates to dislodge the bubble.

  While tapping the plates, do not disturb the binder clips. If the clips are displaced, adjust them immediately after pouring the gel by gently changing their position without taking them off completely

  The ideal location to tap the plates is under the bubble.

  While removing bubbles (if any), place the beaker containing the denaturing sequencing solution away from the gel sandwich to avoid accidently spilling it while rearranging the plates.

• 9Continue pouring the gel. When ~10 mL of the solution is left in the beaker and the denaturing sequencing gel starts to overflow from the gel sandwich, inspect the gel for any additional bubbles. Remove any bubbles by tapping, as described before. Then, while holding the gel sandwich vertically with one hand, use the other hand to insert a 0.4 mm 40-well comb between the two plates at the top of the gel (Figure 4). Insert fully so that the left and right corners of the comb rest on top of the short plate.
• 10Gently lean down the prepared gel (without moving the comb) so that it is almost parallel with the bench top. Pour the left-over (~10 mL) sequencing solution into the area between the comb teeth and short plate.

  When placing the gel (almost) parallel to benchtop surface, the solution will start spilling over from inside the sandwich. Using a bubble level, adjust the incline such that both the left and the right side of the gel are even. Place folded paper towels under the binder clips on the appropriate site to bring it to the same level as the opposite side.

  Do not disturb the gel for at least 6 hours after pouring.

Running the denaturing sequencing gel

• 11The next day (or at least after 6 hours), hold the gel vertically and gently remove each clip, alternating on each side until all clips are removed. Lay the gel on the benchtop and remove the gel-sealing tape.
• 12Symmetric removal of the binder clips from each side of the gel sandwich prevents uneven pressure on the gel.
• 13Holding the gel in a vertical position, use ultrapure water to clean off the surface of the long plate; then, dry with Kimwipes. Lay the gel sandwich on the benchtop on clean paper towels.
• 14While the gel is resting on the benchtop, use ultrapure water to rinse the short plate, and remove all overflowed gel and urea crystals. Do not apply pressure to the gel while wiping it.
• 15Pour ultrapure water on the comb teeth while the comb is still inserted. This makes it easier to remove the comb without disturbing the wells. After a few minutes, gently take out the comb by holding the comb at ~5 cm from each end with both hands and carefully pulling it out in a way that teeth displacement from the gel is equal on both ends of the comb.

  After removing the comb, if there are any polymerized gel fragments blocking the wells, use a P200 pipette tip to gently remove them.

• 16Using ultrapure water and Kimwipes, clean the surface of the long plate exposed after comb removal. Do not allow the Kimwipe to touch the edge of the wells.
• 17Rinse the surface of the short plate with ultrapure water and completely dry with Kimwipes. Do not apply pressure while cleaning.
• 18In a 1-L graduated cylinder, prepare 1 L of 1X TBE using autoclaved 10X TBE. Keep at RT.
• 19Set up the S2 sequencing gel electrophoresis apparatus (Figure 4) by generously applying high-vacuum grease along the entire surface of the silicon gasket. Make sure the grease is covering any openings between the gasket and the aluminum plate as well as the space between the gasket and the upper buffer chamber on the S2 apparatus. Secure the removable buffer tray.

  While preparing the S2 apparatus, inspect the silicon gasket for damages and replace if any tearing is observed. After preparing the S2 apparatus, change gloves, to avoid getting grease on the glass plates.

• 20Place the gel sandwich on the gel supports inside the empty removable buffer tray, with the short plate facing toward the aluminum plate, and carefully lean it against the silicon gasket such that the foam blocks on the side spacers and the ends of the silicon gasket form a sealed space on top of the buffer chamber.
• 21Position the S2 gel clamps (bottom and top on both sides of the S2 apparatus) on the surface of the long plate. Secure by firmly tightening all four screw knobs.

  To prevent irregularities in the gel sandwiched between the two plates, increase the tightening gradually and by alternating between different knobs. Do not overtighten the screw knobs, as it can break the clamps.

  The silicon gasket must not wrinkle while tightening the clamps, but should be pressed firmly and evenly against the short plate and sealing foam blocks.

• 22Using a P200 pipette tip, press the high-vacuum grease against the short plate to seal the space between the short plate and silicon gasket.

  Do not allow the high-vacuum grease to reach the edge of the short plate and enter the wells.

• 23Close the upper buffer chamber drain valve (on the right side of the S2 apparatus). Then pour ~500 mL of 1X TBE (prepared in step 17) into the upper buffer chamber and the rest of the solution into the removable buffer tray at the bottom. The level of the solution in the bottom tray must be above the bottom edge of the gel sandwich.

  The solution on the upper buffer chamber must be ~2 cm above the edge of the short plate and enter the wells completely without any bubbles inside them.

• 24Inspect the upper buffer chamber for leakage.
• 25Draw some 1X TBE from the upper chamber into a 60-mL plastic syringe (needle attached) and use it to gently flush urea from the wells. Urea has a visible difference in viscosity compared with 1X TBE. Care must be taken to avoid damaging the lane walls while flushing.
• 26Close the safety lids (top and bottom) and attach the power cords. Pre-run the gel for at least 45 to 60 minutes at 65 W (constant watts), with the goal of increasing the gel temperature to approximately 50°C (can be checked by a thermometer strip, attached to the outside of the short plate).

  During this step, rinse the comb with DI water and allow it to dry on clean paper towels.

• 27During the pre-run, remove the DNA samples resuspended in small-fragment formamide loading dye (Basic Protocol 3) from −20°C. Keep at RT.
• 28Prepare the sequencing ladder for loading on the gel. First, take out the eight microcentrifuge tubes continuing the dideoxy ladders prepared in Support Protocol 1 (four tubes for each primer). Label four fresh microcentrifuge tubes as A, C, G, and T. In A, combine the ddATP reaction run with the forward primer with the ddATP reaction run with the reverse primer to create the ddATP ladder. Do the same with C, G, and T, respectively, combining the corresponding ddNTP reactions for the two different primers together into the appropriate tubes. For instance, the tube labeled “A” would contain both FAM- and Cy3-labeled didoxy ladders generated via Sanger sequencing in the presence of ddATP. If necessary, pulse-spin the four resulting tubes containing the dideoxy ladders to collect the solutions at the bottom of the tubes. Keep at RT.

  When scanning at two different wavelengths, merging the Sanger reactions with the two different primers according to the ddNTP will save an extra four lanes, thus allowing more lanes for DNA cleavage reactions.

• 29Set a heat block to ~95 °C. Check the water level frequently and fill as needed.
• 30Once the gel pre-run is finished, pause the power supply, disconnect the power cord, open the top safety lid, and flush the wells again as described in step 24.
• 31Using a P10 pipette, immediately load 6.5 μl of each of the four dideoxy ladders mixtures (from Step 27) in four separate wells. While loading the ladders, place two of the 1.5-mL DNA fragment tubes (step 26) inside the 95 °C heat block to warm up for ~1 minute before loading on the gel.

  Flat 0.4 mm pipette tips are not required, but can be used to load the gel.

  Do not heat the dideoxy ladders.

• 32Next, load ~4.3 μL from tubes containing the processed DNA fragments (incubated in the heat block for ~1 minute in step 30) on the sequencing gel. For each microcentrifuge tube taken from the heat block, replace it with another, allowing each new tube to reach to 95°C prior to loading.

  The volume of the solution containing the DNA sample is larger than the amount of small-fragment formamide loading dye added to resuspend the DNA pellet (4 μL) due to expansion of the solution by heating at 95 °C.

  Adjust the number of microcentrifuge tubes placed on the heat block at any one time according to the speed of gel loading. Each DNA fragment sample should be heated for ~1 minute prior to loading on the gel. Do not allow DNA fragments to cool down before loading.

