Abstract
The dynamics of histone-DNA interactions govern chromosome organization and regulates the processes of transcription, replication, and repair. Accurate measurements of the energies and the kinetics of DNA binding to component histones of the nucleosome under a variety of conditions are essential to understand these processes at the molecular level. To accomplish this, we employ three specific single-molecule techniques: force disruption (FD) with optical tweezers, confocal imaging (CI) in a combined fluorescence plus optical trap, and survival probability (SP) measurements of disrupted and reformed nucleosomes. Short arrays of positioned nucleosomes serve as a template for study, facilitating rapid quantification of kinetic parameters. These arrays are then exposed to FACT (FAcilitates Chromatin Transcription), a non-ATP-driven heterodimeric nuclear chaperone known to both disrupt and tether histones during transcription. FACT binding drives off the outer wrap of DNA and destabilizes the histone-DNA interactions of the inner wrap as well. This reorganization is driven by two key domains with distinct function. FD experiments show the SPT16 MD domain stabilizes DNA-histone contacts, while the HMGB box of SSRP1 binds DNA, destabilizing the nucleosome. Surprisingly, CI experiments do not show tethering of disrupted histones, but increased rates of histone release from the DNA. SI experiments resolve this, showing that the two active domains of FACT combine to chaperone nucleosome reassembly after the timely release of force. These combinations of single-molecule approaches show FACT is a true nucleosome catalyst, lowering the barrier to both disruption and reformation.
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Key words
1 Introduction
The nucleosome coordinates the histone octamer and ~2 wraps of DNA to form the basic subunit of chromatin (Fig. 1a). Histones are the most abundant nuclear protein, and four specific types, H2A, H2B, H3, and H4, combine in a symmetric disc that contacts the wrapped DNA in hundreds of histone-DNA and water-mediated histone-DNA contacts [1]. A total of 147 base pairs wrap into the nucleosome, centered on the (H3-H4)2 tetramer, and two H2A/H2B dimers (Fig. 1b). A central 80 base pairs constitute the inner wrap, beginning at the dyad center of the tetramer and extending to each dimer, in direct contact with these core or strong sites [2, 3]. The remaining bases bind ~ 1/2 wraps above and below the inner wrap (Fig. 1c). Though this structure is stable, it is also highly dynamic, as local regions of DNA-histone separation, or breathing, occur. Histone-DNA binding must be disrupted to facilitate many cellular processes including replication, transcription, and repair [4, 5].
Among the variety of proteins that disrupt isolated nucleosomes, the non-ATP-driven heterodimer FACT consists of the 140 kDa SPT16 (SuPpressor of Ty elements 16) and 80 kDa SSRP1 (Structure Specific Recognition Protein 1), joined at dimer domains (DD). Several domains within these subunits contact to both disrupt and reassemble the nucleosome [4, 6,7,8,9,10,11]. Recent cryoEM structures show tetramers bound to FACT (missing the HMGB domain), displacing the H2A/H2B dimer [12], while another series of structures show FACT (plus an accessory protein) tethering histones during the passage of human RNA polymerase II (RNAPII) [13]. Domains of human HMGB exist in the nucleus both as part of FACT and alone, while the HMG box found in yeast, NHP6A, is not directly bound to the yeast analogue of FACT, but coordinates with it. Both HMGB and NHP6A are known to bind to and disrupt DNA wrapped around the nucleosome [21]. Single-molecule experiments on chromatin have yielded insights into chromatin structure ranging from measurements of the mechanical elasticity of a chromosome down to the protein-DNA interactions holding DNA to single nucleosomes [17, 22,23,24]. Here, we will assemble a short array of individual nucleosomes that facilitate the rapid collection of data on both energies and kinetics of histone-DNA interactions. Force disruption (FD) experiments characterize the strength of histone-DNA interactions and facilitate the construction of a free energy landscape. Confocal imaging (CI) experiments probe the kinetics of disrupted DNA-histone interactions. Finally, survival probability (SP) experiments reveal the extent of nucleosome reformation as tension is released.
The following experiments reveal that FACT binds to the nucleosome, releasing the outer 1/2 wraps of DNA and destabilizing the DNA-histone interactions of the remaining inner wrap. Within FACT, two domains, the SPT16 MD and SSRP1 HMGB, function effectively to bind the histone and the DNA, substituting direct DNA-histone contacts with contacts mediated by FACT. SSRP1 HGMB plays a significant role in nucleosome destabilization, while yeast NHP6A shows even greater activity. Yet only the full protein is able to simultaneously destabilize the nucleosome and tether the components for reassembly after release of tension. Full FACT acts as a true nuclear chaperone.
2 Materials
Several studies have detailed the assembly of human histone octamers, as well as their reconstitution onto DNA [25,26,27,28]. Furthermore, protocols for purification of the proteins studied here have been described. The goal of this writing is to present a general technique to study the effectiveness of nucleosome chaperones. Some familiarity with octameter assembly, fluorescent dye labeling, and protein purification is assumed. The following protocol will outline the key steps to create the DNA template and then combine octamers to create a 12× nucleosome array for study. A brief outline of the expression and purification of the proteins specifically is used here, though these techniques are useful for any nucleosome binding ligand.
2.1 Instruments, Reagents, and Buffers for Sample Preparation
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1.
E. coli Rosetta(DE3)pLysS and E. coli BL21(DE3)pLysS cells for expression (MilliporeSigma).
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Plasmid pET28 (modified), pTEV, and pCOLADuet cloning vectors.
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3.
Ni-NTA column (Qiagen).
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Bind buffer: 50 mM NaH2PO4-Na2HPO4, pH 7.5, 300 mM NaCl.
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Wash buffer: 50 mM NaH2PO4-Na2HPO4, pH 7.5, 300 mM NaCl, 20 mM imidazole.
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Elution buffer: 50 mM NaH2PO4-Na2HPO4, pH 7.5, 300 mM NaCl, 200 mM imidazole.
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Superdex 200 column (GE Healthcare) for size-exclusion chromatography with Bind buffer as running buffer.
