Name: in situ nucleosome assembly for single-molecule correlative force and fluorescence microscopy
Uploaded: 2024-09-06T12:50:00.000+00:00
Duration: 5 min 58 s
Description: Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding ...

G. N. L. C. acknowledges support from the National Institute of Mental Health of the National Institutes of Health (NIH) under award number F31MH132306. S. L. is supported by the Robertson Foundation, the International Rett Syndrome Foundation, and the NIH (award number R01GM149862).

The details of the reagents and the equipment used in the study are listed in the Table of Materials.
1. Preparation of biotinylated DNA
<ol>
	<li>Prepare a 120 &#181;L volume reaction containing 20 &#181;g of &#955; DNA, 33 &#181;M of biotin-dCTP, 33 &#181;M of biotin-dUTP, 33 &#181;M of biotin-dATP, 33 &#181;M of dGTP, 10 units of Klenow enzyme, and 1x NEB Buffer 2. 
	NOTE: This procedure specifically utilizes linear double-stranded (ds) methylation-free bacteriophage &#955; genomic DNA (48,502 bps), which contains 12-nucleotide&#160;(nt) 5&#39; single-stranded (ss) DNA overhang23. However, the protocol can be adapted to make use of any length or sequence of linear dsDNA provided it contains at least several nts of ss 5&#39; overhangs, the length and nt sequence dictating the number of biotinylated nts installed on each end for downstream tethering between optical traps. Additionally, the reaction can be scaled down. For example, 5 &#181;g of DNA can be used in a 30 &#181;L volume reaction. Additionally, the DNA substrates generated in this way are not torsionally constrained. For protocols to create torsionally constrained DNA, please refer to the previously published reports24,25.</li>
	<li>Incubate the reaction at room temperature for 15 min.</li>
	<li>Add 10 mM of EDTA and incubate at 75&#160;&#176;C for 20 min.</li>
	<li>Perform ethanol precipitation to obtain the biotinylated DNA product.
	<ol>
		<li>Add 0.1x volume of 3 M sodium acetate and 3x volume of ice cold 100% ethanol. Keep at -20&#160;&#176;C for at least 1 h up to overnight. Centrifuge at 13,500 x g for 30 min at 4&#160;&#176;C.</li>
		<li>Carefully remove the supernatant without disturbing the pellet and add 1 mL of 75% ethanol. Centrifuge at 13,500 x g for 1 min at 4 &#176;C.</li>
		<li>Repeat the supernatant removal, ethanol addition, and centrifugation steps.</li>
		<li>Carefully remove all the supernatant and let the pellet air dry for 10 min. Dissolve the pellet in 100-200 &#181;L of TE buffer or ddH2O (can incubate at 37&#160;&#176;C to facilitate dissolving). Measure the concentration using a Nanodrop spectrophotometer.</li>
	</ol>
	</li>
</ol>
2. Preparation of channels in a flow cell
NOTE: This procedure is based on a commercially available microfluidic chip (see Table of Materials) but can be adapted to any smCFFM instrument with a multi-channel flow cell.
<ol>
	<li>Passivate channels (ch.) 4 and 5 (or any ch. that will contain proteins) (see schematic of channels in Figure 1A). Add 500 &#181;L of 0.1% BSA to each channel and flow at 0.8 bar (~0.45 &#181;L/s for tubing with a 0.010 inch inner diameter) for 8 min. Remove solution.</li>
	<li>Add 500 &#181;L of 0.05% Pluronic-F127 to each channel and flow at 0.8 bar for 8 min. Remove the solution and rinse out the syringes with 1x PBS. Add 2 mL of 1x PBS to each channel and flow at 0.8 barfor 27 min.</li>
	<li>Vortex the stock solution of streptavidin-coated beads (this procedure uses 3.13 &#181;m size beads) and add 2.15 &#181;L of the bead solution to 1 mL of 1x PBS. Add to ch. 1. 
	NOTE: The size and concentration of the streptavidin-coated beads can be adjusted according to user preference.</li>
	<li>Add 30-80 ng of biotinylated &#955; DNA to 1 mL of 1x PBS. Add to ch. 2. 
	NOTE: The concentration of biotinylated DNA can be adjusted to user preference. Greater amounts of DNA will increase the frequency of forming DNA tethers but also raise the frequency of tethering multiple molecules of DNA at once, which can add to the difficulty of the experiment.</li>
</ol>
3. Nap1-mediated in situ nucleosome formation
<ol>
	<li>Add 500 &#181;L of protein-free&#160;image buffer to ch. 3. 
	NOTE: The buffer composition in ch. 3 is flexible and should be appropriate for both nucleosomes and the specific protein(s) of interest. Optionally, users may choose to add oxygen scavenging systems for reduced photobleaching of fluorophores and/or competitor DNA to remove histone octamers that are not properly wrapped into nucleosomes.</li>
	<li>Add 2 nM S. cerevisiae Nap1 and 2-5 nM LD655-labeled H4 histone octamer (HO) to 500 &#181;L of 1x HR buffer18. Add to ch. 4. 
	NOTE: Histone octamers and Nap1 can be commercially purchased or prepared in-house (see Table of Materials). The number of nucleosomes to be formed along each tethered DNA depends on the octamer concentration as well as the time spent in this channel. The user&#39;s desired and approximate nucleosome density along each DNA molecule can be modulated by adjusting either or both of these parameters. Additionally, the user may choose to label histone octamers with their preferred fluorophore.</li>