  The amount of labeled DNA per lane should be less than 2 pmol. Overshooting this amount can result in smeary bands.

• 33After loading the gel, immediately close the safety lid, reconnect the electric cord to the top of the S2 apparatus, and continue the run at 65 W.

  The final loaded gel should contain four wells corresponding to four dideoxy ladder Sanger sequencing mixtures (loaded in step 30) with each subsequent well containing a DNA cleavage reaction from 0.8 pmol OP-labeled nucleosomes (step 31).

  While running, regularly check the power supply to ensure the run is not stopped.

• 34Continue the run until the orange dye used in the small-fragment formamide loading dye runs off the bottom of the gel, which takes ~1.5 hours.
• 35Turn off the power supply and disconnect the power cords. Open the safety lids and, using the upper buffer chamber drain valve (on the right side of the S2 apparatus), completely drain the 1X TBE solution from the upper chamber.
• 36Place four clean paper towels (as in similar to step 4, see Figure 4) on a clean benchtop.
• 37Loosen the S2 gel clamps and carefully remove the gel sandwich from the S2 apparatus. Avoid separating the short and long plates while taking out the gel sandwich.
• 38Place the gel on the long plate facing downward on the paper towels and thoroughly wipe the left-over grease from the short plate, the exposed long plate, and the edge of the gel sandwich. Use 70% (v/v) isopropanol and Kimwipes for this purpose. Dry with Kimwipes once all the high-vacuum grease is removed.

  Change gloves as needed, to prevent transferring the high-vacuum grease from gloves to the gel sandwich.

• 39Thoroughly clean both sides of the gel sandwich, using the protocol described in step 2 above.

  Proper cleaning of the gel sandwich should be very thorough. It can take as long as 30 minutes to complete.

• 40Before scanning the gel, wipe the short and long plate one more time with ultrapure water. Avoid leaving behind lint before scanning the gel.
• 41Scan the gel using a fluorescence scanner such as a Typhoon 5 variable mode imager at excitation wavelengths corresponding to the two fluorophores used to label the DNA (e.g., FAM and Cy3).

  Be sure to use scanner settings where band intensities are not beyond the dynamic range of the detector (typically, saturated signals are highlighted in red by the scanning software). With saturating signals, it may be necessary to reduce sensitivity of the detector, for example, by reducing voltage of the photomultiplier tube (PMT). We typically scan the FAM-labeled DNA fragments extracted from 0.8 pmol OP-labeled nucleosome samples (in Basic Protocol 3) at PMT voltage of 550 v. For scanning with excitation by the Cy3 laser, PMT voltage is often set to 600 v. Sample data are shown in Figure 5.

• 42After scanning is completed, separate the two plates, discard the gel sandwiched between the plates in a proper waste container, and rinse the plates (both sides) with DI water. Check the plates and spacers. If there is any residual grease left, use 70% (v/v) isopropanol to wipe it off. Place the plates on clean paper towels and dry with Kimwipes or air-dry. Store the plates, 0.4-mm spacers, and comb away from dust once they dry.
• 43Empty the 1X TBE solution collected at the removable bottom tray of S2 apparatus and rinse the tray with DI water.

Figure 5. Example sequencing gels of OP-guided DNA cleavages on the nucleosome.

(a) Schematic of H2B(T87C) sites labeled with OP on the nucleosome, and how cleavage sites shift with nucleosome sliding. (b,c) Urea denaturing acrylamide gels showing DNA cleavage products for a nucleosome sliding reaction, using a histone core OP-labeled on H2B(T87C). Nucleosomes (40-601-40) were repositioned by 200 mM Chd1, with time points taken 7 s, 15 s, 30 s, 1 min, 2 min, and 4 min after addition of a sub-saturating amount (50 μM) of ATP (see Basic Protocol 2). Since the DNA was fluorescently labeled on both 5’ ends, the two scans (b, FAM; c, Cy3) report on DNA cleavages for each strand. Each OP-labeled H2B(T87C) site cleaves both DNA strands, allowing histone-DNA contacts to be monitored on both sides of the 601 sequence. Numbers on the right of each gel indicate the cut sites relative to the Widom 601 dyad (position zero). The top bands correspond to the uncleaved, full-length DNA fragment, illustrating the efficiency of OP-labeling and cleavage. (d) Mapping cleavage sites near the nucleosome dyad using Widom 601 nucleosomes labeled with OP at H4(S47C). The dideoxy sequencing ladders are shown by lanes labeled A, C, G, T. The lightning bolts in the nucleosome cartoons represent DNA cleavage events.

REAGENTS AND SOLUTIONS:

Acrylamide:bis-acrylamide (60:1) solution, 30% (w/v)

Using a 250-mL glass beaker, magnetic stir bar and stir plate, and a 250-mL graduated cylinder, dissolve 60 g acrylamide (Bio-Rad, cat. no. 161-0101) and 1 g bis-acrylamide (Bio-Rad, cat. no. 161-0201) in ultrapure water (e.g., Milli-Q) to final volume of 200 mL.

Filter (0.22 μm) the solution and vacuum-degas for at least 30 minutes.

Wrap the solution with aluminum foil and store at 4°C for ≤ 6 months.

CAUTION: Acrylamide is neurotoxic and suspected to be carcinogenic. When working with acrylamide solutions, wear gloves, eye protection and a protective lab coat. While handling dry acrylamide, wear a vapor-protective mask and face shield to avoid inhalation exposure.

TBE, 10X

• 890 mM Trizma base (Sigma-Aldrich, cat. no. T1503)

• 890 mM Boric acid (Fisher, cat. no. A73-500)

• 20 mM EDTA·KOH (pH 8.0) (EDTA: Sigma-Aldrich, cat. no. E9884; KOH: Sigma-Aldrich, cat. no. P5958)

Dissolve 108 g of Trizma base and 55 g of boric acid in ~800 mL of ultrapure water (e.g., Milli-Q). Once dissolved, add 40 mL of 0.5 M EDTA·KOH (pH 8.0) and bring the volume to 1 L using ultrapure water.

Autoclave the solution and store for several months at RT. Discard the solution if precipitation occurs.

TBE, 20X

• 1780 mM Trizma base (Sigma-Aldrich, cat. no. T1503)

• 1780 mM Boric acid (Fisher, cat. no. A73-500)

• 40 mM EDTA·KOH (pH 8.0) (EDTA: Sigma-Aldrich, cat. no. E9884; KOH: Sigma-Aldrich, cat. no. P5958)

In a 50-mL conical tube, add 8.6 g Trizma base (Sigma-Aldrich, cat. no. T1503), 4.4 g boric acid (Fisher, cat. no. A73-500), and 3.2 mL of 0.5 M EDTA·KOH (pH 8.0) filtered (0.22 μm) solution to ultrapure water (e.g., Milli-Q) to final volume of 40 mL.

Vortex vigorously to dissolve completely.

Store the 20X TBE solution at RT for several months and discard when precipitation occurs.

APS solution, 10% (w/v)

Weigh out 100 mg Ammonium persulfate (APS; Bio-Rad, cat. no. 161-0700) directly inside a 1.5-mL microfuge tube. Add 1000 μL ultrapure water (e.g., Milli-Q) to have a 10% (w/v) solution. Vortex to dissolve.

Make fresh before polymerizing gel solutions.

ATP stock solution, 500 mM

• ATP disodium salt (Sigma-Aldrich, cat. no. A2383)

• 10 M NaOH (see recipe)

• pH test strips (e.g., J.T. Baker BAKER-pHIX, cat. no. 4405-01, pH range 6.0–8.1, 0.3 pH Graduations).