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TEV protease (stock in glycerol) with 50 mM sodium phosphate pH 6.5, 50 mM NaCl, for SSRP1 HMGB storage.
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Protein dialysis and storage buffer: 20 mM HEPES pH 7.5, 100 mM KCl, 1 mM EDTA, 1 mM DTT, and 5% glycerol.
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pU19 plasmid and 601 Widom positioning sequences.
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Bsal I restriction enzyme.
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12.
DNA pol I.
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Digoxygenin-11-dUTP (Roche), biotin-16-dUTP (Sigma), biotin-14-dCTP (Invitrogen) and biotin-14-dATP (Invitrogen).
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Human octamers and human octamers carrying Alexa488 or Atto647N on each H2B (T112C).
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Nucleosome dialysis buffer, high salt: 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, with Na+ of 1 M.
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16.
Nucleosome dialysis buffer, intermediate salt: 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, and Na+ of 750 mM.
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17.
Low salt nucleosome dialysis buffer and storage solution: 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, with Na+ of 2.5 mM.
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18.
Individual solution of 5 M NaCl.
2.2 Instruments and Buffers for Single-Molecule Experiments
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A fiber coupled laser, up to 200 mM at 850 nm (see Note 1), split 50:50 and directed to form a dual beam counterpropagating single trap. A confocally arranged pair of water-dip** 1.0 NA objectives is adjusted to overlap the two traps. Sub-picoNetwon forces on the trapped bead are determined using a lateral effect detector (Melles Griot) to measure trap** beam deflection, which is proportional to the applied force.
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A home-built flow cell and sample chamber. A custom plexiglass spacer is machined and covered on the sides with 22 × 30 mm glass cover slips, fastened with UV-curing epoxy. Inlet and outlet tubing is also glued in place at opposite ends of the chamber.
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Pre-pulled glass micropipettes with an opening of 0.5 micrometers (World Precision Instruments), glued into place through a hole drilled at 45 degrees from vertical (see Note 2).
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The LUMICKS dual beam imaging C-trap is a commercial system capable of simultaneous force disruption (FD) experiments (complementing the instrument described above) and confocal imaging (CI).
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Polystyrene beads, 1.0% w/v, functionalized with a coating of streptavidin, and with a diameter of 1.7, 3.1, or 5 micrometers (see Note 3).
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6.
2 μm diameter polystyrene beads, 0.5% w/v, with digoxygenin coating.
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Experimental buffer: 10 mM HEPES, pH 7.5 and Na+ of 50 or 100 mM.
3 Methods
Many excellent reviews summarize various optical tweezer designs and experiments [18,19,20,21, 29, 30]. Here, tweezers will apply tension to reconstituted DNA arrays described above in force disruption (FD), confocal imaging (CI), and survival probability (SP) assays. These experiments determine the stability of DNA-histone contacts and the length of DNA wrapped around the octamer. Protein binding affects both the wrapped length and stability, allowing the binding affinity to be determined. The kinetics of histone-DNA interactions and the chaperone activity of these proteins after the release of tension is characterized.
3.1 FACT Protein and Isolated Domain Production
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Human FACT SPT16 and SSRP1 as well as isolated SPT16 MD and SSRP1 HMGB are to be expressed in E. coli and purified to homogeneity (see Note 4). Full SPT16 and SPT16 MD are cloned in a modified pET28 vector, while SSRP1 HMG and SPT16 CTD are cloned in a pTEV vector. An N-terminal His6 tag is included to facilitate purification. SSRP1 is cloned in a pCOLADuet plasmid, without a tag.
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E. coli Rosetta(DE3)pLysS and E. coli BL21(DE3)pLysS cells are used to express all subunits/domains, grown in LB broth at 37°C to an OD600 nm of approximately 0.6, and then induced with 0.5 mM isopropyl-β-D-thiogalactoside at 15°C for 16–20 h.
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Harvest the cells by centrifugation. Discard the supernatant. Resuspend the pellets in 10 mL of Bind buffer. Lyse the cells using an Avestin EmulsiFlex C5 homogenizer. PMSF is added before cell lysis (see Note 5).
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Centrifuge at 48,000 × g for 45 min to separate supernatant from pellet. Apply the supernatant onto a Ni-NTA column pre-equilibrated with Bind buffer at 4 °C. Extensively wash the column with Wash buffer. Elute the protein with Elution buffer.
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Concentrate the protein solution. Cut the His6-MBP tag of SPT16 (501-1006) using TEV protease at 4 °C overnight.
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Separate the FACT complex from His6-MBP by size-exclusion chromatography with a Superdex 200 column using Bind buffer as running buffer. Pool all the fractions containing FACT, concentrate to proper volume and store at −80 °C until use.
3.2 NHP6A Protein Expression and Purification
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Recombinant untagged yeast NHP6A protein is expressed in bacteria and purified by HPLC [31].
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Further purification is achieved via size exclusion chromatography in phosphate buffered saline on a Superdex 200 10/30 column. Elution occurs at a flow rate of 0.4 mL/min.
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Fractions are finally pooled and dialyzed against protein dialysis buffer.
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Proteins are stored at −20 °C in dialysis buffer and supplemented with 50% (v/v) glycerol.
3.3 Combining Nucleosome Position Sequences and Handles to Form Templates
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Two DNA constructs (pJ1937 and pJ2774) are created utilizing 12 tandem copies of the Widom 601 nucleosome positioning sequences, inserted into a pUC19-based plasmid.
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Each Widom positioning sequence is comprised of 147 base pairs that contact the histone octamer and 60 base pairs to serve as a linker (see Note 6).
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The first construct, a linear “pJ1937” template, is prepared by BsaI restriction endonuclease cleavage to create flanking non-nucleosomal “handles” of 1340 and 1360 base pairs. BsaI, a type IIS restriction endonuclease, creates distinct four-base overhanging termini allowing DNA polymerase I repair of the overhangs in the presence of single digoxygenin (digoxigenin-11-dUTP) and biotin (biotin-14-dATP) nucleotide analogs on opposing termini, enabling capture for force-extension experiments (Fig. 2a, see Note 7).