Make 2 mL of a 100 mM HEPES·KOH solution by adding 200 μL of 1 M HEPES·KOH, pH 7.5 to 1.8 mL of ultrapure water. In a separate 2-mL microfuge tube, dissolve 551 mg of ATP disodium salt in ~1.5 mL of the 100 mM HEPES·KOH, pH 7.5 solution. Add ~170 μL of 10 M NaOH and check pH with pH test strips. Continue adding 10 M NaOH until pH 7.5 is reached. Then, bring up volume to 2 mL total using 100 mM HEPES·KOH, pH 7.5.

Make 10 μL aliquots and store at −80 °C.

CAUTION: 10 M NaOH is caustic. Wear personal protective equipment while handling NaOH.

Cleavage buffer, 1 mL

• 50 mM Tris·HCl, pH 7.5 (Trizma base: Sigma-Aldrich, cat. no. T1503)

• 2.5 mM NaCl (Sigma-Aldrich, cat. no. S9888)

• Ultrapure water (e.g., Milli-Q) to final volume of 1.0 mL.

Make fresh and keep at RT.

CuCl₂, 75 mM

Prepare a 3 M stock solution of CuCl₂ by dissolving 2.56 g CuCl₂ (dihydrate; Sigma-Aldrich cat. no. C3279) in ultrapure water (e.g., Milli-Q) to final volume of 5.0 mL. Vortex vigorously, filter (0.22 μm) and store at RT indefinitely.

Combine 250 μL of 3 M stock solution of CuCl₂ with 9.75 mL ultrapure water (e.g., Milli-Q) to make 10.0 mL 75 mM stock solution of CuCl₂.

Vortex to mix and store at RT for ≤ 1 year.

If CuCl₂ is not available, alternatively use CuSO₄ (Sigma-Aldrich, cat. no. C8027).

D-(+)-Glucose, 40% (w/v) (~2.2 M)

Dissolve 40 g of D-(+)-Glucose (Sigma-Aldrich, cat. no. G7021) in ultrapure water (e.g., Milli-Q) in a total volume of 100 mL. Filter the solution (0.22 μm) and store at 4 °C for ≤ 1 year. Discard if glucose is precipitated out or the solution is cloudy.

Denaturing sequencing gel solution

• 7.8 M Urea (Sigma-Aldrich, cat. no. U1250)

• 8% (w/v) acrylamide:bis-acrylamide solution, 19:1 (using 40% (w/v) solution: BioRad cat. no. 1610144)

• 1X TBE

In a 500-mL beaker containing a magnetic stir bar, add 48 mL of 40% 19:1 acrylamide:bis-acrylamide (BioRad cat. no. 1610144) and 24 mL of autoclaved 10X TBE. While mixing the solution, add 112 g of urea. Increase the solution volume to ~220 mL by addition of ultrapure water. Place a wide-opening plastic container on a stir plate. Add enough warm water(~40 °C; not hot) to make a water bath, then place the beaker with the urea-containing solution in the water bath and continue mixing until urea is completely dissolved. Transfer the solution to a 250-mL graduated cylinder and add ultrapure water to a final volume of 240 mL. Filter (0.22 μm) and vacuum-degas for at least 30 minutes. Transfer into a 500-mL glass bottle covered in aluminum foil. Store at 4 °C for ≤ 6 weeks.

CAUTION: Acrylamide is neurotoxic and suspected to be carcinogenic. When working with acrylamide solutions, wear gloves, eye protection and a protective lab coat.

Glycogen, 20 mg·mL⁻¹

Weigh 20 mg of glycogen (Sigma-Aldrich, cat. no. G8751) directly inside a 1.5-mL microcentrifuge tube. Add 1.0 mL ultrapure water (e.g., Milli-Q) and vortex to dissolve. Store solution at −20 °C indefinitely.

HEPES·KOH, pH 7.5, 1 M

• HEPES (Sigma-Aldrich, cat. no. H3375)

• KOH (Sigma-Aldrich, cat. no. P5958).

• pH meter (e.g., Mettler Toledo SevenGo Duo model SG23 pH meter with InLab routine Pro pH electrode)

Make a 10 M KOH solution by dissolving 5.61 g of KOH in ~5 mL of ultrapure water, then adding more water to bring the total volume to 10 ml.

Make 50 ml of a 1 M HEPES·KOH, pH 7.5 solution by dissolving 11.92 g of HEPES into ~35 ml of ultrapure water. Adjust pH to 7.5 using 10 M KOH solution. After pH 7.5 is reached, bring the total volume to 50 ml with ultrapure water, and then 0.22 μm filter and store at 4 °C.

Hexokinase, 1.0 u·μL⁻¹

• 1 KU bottle of hexokinase (Sigma-Aldrich, cat. no. H4502, ≥ 130 u·mg⁻¹ protein).

• 10 mM HEPES·KOH, pH 7.5

Using the information provided on the purchased hexokinase bottle, determine the activity of the entire solid (lyophilized protein powder). For instance, if the label states that there is 2.74 mg of solid inside the bottle and the solid has an activity of 364 u·mg⁻¹ of solid, the activity associated with the entire solid is 2.74 mg × 364 u·mg⁻¹ ≃ 997 units.

Make 2 mL of 10 mM HEPES·KOH, pH 7.5 solution by adding 20 μL of 1 M HEPES·KOH to 1.98 mL of ultrapure water. Then, add a sufficient amount of 10 mM HEPES·KOH, pH 7.5 to the entire lyophilized hexokinase bottle in order to have a 1.0 u·μL⁻¹ hexokinase solution. In the example above, addition of ≤ 997 μL buffer to entire bottle (2.74 mg solid) results in enzymatic activities ≥ 1.0 u·μL⁻¹.

Since the activity of hexokinase in quenching buffer has to be ≥ 0.04 u·μL⁻¹, make sure that the activity of hexokinase stock solution here is ≥1.0 u·μL⁻¹.

Do not vortex to dissolve, rather gently invert the bottle. Place the prepared solution on ice until ready to aliquot.

Aliquot 10–20 μL of 1.0 u·μL⁻¹ hexokinase stock solution in 0.6-mL or 1.5-mL microcentrifuge tubes. Flash-freeze using liquid nitrogen and store at −80 °C indefinitely.

KOH, 1 M

In a 100-mL graduated cylinder, dissolve 5.61 g KOH (Sigma-Aldrich, cat. no. P5958) in ultrapure water (e.g., Milli-Q), and bring the final volume to 100 mL. Filter the solution (0.22 μm); store at RT indefinitely.

Labeling buffer, 10 mL

• 10 mM Tris·HCl, pH 7.5 (Trizma base: Sigma-Aldrich, cat. no. T1503)

• 1 mM EDTA·KOH (pH 8.0) (EDTA: Sigma-Aldrich, cat. no. E9884; KOH: Sigma-Aldrich, cat. no. P5958)

• 2 M NaCl (Sigma-Aldrich, cat. no. S9888)

• 10% (v/v) Glycerol (Sigma-Aldrich, cat. no. G7893)

• Add ultrapure water (e.g., Milli-Q) to 10 mL in a 15-mL conical tube.

Vortex to mix.

Prepare fresh before the start of labeling reaction and keep on ice.

Mapping buffer, 2X

• 100 mM Tris·HCl, pH 7.5 (Trizma base: Sigma-Aldrich, cat. no. T1503)

• 5 mM NaCl (Sigma-Aldrich, cat. no. S9888)

• 300 μM CuCl₂ (Sigma-Aldrich cat. no. C3279)

Use 4.0 μL of 75 mM CuCl₂ solution as the CuCl₂ stock solution to prepare a 1.0 mL 2X mapping buffer. Use ultrapure water (e.g., Milli-Q) to bring final volume of 1.0 mL.