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The second construct, the linear “pJ2774” template, employs longer (3400 bp) “handles” flanking the 12 Widom 601 sequences (Fig. 2b). The template is digested with BsaI restriction endonuclease, and overhangs are filled in with the Klenow fragment of DNA polymerase I in the presence of biotinylated nucleotide analogs labeling both DNA termini (biotin-16-dUTP, biotin-14-dATP, biotin-14-dCTP; see Note 8).
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5.
The biotin- and digoxigenin-labeled DNA was purified by spermine DNA precipitation and resuspended in water prior to assembly with the human histone octamers.
3.4 Reconstituting Nucleosome Arrays
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Pre-assembled octamers are combined with the DNA constructs at high concentrations (ng/μL, see Note 9), in low salt dialysis buffer, titrated with isolated 5 M Na+ solution to a final concentration of 2 M Na+.
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2.
Combined samples are placed into a small volume dialysis button (Hampton Research, 50 μL, see Note 10), and the button is sealed with dialysis membrane (see Note 11) and placed in 200 mL of high salt dialysis buffer.
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Dialysis decreases Na+ from 2 M to 1 M over 4–6 h.
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Discard the high salt buffer and replace with intermediate buffer, dialyzing for 4–6 h.
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Finally, replace buffer with low salt buffer and dialyze for another 4–6 h.
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Reconstituted arrays remain stored in low salt dialysis buffer for storage and are stable for several weeks at 4 °C (see Note 12).
3.5 Force Disruption (FD) Using Optical Tweezers
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Stock solutions of streptavidin coated and anti-digoxigenated coated beads are diluted 1000:1 and 1000:2, respectively, into the experimental buffer at ambient lab temperature of 22 °C (see Note 13).
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One 3-micron diameter streptavidin coated bead is caught and held in the optical trap, while a 2-micron diameter anti-digoxigenin coated bead is pulled onto the micropipette tip (see Notes 14 and 15).
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Flow reconstituted 12-mer nucleosome arrays diluted 10,000:1 in experimental buffer into the sample cell (see Note 16). Arrays quickly saturate the flow cell chamber. An array is secured between the beads by rubbing the smaller bead on the micropipette tip against the bigger bead on the trap and slightly tugging out and looking for some resistance (see Note 17).
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4.
Once the nucleosome array is tethered, the piezo stage increases the extension stepwise in 4 nm increments, with a force measurement after each step (see Notes 18 and 19). After a short pause at ~40 pN, the array should be gradually released, also in 4 nm steps (see Note 20).
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5.
Data for the nucleosome array stretching shows a smooth increase in extension with force at the beginning (see Notes 21 and 22).
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6.
Nucleosome outer wrap release will be seen as a shoulder in the low force region (~2–5 pN, see Fig. 3a).
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7.
As force increases further (>10 pN), sudden, consecutive increases in extension will be seen. These correspond to nucleosome inner wrap (core) openings. Discrete rips appear, in the which the extension change is due to base pairs unwrapped and released from each nucleosome. All wild-type nucleosomes will be unwrapped once the applied tension reaches ~40 pN.
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8.
Opening events at higher forces are due to 2× or even 3× arrays tethered simultaneously between the same streptavidin-anti-digoxigenin bead complex. It is difficult to resolve the data due to multiple arrays and to physically separate multiple arrays from the same bead. These curves and beads should be discarded, and the fresh beads caught.
3.6 Quantifying Force Disruption Data
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1.
With calibration of the force and extension, the full cycle of extension and release is methodically compared to a well-known model of polymer elasticity. The extensible Worm-Like Chain (eWLC) is used to characterize double-stranded DNA over a wide range of solution conditions. An analytical solution is known for the high force limit [19, 32,33,34,35],
$$ b(F)=B\left[1-\frac{1}{2}{\left(\frac{k_BT}{PF}\right)}^{1/2}+\frac{F}{S}\right]\cdot $$(1)-
The measured extension as a function of the applied tension, b(F), is determined by a set of intrinsic polymeric parameters: the end-to-end contour length (B), the polymeric bending stiffness known as the persistence length (P), and the enthalpic stretch modulus (S, see Note 23). Values of these parameters have been measured in many circumstances (see Note 24), and typical values match those fitted to the constructs measured here: B = 0.340 ± 0.001 nm/bp, P = 42 ± 1 nm, and S = 1000 ± 100 pN.
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2.
Changes in the measured length of the DNA (Fig. 3a) are attributed to DNA unwrap** from the nucleosome. Beginning with the fully unwrapped construct, at maximum extension, a custom algorithm searches for divergences between the eWLC model and the data (see Note 25). Each disruption is attributed to a disruption of the inner wrap, and both the force and extension change (see Note 26) are recorded and plotted against the number remaining (A, see Fig. 3b, c as well as Table 1 for n = 25 arrays).
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3.
Continuing to low force, a small plateau may be observed below 10 pN, and the extension change here is due to collective unwrap** of the outer 1/2 wraps. Table 1 summarizes the sum of the length of all released DNA.
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4.
Unwrap** the core DNA above 10 pN gives distinct events that may be arranged in order of observation (here by the number remaining, A). Despite variations, the averaged force shows a clear increase as the number remaining decreases. This is due to the nonequilibrium rate of release, relative to the loading rate of the instrument, and can be described by a simple model [17]:
$$ F=\frac{k_BT}{x_{\mathrm{core}}^{\dagger }}\cdot \ln \left[\frac{\mathrm{d}F}{\mathrm{d}t}\cdot \frac{x_{\mathrm{core}}^{\dagger }}{k_BT\cdot {k}_0\cdot A}\right]\cdot $$(2)-
The observed rip** force (F) will increase as the number remaining (A) decreases, depending upon the loading rate (dF/dT), the distance from the wrapped state to the barrier (x†core), and the natural (zero force) rate of DNA-histone opening fluctuations (ko). Fits to the data averaged at each A (Fig. 3b) determine this rate and the distance to the barrier.