Make fresh and keep at RT.

NaOAC/HOAC, pH 5.2, 3 M

• 26.6 g sodium acetate (Sigma-Aldrich, cat. no. 241245)

• Glacial acetic acid (Fisher, cat. no. A38-212)

• pH-meter (e.g., Mettler Toledo SevenGo Duo model SG23 pH meter with InLab routine Pro pH electrode)

In a 250-mL beaker with a magnetic stir bar, dissolve 26.6 g of sodium acetate in ~80 mL of ultrapure water (e.g., Milli-Q). Using a pH-meter, check the pH of the solution and adjust it to pH 5.2 by dropwise addition of concentrated acetic acid. Transfer the solution to a 100-mL graduated cylinder and bring to a final volume of 100 mL with ultrapure water. Filter (0.22 μm) and store indefinitely at RT.

CAUTION: concentrated acetic acid is extremely corrosive. Wear protective gloves, a lab coat and eye protection while handling it. Work under a well-ventilated hood and avoid inhaling the fumes.

NaOH, 10 M

In a 50-mL conical tube, dissolve 16 g NaOH pellets (Fisher, cat. no. BP359) by vortexing in ultrapure water (e.g., Milli-Q) to final volume of 40 mL. Filter the solution (0.22 μm) and store at RT indefinitely.

Do not store NaOH solutions in glass containers.

CAUTION: 10 M NaOH is caustic. Wear personal protective equipment while handling NaOH.

Neocuproine, 25 mM

In a 15-mL conical tube, add 52.1 mg Neocuproine (Sigma-Aldrich, cat. no. N1501) to 10.0 mL Dimethyl Sulfoxide (DMSO; Fisher, cat. no. D128-500). Dissolve by vortexing.

Neocuproine is light sensitive; thus, wrap the solid neocuproine bottle and the 15-mL conical tube in aluminum foil. Store the purchased solid neocuproine indefinitely at RT. The 25 mM solution can be stored at RT for ≤ 1 year. Do not store the solutions containing DMSO at temperatures ≤ 18 °C.

OP solution, 4.5 mM

• N-(1,10-Phenanthrolin-5-yl)iodoacetamide (Iodoacetyl OP; Biotium, cat. no. 92015)

• Dimethyl sulfoxide (DMSO; Fisher, cat. no. D128-500)

To prepare the OP solution, add 3060 μL of DMSO to the entire 5 mg bottle of Iodoacetyl OP. Vigorously mix the bottle to dissolve the product completely. Re-wrap the bottle in aluminum foil.

Store the OP solution at room temperature for ≤ 6 months in a dark and dry environment.

Do not store the solutions containing DMSO in temperatures ≤ 18 °C.

Do not unseal the purchased Iodoacetyl OP bottle until ready to make this solution.

Iodoacetyl is light-sensitive. Wrap the unsealed bottle in aluminum foil. Store it with a desiccant at −20 °C. When preparing this solution, work in a dark room with lamp that has a low wattage (e.g., 25 W) incandescent light bulb. Minimize exposure to ambient light.

Purified Chd1 remodeler

Express and purify a truncated construct of the Saccharomyces cerevisiae Chd1 protein (residues 118-1274) according to Nodelman et al., 2017.

Determine the protein concentrations using a spectrophotometer. The extinction coefficient of this construct at 280 nm is 137350 M⁻¹·cm⁻¹.

Aliquot the samples in 0.6-mL or 1.5-mL microcentrifuge tubes at final concentrations ≥ 50 μM in 30 mM Tris·HCl, pH 7.5, 150 mM NaCl, 10% (v/v) glycerol, and 1 mM dithiothreitol (DTT) storage buffer (Trizma base: Sigma-Aldrich, cat. no. T1503; HCl: Fisher, cat. no. A144S-212; NaCl: Sigma-Aldrich, cat. no. S9888; glycerol: Sigma-Aldrich, cat. no. G7893; DTT: Sigma-Aldrich, cat. no. D9163).

Snap-freeze in liquid nitrogen and store at −80°C indefinitely.

Purified folded histones (dimer, tetramer, octamer)

Using PCR-based mutagenesis, introduce a single-cysteine mutation on recombinant histone DNA (see Strategic Planning). Purify the generated plasmid and confirm the DNA sequence.

Express and purify recombinant histones as described previously (Dyer et al., 2004).

To obtain histone dimer, tetramer, or octamer, combine purified histones at equimolar ratios, refold and purify as described previously (Dyer et al., 2004).

Calculate the extinction coefficients of purified histone core proteins according to primary sequence, and determine protein concentrations using a spectrophotometer.

Aliquot the samples in 0.6-mL or 1.5-mL microcentrifuge tubes at final concentrations of 50 to 500 μM in storage buffer of 10 mM Tris·HCl, pH 7.5, 1 mM EDTA·KOH (pH 8.0), 2 M NaCl, 10% (v/v) glycerol, and 5 mM βME (Trizma base: Sigma-Aldrich, cat. no. T1503; HCl: Fisher, cat. no. A144S-212; EDTA: Sigma-Aldrich, cat. no. E9884; KOH: Sigma-Aldrich, cat. no. P5958; NaCl: Sigma-Aldrich, cat. no. S9888; glycerol: Sigma-Aldrich, cat. no. G7893; βME; Sigma-Aldrich, cat. no. M6250).

Flash-freeze in liquid nitrogen and store at −80 °C indefinitely.

CAUTION: concentrated HCl is extremely corrosive. Wear protective gloves, a lab coat and eye protection while handling HCl. Work under a well-ventilated hood and avoid inhaling the fumes.

Purified nucleosomes

Perform large-scale DNA amplification similar to (Nodelman et al., 2020) using fluorescently-labeled DNA oligos (see Strategic Planning). If possible, using FAM as the fluorophore is highly recommended.

Purify the DNA as described in (Nodelman et al., 2020) and use it to reconstitute the desired nucleosome, as in (Luger, Rechsteiner, & Richmond, 1999; Nodelman et al., 2020).

Purify the nucleosome using native PAGE, as described in (Nodelman et al., 2020).

Aliquot 5–10 μL of 2–10 μM sample in 0.6- or 1.5-mL prechilled microcentrifuge tubes.

Flash-freeze in liquid nitrogen and store at −80°C indefinitely.

Post-cleavage buffer

50 mM Tris·HCl, pH 8.0 (Trizma base: Sigma-Aldrich, cat. no. T1503)

2% (w/v) sodium dodecyl sulfate (SDS; Sigma-Aldrich, cat. no. L6026)

100 mM NaCl (Sigma-Aldrich, cat. no. S9888)

Add Ultrapure water (e.g., Milli-Q) to final volume of 40 mL in 50-mL conical tube.

Store at RT for several months.

Post-labeling dialysis buffer (2 × 2 L = 4 L total)

• 10 mM Tris·HCl, pH 7.5 (Trizma base: Sigma-Aldrich, cat. no. T1503)

• 1 mM EDTA·KOH (pH 8.0) (EDTA: Sigma-Aldrich, cat. no. E9884; KOH: Sigma-Aldrich, cat. no. P5958)

• 2 M NaCl (Sigma-Aldrich, cat. no. S9888)

• 10% (v/v) Glycerol (Sigma-Aldrich, cat. no. G7893)

• 5 mM 2-Mercaptoethanol (βME; Sigma-Aldrich, cat. no. M6250, ≥99.0% purity, 14.3 M).

• Add ultrapure water (e.g., Milli-Q) to 2 L (2X) or 4 L.

Prepare fresh on day 1 of histone labeling (without βME). Cover each 2-L beaker with parafilm and store at 4 °C (e.g., cold room) with the stir bar.