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5.
The measured extension change, once corrected for force dependent polymer elasticity (Eq. 1), does not vary within uncertainty across the number released (Fig. 3c) and may be simply averaged across all A.
3.7 Evaluating the Free Energy and Transition Barrier
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1.
The free energies of the inner and core releases are found in distinct ways. The release of the outer 1/2 wraps, though indistinguishable, is in equilibrium, and the free energy is simply determined by the area between the measured change in the extension (Fig. 3a shows ΔGouter and see Note 27). The measured value of ΔGouter = 14 ± 2 kBT compares well with previous measurements (see Table 2) [3].
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2.
The inner wrap release is a nonequilibrium event, and the area between the polymer models is now formally the work done by the instrument (WA) at each release,
$$ {W}_A=\Delta {G}_A-\Delta {G}_{A-1}+{W}_{\mathrm{stiffness}}\cdot $$(3)
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This work is the difference between energy required to extend the array with A nucleosomes remaining less the array with A − 1 nucleosomes (shaded in blue in Fig. 3a, see Note 28).
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3.
The free energy of the inner wrap release is deduced from the work, using the method of Jarzynski (see Note 29) [36],
$$ \Delta {G}_{\mathrm{core}}=-{k}_BT\cdot \ln \left[\sum {e}^{-W/{k}_BT}\right]. $$(4)
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The measured value of ΔGcore = 62 ± 2 kBT is substantially stronger than the energy holding the outer 1/2 wraps.
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4.
Distributions of each force release (at each A, shown in Fig. 3b) can determine transition state parameters, using a simplified form of the solution of the transition state theory of Dudko [37, 38],
$$ {G}_{\mathrm{core}}^{\dagger }=\frac{F_{\mathrm{max}}\cdot {x}_{\mathrm{core}}^{\dagger }}{k_BT}\left[\frac{x_{\mathrm{core}}^{\dagger }}{x_{\mathrm{core}}^{\dagger }-{P}_{\mathrm{max}}\cdot {k}_BT\cdot e}\left(1-\nu \right)\right]\cdot $$(5)
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The transition energy barrier (G†core) is calculated from the distribution peak probability (Pmax, see Note 30) at the peak probability force (Fmax, see Note 31). The value of the distance to the transition state (x†core) is chosen using the values determined from fits to Eq. 2 (see Notes 32 and 33).
3.8 Characterizing the Effects of a Chaperone Protein
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Nucleosomes are incubated in specific protein concentration and tethered in the flow cell. A full FD cycle is sought and analyzed as for isolated arrays shown above (see Note 34).
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Force extension curves (Fig. 4a for examples) are scrutinized for changes in length due to unwinding of the outer 1/2 wraps and the inner wrap, as described above.
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The measured force of the inner wrap release is the most sensitive method for judging ligand activity, as compared to the measured length, or other parameters that result from additional analysis steps. Forces are averaged across the entire array (Fig. 4b) for a single cycle of extension and release, and fresh arrays are averaged together (n is typically three or greater). Proteins that destabilize core DNA-histone interactions include human HMGB isolated from SSRP1 and yeast NHP6A, and lower forces are required to fully disrupt the core wrap. In contrast, STP16 MD stabilizes histone-DNA interactions as this protein must bind both histones and DNA and must be removed to facilitate further unwrap**.
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4.
The concentration-dependent changes to the measured disruption force is fit to a simple model that combines the Hill equation with a linear model of changing force with protein binding [17, 35],
$$ F\left(\Theta \right)={F}_{\mathrm{nucl}}-\left({F}_{\mathrm{nucl}}-{F}_{\mathrm{protein}}\right)\cdot \Theta \cdot $$(6)
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Here, F(Θ) is the measured average force of inner wrap disruption for a given fractional occupancy Θ (Θ, which is related to the ligand concentration through the Hill Eq. (see Note 35). Fnucl is the average force in the absence of any ligand, and Fprotein is the value determined at saturation. Nonlinear fits determine the equilibrium binding dissociation constant (KD), summarized in Table 1 (see Note 36). The equilibrium dissociation constant for full FACT (KD = 26 ± 3 nM) is less than the individual active domains and compares well with other techniques [39].
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5.
Both the released length of the outer 1/2 wraps and the core wrap are measured to find the total length (Fig. 4c and see Note 37). Saturating concentrations of protein clearly displace the outer 1/2 wraps and modestly release of the core wrap is seen, though the histone-DNA interactions of the core sites (the tetramer and the dimer) appear largely intact.
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6.
To separately distinguish protein affinity for the nucleosome and for free DNA, the titration may be repeated on the DNA construct in the absence of any nucleosome. As there is no nucleosome disruption to characterize, measured force extension curves are fit to the eWLC model of Eq. (1). Protein binding will alter the flexibility of the DNA, and this is quantified through fitted changes to the measured persistence length, P(Θ) (see Note 38) [17, 35],
$$ \frac{1}{P\left(\Theta \right)}=\frac{1-\Theta}{P_{\mathrm{DNA}}}+\frac{\Theta}{P_{\mathrm{protein}}}. $$(7)
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As above, Θ is the fractional occupancy of DNA binding sites. In the absence of any protein, DNA has a persistence length of PDNA (see Note 39), and when saturated with protein, the value will be Pprotein. Measured and fitted changes in the persistence length vary with different protein concentrations (Fig. 4d). Nonlinear fits find the equilibrium binding dissociation constant (KD), describing the affinity to DNA, summarized in Table 2 (see Note 40).
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7.
For all proteins studied here, the values of KD reveal a stronger affinity for the complete nucleosome than for bare DNA (Fig. 4e and Table 1). The reason varies by protein. In the case of both NHP6A and SSRP1 HMGB, this is due to the well-known preference of HMG for bent DNA, which binds the entry/exit points of the nucleosome, while binding more weakly to bare DNA. By contrast, SPT16 MD simultaneously binds both DNA and the histones of the nucleosome. The activity of full FACT is driven by a combination of these effects, plus the coordinated DNA/histone binding of other subunits (notably SPT16 CTD) [39].