Add 700 μL of 14.3 M βME to each 2 L buffer immediately before dialysis.

Quenching buffer, 800 μL

• 50 mM Tris·HCl, pH 7.5 (Trizma base: Sigma-Aldrich, cat. no. T1503)

• 50 mM KCl (Sigma-Aldrich, cat. no. P3911)

• 10 mM MgCl₂ (Sigma-Aldrich, cat. no. M9272)

• 0.1 mg·mL⁻¹ Bovine serum albumin (BSA: Sigma-Aldrich, cat. no. A7906)

• 0.02 u·μL⁻¹ hexokinase

First, make quenching buffer without hexokinase, by adding all components and bringing total volume to 768 μL. Vortex and keep the solution on ice while ready to add hexokinase. Before the start of sliding reactions, thaw and add 32 μL of 1.0 u·μL⁻¹ hexokinase stock solution to pre-chilled quenching buffer. Mix gently by pipetting up and down. Do not vortex the quenching solution after hexokinase addition.

The left-over thawed 1.0 u·μL-1 hexokinase stock solution can be stored at −20 °C for ≤ 1 month, as it retains activity for a month after freezing and thawing. Avoid repetitive freezing and thawing.

Sliding buffer, 1.5 mL

• 50 mM Tris·HCl, pH 7.5 (Trizma base: Sigma-Aldrich, cat. no. T1503)

• 50 mM KCl (Sigma-Aldrich, cat. no. P3911)

• 10 mM MgCl₂ (Sigma-Aldrich, cat. no. M9272)

• 0.1 mg·mL⁻¹ Bovine serum albumin (BSA: Sigma-Aldrich, cat. no. A7906)

• 100 mM D-(+)-Glucose

Add components together in a 2-mL microcentrifuge tube. Use 68.2 μL of a 40% (w/v) D-(+)-glucose solution. Use ultrapure water (e.g., Milli-Q) to bring total volume to 1.5 mL. Vortex to mix thoroughly.

Prepare solution fresh and keep on ice.

Small-fragment formamide loading dye

• 20X TBE

• 95% (v/v) formamide (Sigma-Aldrich, cat. no. 221198)

• 2 mg·mL⁻¹ Orange G dye (Sigma-Aldrich, cat. no. O3756)

• 0.75 g AG 501-X8 ion-exchange resin (Bio-Rad, cat. no. 1437424)

First, add ion-exchange resin to ~15 mL of 100% formamide in a beaker with a stir bar. Mix for at least 1 hour to deionize formamide. Filter (0.22 μm) to remove the beads.

To 9.5 mL of deionized and filtered formamide, add 500 μL of 20X TBE buffer. Add 20 mg of Orange G dye and mix.

Store in 125 μL aliquots at −20 °C indefinitely.

CAUTION: Formamide is toxic, suspected to be carcinogenic and may damage the unborn child. When working with formamide, wear gloves, eye protection, face shield and a protective lab coat. Avoid inhalation by working under a ventilated hood and prevent skin contact.

TCEP, 500 mM

In a 15-mL conical tube, dissolve 1.43 g of TCEP·HCl (Tris(2-carboxyethyl)phosphine hydrochloride; Pierce, cat. no. PG82080) in ultrapure water (e.g., Milli-Q) to a total volume of 10.0 mL. Filter (0.22 μm) and aliquot 500 μL of 500 mM solution into 1.5-mL microcentrifuge tubes.

Store at −20 °C for several years. Do not subject aliquots to repetitive cycles of freezing and thawing.

TCEP (pH-balanced), 5 mM

Add 40 μL of a 1 M KOH solution to 200 μL of a 500 mM TCEP solution. Check the pH with pH test strips (e.g., J.T. Baker BAKER-pHIX, cat. no. 4405-01, pH range 6.0–8.1, 0.3 pH graduations). Adjust the pH to final value between 7.2 to 7.5 by stepwise 1 μL additions of 1 M KOH solution. Vortex the sample after each KOH-addition.

The initial pH of TCEP solution is ~2.5. After the initial 40 μL addition, raising the pH to 7.2 can be achieved in approximately five 1-μL additions (~5 μL) of 1 M KOH solution (total of ~45 μL 1 M KOH addition).

The pH-adjusted TCEP solution can be stored at −20 °C.

Calculate the new concentration of TCEP solution after pH adjustment using the following equation:

$$\text{Concentration of pH balanced TCEP solution }\left(\text{mM}\right)\text{= }\frac{\text{500 }\left(\text{mM}\right)\text{×200 }\left(\mu \text{L}\right)}{\text{200 }\left(\mu \text{L}\right)\text{+Volume of added 1 M KOH solution to adjust the pH }}$$

For example, if 45 μL of 1 M KOH solution (see recipe) was added to pH-balance 200 μL of 500 mM TCEP solution (see recipe), the concentration of pH-balanced TCEP solution would be ~408.2 mM.

Before starting the OP-labeling reactions (Basic Protocol 1), use the pH-balanced solution and labeling buffer to prepare a fresh stock of 5 mM TCEP (pH-balanced), pH 7.5 solution

COMMENTARY

Background Information

By blocking easy access to DNA, the precise positioning of nucleosomes can regulate many processes (Kornberg & Lorch, 1999; Li, Carey & Workman, 2007; Nocetti & Whitehouse, 2016). Nucleosome positioning, as well as contact points around the histone core, have long been studied using hydroxyl radicals (Brown & Fox, 1996; Schwanbeck, Xiao & Wu, 2004). Through a Fenton reaction, iron(II) EDTA is oxidized by H₂O₂ to generate highly reactive hydroxyl radicals. The iron can be returned to its initial oxidation state with a coupled ascorbic acid/dehydroascorbate redox reaction, allowing multiple cycles of hydroxyl radical production. When produced in bulk solution, hydroxyl radicals cleave exposed regions of the DNA duplex, and, on nucleosomes, protection from histone contacts produces a characteristic footprinting pattern with ~10 bp periodicity. Although useful in revealing all points of contact around the nucleosome, cleavage from bulk hydroxyl radicals requires uniquely positioned nucleosomes, as overlapping footprints from multiple placements of the histone core are challenging to interpret. Footprinting with hydroxyl radicals produced in bulk solution is also sensitive to glycerol and other sugars, which can, even at very low (0.5% (v/v)) concentrations, scavenge hydroxyl radicals before they can migrate from bulk solution to DNA (Tullius et al., 1987).

Using an iron(II) EDTA derivative that can be attached to cysteine residues, Richmond and colleagues targeted production of hydroxyl radicals to specific sites on the histone core (Flaus et al., 1996; Flaus & Richmond, 1999). They showed that such tethered iron(II) EDTA moieties produce site-specific cleavage patterns that reflect histone-DNA contacts at bp resolution. This site-specific creation of hydroxyl radicals is advantageous over classical footprinting, as it can cleanly reveal multiple distinct positions of the histone core on DNA. Due to its proximity to DNA, site-specific targeting of iron(II) EDTA also allows for DNA cleavage in the presence of glycerol and other scavengers that interfere with footprinting from bulk solution. Although this method offers single-bp resolution in determining where specific histone sites contact DNA, and, thus, the positioning of the histone core on DNA, its application is limited by the need to chemically synthesize the EDTA derivative used to label the cysteine residue. Additionally, since EDTA is a strong chelator of divalent cations, including Mg²⁺ (a key cofactor in ATP hydrolysis reactions), buffer conditions containing divalent cations can potentially interfere, as they can replace Fe²⁺ cations and inhibit the site-specific DNA cleavage reaction.