3.9 Quantifying Changes to the Energy Landscape
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Fitting the release forces to averaged arrays in the presence of protein shows increases in the kinetics of histone-DNA interactions (Fig. 5a, b) for full FACT and SSRP1 HMGB.
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In the presence of saturating concentrations of protein, the measured energy of nucleosome release increases for SPT16 MD, while increasing for all other proteins.
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3.
The transition state barrier is found as above. Values are summarized in Table 2 and graphically in Fig. 5c and compare to the values obtained for the nucleosome alone (see note).
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4.
These landscape parameters confirm the stabilizing role of SPT16 MD, while SSRP1 HMGB both weakens DNA-protein interactions across the core sites, increasing the kinetics of histone-DNA fluctuations.
3.10 Confocal Imaging (CI) and Kymographs in Combined Instrument
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1.
Experiments begin with construct assembly in a laminar flow cell, commercially produced for the LUMICKS confocal C-Trap (Fig. 6a). Streptavidin-coated beads are caught at the focus of a split 1064 laser (see Note 41). Nucleosome arrays are tethered, translated into a buffer (protein and array free) channel. A quick confocal image verifies a clean catch of a single array (Fig. 6a, inset).
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2.
Confocal images are first collected at a fixed force of ~1 pN (Fig. 6b, see Note 42) and then at 40 pN (Fig. 6c), after progressive disruption of the full array (with all released). Though the images are diffraction limited, the extension change due to wrap** may be observed (Fig. 6b, c).
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3.
Once a fresh array is tethered and translated into a free channel (see Note 43) or a channel containing saturating values of protein (see Note 44), a kymograph measures array intensity versus time (see Notes 45 and 46). Kymographs are initiated at 1 pN, and the force is increased again to 40 pN to complete core disruption (the resulting change in extension is visible in Fig. 6d, e, see Note 47).
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4.
Kymograph data is first processed in custom LAKEVIEW software (see Note 48), to produce an image of photon counts. From there, the image is quantified in FIJI, to plot the photon count versus time (Fig. 6f).
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5.
This plot is fitted using the single exponential algorithm in FIJI (results in Fig. 6f), which gives a half-life of observed fluorescence. Results for nucleosomes and those exposed to FACT and SSRP1 HMGB and SPT16 MD are compared (Fig. 6g).
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6.
Upon disruption, histones are not immediately lost to solution, a result seen previously for nucleosomes labeled on H3 and more generally across all of the octamer (see Note 50) [40]. Interestingly, both SSRP1 HMGB and SPT16 MD induce rapid loss of disrupted histones, with full FACT driving the fastest loss (see Note 51).
3.11 Survival Probability (SP) Across Cycles of Disruption and Release
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1.
In the FD experiments above, only the first extension release cycle is collected/analyzed. Yet subsequent cycles can also be collected and analyzed as above (Fig. 7a, b) (see Note 52).
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2.
The survival probability (Fig. 7c) is scaled to the number of inner wrap releases observed compared to the first cycle (see Note 53), and the rupture forces (Fig. 7d, see Note 54) and extensions (Fig. 7e, see Note 55) are analyzed as above.
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3.
Only full FACT, combining SPT16 MD and SSRP1 HMGB, restores the nucleosome after release, over as many as 30 cycles (see Note 56). Note that here, failure to observe any rewrap** indicates not only dimer loss but likely tetramer loss as well, though it should be noted that the tetramer may remain bound to the DNA but not able to wrap it (due to displacement of the of the DNA from the central dyad).
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4.
Decreases in the observed rupture forces and the opening length indicate some progressive weakening/loss of the histone/DNA interactions in the core holding the inner wrap.
3.12 Comparisons
The results of combined FD, CI, and SP experiments show that FACT both destabilizes the nucleosome and chaperones its reassembly after disruption (Fig. 8). Furthermore, destabilization is driven chiefly by two domains, SSRP1 HMGB and STP MD, though each acts distinctly. SPT16 MD stabilizes overall nucleosome interactions, via binding to both DNA and the octamer. SSRP1 HGMGB destabilizes the nucleosome, through binding to the bent DNA of the entry/exit points. Interestingly, yeast NHP6A is shown to have an even stronger effect compared to the human analogue. Though the binding of each subunit disrupts the nucleosome and causes the release of the outer 1/2 wraps of DNA, individually neither can reunite the disrupted DNA with the histones. Only full FACT can restore the nucleosome after disruption, increasing the kinetics of DNA-histone interactions and acting as a true chaperone. The combinations of FD, CI, and SP experiments here can characterize the activity of a wide range of nuclear chaperones and determine their role in transcription, replication, and repair.
4 Notes
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1.
The wavelength of this laser is specified to be in the near IR (850 nm), to avoid water absorption lines at longer wavelengths, which would generate local heating. Systems that use 1064 nm lasers for trap** must be carefully adjusted to minimize this effect.
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2.
A 45-degree angle minimizes protein/DNA aggregation that accumulates at the bead-tip interface.
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3.
Smaller beads provide better resolution in FD and CI experiments, though larger beads facilitate catching and allow higher trap** forces and require smaller corrections to the stiffness.
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4.
Full FACT lacks the SPT16 NTD and SSRP1 CTD, as these constructs are difficult to reliably purify without significant aggregation.
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5.
For the full protein, cell suspensions containing the two subdomains are combined at a 1:2 volume ratio (SPT16:SSRP1) just before lysis and purification. This ratio ensures proper stoichiometry of the SPT16-SSRP1 complex after purification using the N-terminal His6-MBP tag on SPT16.
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6.
Linker length determines the degree of interaction between adjacent nucleosomes. The 60 base pair linkers shown here lead to ~20 nm separation between adjacent nucleosomes, even at low external forces of ~1 pN. Thus, adjacent nucleosomes are expected to have minimal direct interaction.
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7.