A similar method for site-directed hydroxyl-radical DNA cleavage uses a derivative of ortho-phenanthroline (OP) to chelate copper cations. In this case, hydroxyl radicals result from H₂O₂ oxidizing Cu⁺, with repeated rounds of hydroxyl radical production achieved through Cu⁺ regeneration by a thiol such as 3-mercaptopropionic acid (MPA) (Marshall et al., 1981; Brogaard et al., 2012; Pope et al., 1982; Sigman et al., 1991). This creates a cycle of one-electron redox reactions (Figure 3) in a manner similar to the iron-catalyzed Haber-Weiss cycle (Marshall et al., 1981; Haber & Weiss, 1934; Liochev & Fridovich, 2002). Using an iodoacetyl derivative, the OP moiety can be covalently linked to cysteine residues through alkylation of the sulfhydryl group. A significant advantage of OP-directed DNA cleavage over that of EDTA-mediated cleavage is that an iodoacetyl derivative of OP is commercially available. Additionally, OP has a low affinity for Mg²⁺, making the DNA cleavage reaction relatively insensitive to Mg²⁺-containing buffers. At high (>100 mM) concentrations, Mg²⁺ may replace Cu²⁺ and inhibit the cleavage reaction, as Mg²⁺ does not undergo a Fenton reaction to create reactive radicals (Sigman et al., 1991). However, even at such high concentrations, increasing the Cu²⁺ concentration is proposed to facilitate the site-specific DNA cleavage reaction (Sigman et al., 1991).

Mapping nucleosome positions with OP has been reported using three labeling positions: H4(S47C), H2B(T87C), and H3(Q85C) (Krietenstein et al., 2016; Ghassabi Kondalaji & Bowman, 2022; Henikoff et al., 2014; Fei et al., 2015; Ramachandran & Henikoff, 2016; Chereji et al., 2018; Ramachandran, Zentner & Henikoff, 2015; Moyle-Heyrman et al., 2013; Thakur, Talbert & Henikoff, 2015; Voong et al., 2016) (Figure 2). For both in vitro and in vivo experiments, the site-specific cleavage sites allow one to infer the position of the histone core on DNA. In vitro, cleavage sites are typically analyzed by sequencing gels, whereas cleavage sites from in vivo experiments are mapped with high-throughput sequencing. Whereas all cleavage sites are mapped separately on each DNA strand, the cleavage by H3(Q85C) on both strands offers the additional advantage of producing a 51-bp DNA fragment. By enriching for this 51-bp fragment, which provides confirmation of an intact H3/H4 tetramer, nucleosome positioning and occupancy can be determined with significantly reduced background and greater precision (Chereji et al., 2018).

When nucleosome positions are mapped with OP in vitro, as described in this article, the cleavage sites reveal key characteristics of select factors included in the experiment. For chromatin remodelers, for example, mapping nucleosome positions with OP can reveal characteristics of the enzyme, such as step size, speed, and direction of nucleosome sliding. For nucleosomes, OP mapping can show how nucleosome positioning is affected by characteristics such as flanking DNA length, histone variants, and post-translational modifications. By providing the positioning of histone contacts at bp resolution, mapping with OP also reveals fundamental characteristics of the nucleosome itself, in particular the preferred rotational and translational phasing of a particular DNA sequence on the histone core. High resolution mapping with OP also has the potential to reveal changes in histone-DNA contacts that reflect small and dynamic changes in nucleosome structure that can occur upon binding of chromatin binding factors (Winger et al, 2018; Michael et al., 2020; Matsumoto et al., 2019). Mapping by OP, therefore, provides a relatively quick and efficient means to obtain an overall distribution of favored nucleosome positions for a specific DNA sequence, while also revealing subtle structural changes that reflect local energetics of nucleosomal DNA.

Critical Parameters

Basic Protocol 1

Perhaps the most important consideration for success with OP-mediated mapping is the choice of the histone residue to mutate to cysteine. DNA cleavage is limited to a close proximity (~4 Å) from the OP moiety (Ramachandran & Henikoff, 2016). Three sites that have been demonstrated to work well for OP-mediated mapping are H2B(T87C), H4(S47C), and H3(Q85C) (Fei et al., 2015; Ghassabi Kondalaji & Bowman, 2022; Krietenstein et al., 2016; Ramachandran & Henikoff, 2016; Ramachandran, Zentner & Henikoff, 2015;Chereji et al., 2018) (Figure 2).

Labeling with OP can be significantly affected by the quality of the iodoacetyl OP derivative solution and its concentration in the histone labeling reaction. Using an iodoacetyl OP derivative solution that has been stored > 6 months significantly reduces the efficiency of the labeling reaction. The DNA cleavage patterns obtained from nucleosomes prepared from an old OP solution stock show weak or no cleavage patterns, especially at minor cleavage sites (e.g., +6 in OP-H4(S47C) cleavage patterns; see Understanding Results). Additionally, the labeling reactions and nucleosome reconstitutions from the labeled histones should be performed in the dark, as the OP and iodoacetyl OP derivative are light sensitive. The overnight labeling reactions should not proceed for more than 14 hours, which is sufficient to achieve ~70% labeling of folded histones (Brogaard, 2011). It is recommended to eliminate native cysteines that are incompatible with site-specific labeling (e.g., Xenopus laevis H3(C110)).

To ensure proper mixing of the labeling mixture during incubation time with the OP solution, the tube should have enough room (half-full) to allow the reaction solution to be mixed efficiently on a tube rotator. Care must be taken to minimize sample loss during this process or in other steps, including transferring the quenched labeling mixture in and out of (micro)dialysis.

Basic Protocol 2

When performing nucleosome sliding reactions, consistent timing in quenching the reactions is crucial. Sufficient mixing of the sliding mixture immediately after ATP addition is also critical, to ensure that all ATPases are exposed to the same ATP concentration throughout the time course. When the reactions are to be further processed using Basic Protocol 3, the final quenched reactions should be free from EDTA and, if possible, from salmon sperm (competitor) DNA.

Basic Protocol 3

The quenching of the cleavage reaction takes place by addition of a ligand that coordinates strongly with Cu²⁺ (i.e. neocuproine). Thus, it is highly recommended to avoid buffer compositions that interact with copper and make it inaccessible for OP. For instance, Cl⁻ anions at concentrations higher than 100 mM are suspected to prevent site-specific DNA cleavage, unless the concentration of Cu²⁺ is purposefully increased in the reaction mixture (Sigman et al., 1991). Importantly, conventional thiols such as DTT and βME must be avoided, as they are strong copper-chelating agents, unlike MPA (Sigman et al., 1991). EDTA, a necessary buffer component in many nucleic acid mixtures, has also been shown to interfere with the cleavage reaction through Cu²⁺ chelation (Ghassabi Kondalaji & Bowman, 2022; Sigman et al., 1991) and it is recommended to be absent in the DNA cleavage reaction.

A crucial factor in performing the DNA cleavage reaction is to add the thiol (MPA) and oxidizing agent (H₂O₂) simultaneously. Addition of MPA to the reaction mixture containing Cu²⁺ prior to H₂O₂ addition results in cupric reduction to Cu⁺ without its recycling to the initial oxidation state (Cu²⁺) by H₂O₂. Consequently, MPA is consumed without productive generation of hydroxyl radicals.

Following the cleavage reaction, it is critical to minimize sample loss during DNA extraction and preparation for sequencing gels. The most susceptible steps include the following: (i) taking too little of the aqueous upper layer containing DNA during the phenol-chloroform DNA extraction step; (ii) losing sample by vortexing the container tube with an unsecured lid; (iii) allowing precipitated DNA to resolubilize in the 75% ethanol wash step due to warming up; and (iv) accidentally removing the precipitated DNA along with supernatant in the 75% ethanol wash step.

Support Protocol 1

To accurately identify the DNA fragments extracted from OP-guided cleavage reactions, the dideoxy sequencing ladders must have the same sequence, 5’ end, and fluorophore as the cleaved sample. To achieve this, the Sanger reactions should use the same primers and DNA plasmid used for the large-scale PCR-based synthesis of nucleosomal DNA. Unlike standard PCR reactions, each Sanger reaction uses only one primer (instead of both) so that only one strand is linearly amplified.

When loading the sequencing gel, the volume of the dideoxy ladder mixture should not be more than 7 μL per each well, as the dideoxy ladder mixture will overflow from its corresponding well and enter the adjacent wells.

Support Protocol 2

The significance of a well-prepared denaturing sequencing gel in obtaining high-quality data must not be underestimated. Thorough cleaning of the glass gel plates is crucial, both before pouring the sequencing gel solution and prior to scanning. To prevent irregularities in wells, the comb must not be disturbed before the gel solution is fully polymerized. The temperature of the denaturing gel solution must be kept near 4°C, as fluctuations can stimulate urea crystallization before and during gel pouring. Trapped bubbles inside the gel disrupt the migration of DNA fragments during electrophoresis and it is, therefore, crucial that they be eliminated, if possible, while pouring the gel. Finally, one should consider what lengths of cleaved DNA fragments are expected, and be wary of inadvertently running short DNA fragments off the bottom of the gel.

Troubleshooting

The most common issues associated with described protocols in this article, their possible causes, and strategies to resolve them are listed in Table 1.

Table 1.

Troubleshooting guide for mapping histone repositioning by chromatin remodeling enzymes via OP-directed DNA cleavage reactions

Problem	Possible cause	Solution
Loss of sample (Basic Protocol 1)	Dialysis-related sample loss	Check dialysis tubing for possible leakage
		Avoid vigorous mixing of (micro)dialysis setup to prevent disassembly of (micro)dialysis setup (e.g., bag or chambers)
		Avoid overloading microdialysis chambers (≤ 150 μL per tube)
		Do not use dialysis tubing with large molecular weight cutoff value (> 3.5 kDa) to prevent passing the labeled histone core through the membrane
	Miscellaneous sample loss	While concentrating the labeled histone core for long-term storage, use the centrifuge speeds that are less than the maximum g force of the concentrators.
Low intensity or an absence of cleavage bands (Basic Protocol 3)	Inadequate OP-labeling of the histone core (Basic Protocol 1)	Use a fresh OP solution (less than 6 months old)
		Check the pH, concentration, and volume of TCEP solutions to assure ensure complete reduction of disulfide bonds
		Check the concentration and volumes of OP solution added to labeling reaction mixture
	High amounts of EDTA in sliding/quenching mixtures containing OP-labeled nucleosomes	If possible, replace EDTA. For instance, use hexokinase instead of EDTA to quench sliding reactions, as described in Basic Protocol 2
High background in cleavage pattern obtained from scanning the denaturing sequencing gel (Support Protocol 2)	Large amounts of non-target DNA in buffer used to quench reactions	Avoid using competitive DNA, and instead use hexokinase to quench sliding reactions
	Unclean gel plates	Thoroughly clean glass plates, as described in Support Protocol 2.
Gel solution polymerized while pouring gels	High amount of added APS and TEMED	If the room temperature is ≥ 25°C, pour the gel with 10% lower amounts of APS and TEMED
	Warmed-up gel solution	Keep the acrylamide solution at 4°C until ready to add APS and TEMED, and pour the gel immediately
Malformed wells in sequencing gel (Support Protocol 2)	Leakage of the gel solution from in between the gel sandwich	Leakage of the gel equally affects all the lanes. If the wells are too small to hold the individual sample solutions, or are not formed at all, prepare a new gel. Use the wells if the leakage is not severe and the volume of the formed wells is enough to hold the loaded sample
	Disturbing the comb before the gel is fully polymerized or inappropriate removal of the comb after polymerization	Remove the comb by slowly pulling it from the two ends simultaneously, to prevent disruption of the wells. If the wells are formed properly but are bent, use a P10 or P20 pipette tip to straighten the lanes.
Running buffer leaking between the short plate and aluminum plate, or from the contact sites between the silicon gasket and sealing foam blocks of the spacers (Support Protocol 2)	Sequencing gel is not completely sealed against the S2 apparatus	Slightly loosen the clamps by unscrewing the knobs at the top of the gel by half a turn. This may release an over-pressed and wrinkled silicon gasket. If the leakage does not stop, tighten the screw knobs.
		Apply grease to the side of the gel sandwich where the solution is leaking. Normally, the leakage happens at the border of aluminum plate, silicon gasket and edge of gel at the top.
		If the leakage does not stop, remove the gel sandwich. Inspect the silicon gasket and sealing foam blocks for any tearing or damage, apply extra grease to previously leaking sites and put the gel back on S2 apparatus.
Smeared or blurred bands on sequencing gel (Support Protocol 2)	Debris, particles, or bubbles inside the wells or within the polymerized gel	Filter (0.22 μm) and degas the gel solution. Check the gel immediately after placing the comb for the bubbles in the vicinity of the comb teeth. If there are bubbles and extra gel solution is available, take out the comb, immediately add more gel solution, and replace the comb
	Urea crystals inside the wells while loading the DNA sample	Flush the wells twice (before pre-run and immediately before loading).
	Small fragment formamide loading dye is too old or not properly stored	To avoid formamide degradation, store aliquots with fragment formamide loading dye at −20 °C. Avoid freeze/thawing
	High salt content from precipitated DNA	Avoid using NaCl instead of 3M NaOAC/HOAC to precipitate DNA
	Overloaded DNA	Keep the amount of nucleosome/hexasome per lane below 2 pmol.
	Overheated gel sandwich	Run the gel at constant 65 W, not constant voltage.
In the sequencing gel, different migration of the bands in the middle compared with the sides of the gel	Uneven pressure applied to the sides of the gel during polymerization	Place binder clips symmetrically on both sides of the gel and have equal spacing between the placed clips.
	Uneven pressure applied to the sides of the gel during electrophoresis	Do not overtighten the internal gel clamps on S2 apparatus.
Appearance of dark spots after gel scan with excitation at wavelengths corresponding to one or both fluorophores (Basic Protocol 3, Support Protocol 2)	Contamination in Ultrapure water (e.g., mili-Q) used to prepare the gel solutions or running buffers	Change the filter on ultrapure water source or use a different ultrapure water source

Understanding Results

The sites of DNA cleavage are revealed by scanning DNA sequencing gels, as described in Basic Protocol 3. When each DNA strand is labeled with a different fluorophore, two scans can be obtained from one gel, with each scan corresponding to one DNA strand. The gel scans are then analyzed using an image processing software such as ImageJ (Schneider, Rasband & Eliceiri, 2012). If the signals are not saturated, their intensities can be used to quantify the relative proportion of different cleaved species.

As shown in Figures 2 and 5, histone-DNA contacts in the vicinity of the H2A/H2B dimer can be determined by OP-labeling H2B(T87C) (Ghassabi Kondalaji & Bowman, 2022; Fei et al., 2015), which produces two cut sites on each strand. Appearance of these two cut sites corresponds to dimer-DNA contacts on both side of the nucleosome (one cut for each labeled H2A/H2B dimer) and indicates that the DNA is wrapped on both sides. These cleavage pairs are separated by ~70 bp from one another. If the position of the histone core on DNA changes (e.g., by remodelers) or if the nucleosomal DNA does not contain a well-positioning sequence, such as the Widom 601 (Lowary & Widom, 1998), multiple cleavage pairs (each ~70-bp apart) should be observed. Each pair would represent a unique positioning of the histone core on DNA

For the canonical 601 nucleosome, the H2B(T87C)-directed cleavages occur at nucleotides −36 and +34 relative to the dyad on one strand, and +/− 35 on the other strand (Figure 2) (Ghassabi Kondalaji & Bowman, 2022). The DNA cleavage pattern obtained from OP-labeled H4(S47C) nucleosomes shows two single stranded cuts at nucleotides positioned −1 and +6 relative to dyad on each DNA strand (Figure 2) (Krietenstein et al., 2016; Ghassabi Kondalaji & Bowman, 2022; Ramachandran & Henikoff, 2016). Figure 5 shows examples of both OP-cleavage sites for well-positioned Widom 601 nucleosomes. The cleavage sites are identified by comparing the bands with the dideoxy ladders matching the template DNA sequence.