Distinct labels are required on each terminus for constructs whose length is very small relative to the size of the beads used in the force disruption experiments. Generally, the construct length should be comparable to the diameter of the smallest bead, and here, the reconstituted array construct is ~1 micron at 1 pN stretching force, while the smallest bead is 2.11 microns in diameter.
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8.
Longer “handles” facilitate clearer confocal images, which may be otherwise complicated by bead autofluorescence. Longer “handles” also allow the same label to be used on each terminus. Here, the reconstituted array construct is ~3 microns long at 1 pN stretching force, while the beads are 1.76 microns in diameter.
-
9.
A mass ratio 1.2 octamers to DNA sites (excluding handles) results in arrays with nearly 12× octamers on the positioning sites while they are minimized on the handles.
-
10.
While smaller volumes require less sample, sealing the dialysis membrane over a smaller volume is difficult to do without also entrap** an air bubble that will inhibit dialysis.
-
11.
Dialysis membranes were chosen with a cutoff of 6–8 kDa and secured in place with an O-ring.
-
12.
Sample quality was checked in AFM images, where ~12× octamers could be counted.
-
13.
The experimental buffer used here includes 100 mM Na+, 10 mM HEPES at 7.5 pH at room temperature. Slightly lower salt buffer (50 mM Na+) is also used as the lower salt stabilizes the nucleosomes arrays. However, at very low salt buffer, the biotin–streptavidin link is more difficult to establish.
-
14.
For the experiments shown throughout this work, both commercial anti-digoxigeninated (A-D) beads from the LUMICKS company and in house beads are used. Custom A-D beads can be made by coating anti-digoxygenin to protein G labeled 2 micron diameter beads. We have found that the commercial beads are more uniform in size and have better tethers to the nucleosome arrays.
-
15.
Varying sizes of polystyrene beads are useful, though limited by the laser trap diameter. Larger bead sizes are easier to catch and offer more surface area for tethers but are more likely to simultaneously catch multiple arrays. It is also preferable that differently labeled beads be of easily distinguishable sizes. Furthermore, within each stock solution, the beads should not vary strongly (>10%) in diameter, to prevent error in either instrument calibration or stiffness as described.
-
16.
The reconstituted nucleosome array should be freshly diluted 1:10,000 in experimental buffer on every data collection state. Diluted nucleosomes last 5–6 h, after which they steadily degrade.
-
17.
It is helpful to flow in buffer in between cycles of catching fresh beads as the large concentration of nucleosome array constructs quickly saturate the flow cell. The constructs coat the perimeter of the micropipette tip and the sides of the flow cell.
-
18.
Detectors, running at 20 kHz, typically each measure 100 data points of the trap** laser deflection (linearly proportional to the trap** force). Data shown here is averaged at this rate.
-
19.
The rate of the force increase is the loading rate. Here, that rate is ~10 pN/s, in the middle of the useful range of this instrument and typical for instruments of this type.
-
20.
The time for a full cycle is 5–10 s, depending on the number of data points required to reach the fully disrupted state from the starting extension.
-
21.
The instrument is calibrated via overstretched long DNA (phage-λ, 48,500 base pairs), which provides a clear force plateau for calibration of 60–65 pN (including known variations due to experimental conditions).
-
22.
Measuring the trap stiffness corrects for the movement of the trapped bead with increasing force. To find this value, first a bead should be pulled on the micropipette tip. Then, the piezo moves the bead across the length of the empty optical trap. The stiffness correction is defined by the displacement of the bead as the trap** laser is deflected over the relevant trap** range. For the 3 micron diameter beads (in the center range of those used here), this stiffness was measured to be ~110 pN/μm.
-
23.
The enthalpic stiffness is a phenomenological addition, not found in the original solution.
-
24.
Variations due to construct length and solution conditions are well known [41, 42].
-
25.
The force extension data follows the eWLC model until a “rip” is observed, which presents as both a sudden increase in the extension coupled with a drop in the measured force (due to the finite instrument stiffness and averaging rate). Thus, simultaneous force and length increases (relative to the unwrapped state) are sought to discriminate unwrap** events from noise. Typical thresholds were 0.3 pN and 7 nm, and note the length threshold is significantly lower than the typical lengths measured (between 20 and 25 nm). Finally, as mentioned above, the instrument is averaging the data during collection.
-
26.
The change in extension is converted to the number of base pairs, using Eq. (1).
-
27.
In practice, the extension change between the force thresholds of 5 and 2 pN is found, corrected for the expected change due to Eq. (1) and converted to bases.
-
28.
An estimate of the small correction due to the finite instrument stiffness (Wstiffness) and representing the area during the drop in force as the extension increases is also added.
-
29.
The more robust methods of Crooks and Bennett are difficult to implement as closing events were difficult to reliably observe [43, 44]. Where data could be taken, preliminary calculations produced values that appeared to agree with the results shown here.
-
30.
Pmax is formally a probability density, with units of (pN−1).
-
31.
Fmax and Pmax are both a function of the loading rate.
-
32.
Values are determined at each value of A and averaged. The values should be similar across all A, though there are often variations due to experimental uncertainty, especially in Pmax.
-
33.
The shape factor, ν, is formally 1/2 or 2/3. The value of 1/2 is used here.
-
34.
Here, arrays are selected where the number of inner wrap releases (N) was nine or greater, though N > 10 was typical. For every concentration, at least two arrays where N = 12 was sought to avoid biasing the measured force toward higher values.
-
35.
The simple Hill Eq. relates the occupancy (Θ) at each binding site to the applied ligand concentration (c) through the equilibrium dissociation constant (KD),
$$ \Theta =\frac{1}{1+{\left(\frac{K_{\mathrm{D}}}{c}\right)}^H}. $$(8)
-
When KD is equal to c, the occupancy is 1/2. As the sites are assumed to be non-interacting, H is set to unity, which roughly agrees with previous affinity measurements [39]. However, in the case of NHP6A binding, a value of H = 2 fit the data best implying a small degree of cooperativity.
-
36.
While in principle all three parameters could be independently fit to the data, in practice, Fnucl is well characterized and thus is fixed.
-
37.
Errors in the total length will be largely determined by the error in the measuring the outer 1/2 release.
-
38.
Equation (1) is fitted to the extension data using a nonlinear χ2 Levenberg–Marquardt algorithm, and details of this analysis have been shown previously.
-
39.
The value of PDNA was measured here, and the value of 42 ± 2 pN agrees with previous work that measured variations for shorter lengths of DNA.
-
40.
Now PDNA is fixed and KD and Pprotein are determined by the fits. The Hill parameter (H) was fixed at one, though in the case of NHP6A, H = 1.5 gave the best fits and indicate weak cooperativity.
-
41.
Trap** laser power was reduced to ~200 mW, to minimize heating due to absorption by water at 1064 nm.
-
42.
Confocal laser power must be systematically reduced to minimize photobleaching while maximizing signal intensity.
-
43.
The free channel may be passivated with a sequential application of BSA and Pluronic. An alternative and simpler approach prerinses each channel with 0.2% Tween20.
-
44.
To minimize protein attachment (and dye contamination) of the flow cell chamber, arrays may be caught and then translated into a protein containing chamber previously passivated. Alternatively, arrays may be incubated with the protein, as this binding is stable. Protein exposed arrays are then translated into a buffer only chamber to minimize background.
-
45.
The range of the kymograph is shown in the red box on the image in the inset to Fig. 6a.
-
46.
The green blocks correspond to autofluorescence of the beads, also visible in the confocal images.
-
47.
Kymographs are also collected at 1 pN, to check the rate of photobleaching (kbleach). The measured rate is used to correct all subsequent measured release rates (krelease), according to [45],
$$ I={I}_{\mathrm{o}}\cdot {e}^{-{t}_{\mathrm{release}}\cdot {k}_{\mathrm{release}}}\cdot {e}^{-{t}_{\mathrm{bleach}}\cdot {k}_{\mathrm{bleach}}}\cdot $$(9) -
48.
A more robust technique combines an oxygen scavenger (GODCAT, glucose oxidase and catalase) with a triplet suppressor (Trolox) [46].
-
49.
As the relative intensity may be saturated or washed-out during image processing, image data is processed on the photon counts for this comparison.
-
50.
In this study, H3 is labeled with an anti-H3 antibody attached to a dye, and NHS ester labeling is used to label histones generally.
-
51.
Critically, the H2A are dye labeled here, and the loss in signal here indicates only dimer loss. The tetramers may be lost at the same time or may also be lost shortly after (see results below).
-
52.
The measured half-lives from the previous sections indicate that to forestall histone loss, the arrays should remain unwrapped for less than 1–2 s.
-
53.
Only arrays were N > 10 are extended/released over these experiments, to minimize any potential disruption from the tethering process.
-
54.
As the cycle number increases and the number of nucleosomes left on each array decreases, Eq. (2) predicts that the average disruption force should increase. Yet this is not observed, indicating some progressive destabilization relative to native nucleosomes.
-
55.
The observed unwrap** length appears nearly constant with increasing cycle number, in contrast to the observed force. While DNA appears to wrap up to the dimers, the rewrapped state is not fully stable compared to the non-disrupted nucleosome.
-
56.
Restoration was limited to 30 cycles as the observed release force gradually decreased with each cycle, eventually crossing below the low force limit of the threshold chosen here.
References
Luger K, Mader AW, Richmond RK et al (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389(6648):251–260
Luger K, Richmond TJ (1998) DNA binding within the nucleosome core. Curr Opin Struct Biol 8(1):33–40
Brower-Toland BD, Smith CL, Yeh RC et al (2002) Mechanical disruption of individual nucleosomes reveals a reversible multistage release of DNA. Proc Natl Acad Sci U S A 99(4):1960–1965
Formosa T (2008) FACT and the reorganized nucleosome. Mol BioSyst 4(11):1085–1093
Belotserkovskaya R, Oh S, Bondarenko VA et al (2003) FACT facilitates transcription-dependent nucleosome alteration. Science 301(5636):1090–1093
Reinberg D, Sims RJ 3rd (2006) de FACTo nucleosome dynamics. J Biol Chem 281(33):23297–23301
Singer RA, Johnston GC (2004) The FACT chromatin modulator: genetic and structure/function relationships. Biochem Cell Biol 82(4):419–427
Winkler DD, Luger K (2011) The histone chaperone FACT: structural insights and mechanisms for nucleosome reorganization. J Biol Chem 286(21):18369–18374
Chen P, Dong L, Hu M et al (2018) Functions of FACT in breaking the nucleosome and maintaining its integrity at the single-nucleosome level. Mol Cell 71(2):284–293. e284
Gurova K, Chang HW, Valieva ME et al (2018) Structure and function of the histone chaperone FACT - resolving FACTual issues. Biochimica et biophysica acta Gene regulatory mechanisms
Wang T, Liu Y, Edwards G et al (2018) The histone chaperone FACT modulates nucleosome structure by tethering its components. Life Sci Alliance 1(4):e201800107
Liu Y, Zhou K, Zhang N et al (2020) FACT caught in the act of manipulating the nucleosome. Nature 577(7790):426–431
Ehara H, Kujirai T, Shirouzu M et al (2022) Structural basis of nucleosome disassembly and reassembly by RNAPII elongation complex with FACT. Science 377(6611):eabp9466
McCullough LL, Connell Z, **n H et al (2018) Functional roles of the DNA-binding HMGB domain in the histone chaperone FACT in nucleosome reorganization. J Biol Chem 293(16):6121–6133