In Basic Protocol 2, a method for repositioning nucleosomes using the Chd1 remodeler is described, with quenched reactions providing material that can mapped using Basic Protocol 3. As the remodeler repositions the nucleosomes, the histone core changes its position on the DNA (Figure 5a,b). When centered nucleosomes made with the Widom 601 sequence are remodeled by Chd1, the histone core moves toward the TA-poor side in ~10 bp steps (Winger & Bowman, 2017). As a result, the initial sites of cleavage shift over time, consistent with repositioning of the histone core along DNA. These shifts can be visualized by following the cleavage pattern on either strand of DNA.

In addition to histone repositioning, the expected OP-cleavage pattern may change due to loss of histone-DNA contacts. Hexasomes made with the Widom 601 sequence preferentially lack the H2A/H2B dimer on the TA-poor side (Levendosky et al., 2016). Thus, hexasomes only produce the cut sites on the TA-rich side of the 601 sequence (Ghassabi Kondalaji & Bowman, 2022). Transcription factors may interfere with DNA cleavage when their binding sites are close to the site of cleavage (Ghassabi Kondalaji & Bowman, 2022).

Time considerations

The protocols described here for labeling folded histone(s) (Basic Protocol 1), repositioning nucleosomes with a chromatin remodeler (Basic Protocol 2), cleaving and processing DNA (Basic Protocol 3), and resolving DNA fragments on a denaturing sequencing gel (Support Protocols 1 and 2) span approximately 7 to 9 days. This timeframe includes nucleosome reconstitution and purification steps, which are described elsewhere (Nodelman et al., 2020). Before beginning these protocols, additional time is also required for producing and purifying the histones and DNA, and then folding and purifying histone dimer, tetramer, and/or octamer (Dyer et al., 2004). Once all reagents are in-hand, a main factor affecting the time between OP labeling and resolving cleavage products on a sequencing gel will depend on the needs for intermediate experiments designed to alter histone-DNA contacts (e.g., Basic Protocol 2, Figure 1). Any such intermediate experiments should be followed immediately by DNA cleavage, extraction, and precipitation steps (day 1) in Basic Protocol 3.

The timing of major steps is described in the following sections.

Basic Protocol 1

The entire process can be completed in ~42 hours. Multiple histone core proteins (H2A/H2B dimer, H3/H4 tetramer and histone octamer) can be labeled simultaneously.

• Day 1: Preparing solutions and materials (≤ 2 hours); OP labeling at RT (2 hours); overnight labeling (12 to 14 hours).

• Day 2: Incubation of labeling mixture with quencher (≤ 1 hour); preparing labeling reactions for dialysis (≤ 1 hour); 1^st dialysis of quenched labeling mixture against post-labeling dialysis solution (overnight, ≥12 hours).

• Day 3: 2^nd dialysis (≥3 hours); preparing OP-labeled histones for long-time storage (2–3 hours).

Basic Protocol 2

As mentioned earlier, Basic Protocol 2 is introduced in this article as an example of an in vitro experiment that alters histone-DNA contacts, and is compatible with DNA cleavage reactions in Basic Protocol 3. The time required for Basic Protocol 2 is primarily dependent on the number of quenched sliding reactions and the length of the time course, but, generally, it can be carried out in ~3 hours. A few additional hours should be allotted for processing quenched sliding reactions as part of Basic Protocol 3. Generally, it is feasible to carry out as many as 72 sliding reactions and subsequently perform DNA cleavage reactions (start of Basic Protocol 3) on the same day.

Basic Protocol 3 and Support Protocols 1 and 2

The first steps of Basic Protocol 3 (DNA cleavage, processing, and DNA precipitation) should be carried out as soon as samples are ready to be processed (e.g., quenched sliding mixtures in Basic Protocol 2, or directly from OP-labeled nucleosomes). Two stages where the procedure can be paused are during DNA precipitation (where samples can be stored at ≤ −20 °C indefinitely; end of day 1 of Basic Protocol 3) and after the DNA pellet has been resuspended in small fragment formamide loading dye (also to be stored at −20 °C until needed; day 2 of Basic Protocol 3). A major time constraint is the preparation of the DNA sequencing gel (Support Protocol 2). In addition to preparation and assembly of the glass plates and pouring the gel, full polymerization requires at least 6 hr, and can be left overnight. Preparation of a dideoxy sequencing ladder (2–3 hours; described in Support Protocol 1) can be performed during downtime on day 2 of Basic Protocol 3. Alternatively, a dideoxy sequencing ladder can be prepared days in advance, as the ladders can be stored at −20°C indefinitely. The entire process from start of the DNA cleavage reaction to scanning the denaturing sequencing gel can, therefore, be performed in one to three days, depending on when the sequencing gel is prepared.

Below is the detailed breakdown of the process from OP-directed DNA cleavage to scanning the sequencing gel:

• Performing the DNA cleavage reaction on OP-labeled nucleosome mixtures such as quenched sliding aliquots (~0.5–2 hours); post-cleavage heating and deproteinization of cleaved DNA solutions (20–30 minutes); phenol-chloroform extraction (1–3 hours, depending on the number of cleaved DNA samples); DNA precipitation (1 hr to overnight); pelleting the precipitated DNA (≤ 1 hour); air-drying and resuspending the DNA pellet (≥ 1.5 hours); making the dideoxy sequencing ladders (2–3 hours); setting up and pouring the denaturing sequencing gel (~2 hours); polymerizing the denaturing sequencing gel (6 hours to overnight); preparing the polymerized gel for run (1 hour); pre-running the sequencing gel (~45 minutes); loading and running the sequencing gel (1.5–2 hours); cleaning the gel and scanning (Typhoon imager) at two wavelengths (≤ 1.5 hours).

Abbreviations

APB

4-azidophenacyl bromide

APS

Ammonium persulfate

ATP

Adenosine triphosphate

βME

2-Mercaptoethanol

BSA

Bovine serum albumin

Chd1

Chromodomain helicase DNA binding protein 1

Cy3

Cyanine-3

Cy5

Cyanine-5

DI

Deionized

dATP

deoxyadenosine triphosphate

dCTP

deoxycytidine triphosphate

dGTP

deoxyguanosine triphosphate

dTTP

deoxythymidine triphosphate

dNTP

deoxynucleotide triphosphate

ddNTP

dideoxynucleotide

DMSO

Dimethyl sulfoxide

DTT

Dithiothreitol

FAM

Fluorescein amidite

MPA

3-mercaptopropionic acid

NCP

Nucleosome core particle

OP

ortho-phenanthroline

PCR

Polymerase chain reaction

PMT

Photomultiplier tube

RT

Room temperature

SDS

sodium dodecyl sulfate

TAU

Triton acetic acid urea

TBE

Tris-borate-EDTA

TCEP

Tris(2-carboxyethyl)phosphine

TEMED

Tetramethylethylenediamine

TF

Transcription factor

DATA AVAILABILITY STATEMENT

Data sharing not applicable to this article as no datasets were generated or analyzed during the current study