Valieva ME, Armeev GA, Kudryashova KS et al (2016) Large-scale ATP-independent nucleosome unfolding by a histone chaperone. Nat Struct Mol Biol 23(12):1111–1116
Stillman DJ (2010) Nhp6: a small but powerful effector of chromatin structure in Saccharomyces cerevisiae. Biochim Biophys Acta 1799(1–2):175–180
McCauley MJ, Huo R, Becker N et al (2019) Single and double box HMGB proteins differentially destabilize nucleosomes. Nucleic Acids Res 47(2):666–678
Bustamante CJ, Chemla YR, Liu S et al (2021) Optical tweezers in single-molecule biophysics. Nat Rev Methods Primers 1(1):25
Chaurasiya KR, Paramanathan T, McCauley MJ et al (2010) Biophysical characterization of DNA binding from single molecule force measurements. Phys Life Rev 7:299–341
Heller I, Hoekstra TP, King GA et al (2014) Optical tweezers analysis of DNA–protein complexes. Chem Rev 114(6):3087–3119
Neuman KC, Block SM (2004) Optical trap**. Rev Sci Inst 75(9):2787–2809
Brower-Toland B, Wang MD (2004) Use of optical trap** techniques to study single-nucleosome dynamics. Methods Enzymol 376:62–72
Sheinin MY, Li M, Soltani M et al (2013) Torque modulates nucleosome stability and facilitates H2A/H2B dimer loss. Nat Commun 4:2579
Meijering AEC, Sarlós K, Nielsen CF et al (2022) Nonlinear mechanics of human mitotic chromosomes. Nature 605(7910):545–550
Dyer PN, Edayathumangalam RS, White CL et al (2004) Reconstitution of nucleosome core particles from recombinant histones and DNA. Methods Enzymol 375:23–44
Luger K, Rechsteiner TJ, Richmond TJ (1999) Preparation of nucleosome core particle from recombinant histones. Methods Enzymol 304:3–19
Muthurajan UM, McBryant SJ, Lu X et al (2011) The linker region of macroH2A promotes self-association of nucleosomal arrays. J Biol Chem 286(27):23852–23864
Rogge RA, Kalashnikova AA, Muthurajan UM et al (2013) Assembly of nucleosomal arrays from recombinant core histones and nucleosome positioning DNA. J Vis Exp 79
Perkins TT (2014) Angstrom-precision optical traps and applications. Annu Rev Biophys 43(1):279–302
Chemla YR (2016) High-resolution, hybrid optical trap** methods, and their application to nucleic acid processing proteins. Biopolymers 105(10):704–714
Peters JP, Becker NA, Rueter EM et al (2011) Quantitative methods for measuring DNA flexibility in vitro and in vivo. Methods Enzymol 488:287–335
Odijk T (1995) Stiff chains and filaments under tension. Macromolecules 28:7016–7018
Bustamante C, Marko JF, Siggia ED et al (1994) Entropic elasticity of lambda-phage DNA. Science 265(5178):1599–1600
Smith SB, Cui YJ, Bustamante C (1996) Overstretching B-DNA: the elastic response of individual double-stranded and single-stranded DNA molecules. Science 271:795–799
McCauley MJ, Rueter EM, Rouzina I et al (2013) Single-molecule kinetics reveal microscopic mechanism by which High-Mobility Group B proteins alter DNA flexibility. Nucleic Acids Res 41(1):167–181
Jarzynski C (1997) Nonequilibrium equality for free energy differences. Phys Rev Lett 78(14):2690–2693
Dudko OK, Hummer G, Szabo A (2006) Intrinsic rates and activation free energies from single-molecule pulling experiments. Phys Rev Lett 96(10):108101
McCauley MJ, Rouzina I, Manthei KA et al (2015) Targeted binding of nucleocapsid protein transforms the folding landscape of HIV-1 TAR RNA. Proc Natl Acad Sci U S A 112(44):13555–13560
Winkler DD, Muthurajan UM, Hieb AR et al (2011) Histone chaperone FACT coordinates nucleosome interaction through multiple synergistic binding events. J Biol Chem 286(48):41883–41892
Spakman D, King GA, Peterman EJG et al (2020) Constructing arrays of nucleosome positioning sequences using Gibson Assembly for single-molecule studies. Sci Rep 10(1):9903
Seol Y, Li J, Nelson PC et al (2007) Elasticity of short DNA molecules: theory and experiment for contour lengths of 0.6–7 μm. Biophys J 93(12):4360–4373
Wenner JR, Williams MC, Rouzina I et al (2002) Salt dependence of the elasticity and overstretching transition of single DNA molecules. Biophys J 82:3160–3169
Crooks GE (2000) Path-ensemble averages in systems driven far from equilibrium. Phys Rev E 61(3):2361–2366
Bennett CH (1976) Efficient estimation of free energy differences from Monte Carlo data. J Comput Phys 22(2):245–268
Farge G, Laurens N, Broekmans OD et al (2012) Protein sliding and DNA denaturation are essential for DNA organization by human mitochondrial transcription factor A. Nat Commun 3(1):1013
Rasnik I, McKinney SA, Ha T (2006) Nonblinking and long-lasting single-molecule fluorescence imaging. Nat Methods 3(11):891–893
Davey CA, Sargent DF, Luger K et al (2002) Solvent mediated interactions in the structure of the nucleosome Core particle at 1.9Å resolution. J Mol Biol 319(5):1097–1113
Ohndorf U-M, Rould MA, He Q et al (1999) Basis for recognition of cisplatin-modified DNA by high-mobility-group proteins. Nature 399(6737):708–712
Masse JE, Wong B, Yen Y-M et al (2002) The S. cerevisiae architectural HMGB protein NHP6A complexed with DNA: DNA and protein conformational changes upon binding. J Mol Biol 323(2):263–284
McCauley MJ, Morse M, Becker N et al (2022) Human FACT subunits coordinate to catalyze both disassembly and reassembly of nucleosomes. Cell Reports 41(13):111858
Acknowledgments
This work was supported by grants NIH AI167700 and NSF MCB-1243883 to M.C.W.
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McCauley, M.J. et al. (2024). Quantifying ATP-Independent Nucleosome Chaperone Activity with Single-Molecule Methods. In: Heller, I., Dulin, D., Peterman, E.J. (eds) Single Molecule Analysis . Methods in Molecular Biology, vol 2694. Humana, New York, NY. https://doi.org/10.1007/978-1-0716-3377-9_2
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