	<li>Optional: Add chromatin-binding protein or other protein of interest to ch. 5 in an appropriate buffer.</li>
	<li>Flow all channels at 1.0 bar (~0.55 &#181;L/s in the current experimental setup) for 30 s to flush the flow cell channels with samples.</li>
	<li>Set the flow to 0.2 bar (~0.16 &#181;L/s in the current experimental setup). Move the optical traps (OTs) to ch. 1 and catch a suitable pair of streptavidin-coated beads.</li>
	<li>Move the OTs to ch. 3, conduct a force calibration for the bead pair, turn on the red confocal laser (or another laser line of choice), and calibrate the imaging area.</li>
	<li>Set the flow to 0.2 bar and move OTs to ch. 2. Catch biotinylated DNA.</li>
	<li>Move the OTs to ch. 3 and stop the flow. Increase the distance between the OTs to stretch the DNA and monitor the associated force-distance (FD) curve to confirm the presence of a single DNA tether (as opposed to multiple tethers). The curve should follow the worm-like-chain model for dsDNA at the given contour length26.</li>
	<li>Adjust the distance between the OTs so that the DNA tension reads approximately 1 pN. 
	NOTE: 1 pN of tension is low enough to allow nucleosomes to remain wrapped by DNA27 but is high enough to place the entire DNA molecule within the line scanning imaging axis.</li>
	<li>Turn on line scanning to start collecting a kymograph with the red laser on. 
	NOTE: Kymographs or 2D scans of tethered DNA can be obtained, as per user preference.</li>
	<li>Move the OTs to ch. 4 with the flow off. Facilitated by Nap1, HOs are loaded onto DNA and wrapped to form nucleosomes in real-time. Nucleosome loading is visualized as fluorescent trajectories appearing on the DNA within kymographs. Concurrently, nucleosome formation on the DNA can be monitored by the force increase as more HOs are loaded when the positions of OTs are held constant. 
	NOTE: Fluorescence excitation laser can be turned off during the nucleosome formation steps to avoid photobleaching of fluorophores. Additionally, non-specific binding of histone octamers to DNA wherein the octamers are not wrapped into nucleosomes could occur using this protocol. These binding events will not generate a &#34;rip&#34; when the DNA is stretched, as described in the Unwrapping Nucleosomes (step 4) protocol below, and can sometimes be washed off by a gentle buffer flow, particularly if the buffer contains free competitor DNA.</li>
	<li>When the approximate desired number of nucleosomes is formed, start collecting data.</li>
</ol>
4. Unwrapping nucleosomes
<ol>
	<li>Complete step 2 and step 3.</li>
	<li>After forming a DNA tether containing nucleosomes, move the OTs to ch. 3.</li>
	<li>Prepare for nucleosome unwrapping experiments by reducing the distance between OTs until the force is approximately zero. Zero the force.</li>
	<li>Begin moving the OT 1 away from the OT 2 at a constant speed of 0.1 &#181;m/s. On a kymograph, the bead in OT 1 will visually move farther from the bead in OT 2. On the FD curve, the force will rise as the distance increases. As nucleosomes unwrap, there will appear &#34;rips&#34; or transitions in the force (~8-37 pN) that signify the unraveling of the inner wrap of individual nucleosomes27,28,29. 
	NOTE: The force at which unwrapping occurs depends on the pulling rate, which is expected for non-equilibrium processes. Sometimes, multiple nucleosomes unwrap at once, producing a larger distance change in the transition on the FD curve. Additionally, the unraveling of the outer wrap of the nucleosome is not readily visible in this experimental setting (see the Discussion section).</li>
</ol>
5. Visualizing protein binding to chromatin
<ol>
	<li>Complete step 2 and step 3.</li>
	<li>Add 10 nM of Cy3-labeled linker histone H1.4 to 500 &#181;L of image buffer in ch. 5. 
	NOTE: Any protein of interest can be added to ch. 5 at a concentration of choice (typically below 100 nM for single-molecule visualization).</li>
	<li>After forming a DNA tether containing nucleosomes, turn on the green laser in addition to the red laser and move OTs to ch. 5 for real-time imaging of H1 binding to chromatin.</li>
	<li>Continue collecting kymographs or scans until sufficient statistics have been achieved (at least ~10-20 kymographs depending on the frequency of events and experimental difficulty).</li>
</ol>

In Situ Nucleosome Reconstitution for Single-Molecule Studies

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The authors declare no competing interests.

The described protocol provides several advantages for nucleosome reconstitution including minimizing reagents and time as well as enabling chaperone-dependent nucleosome formation along any (potentially native) DNA sequence. Moreover, the in situ method allows simpler experimental workflow and a convenient transition into single-molecule assays of nucleosome mechanics and protein-chromatin interaction. On the other hand, limitations of this approach include the inability to direct nucleosome positioning along the DNA (even with the incorporation of strong nucleosome positioning sequences) and precisely control the number of nucleosomes formed per DNA tether (although this can be empirically tuned by standardizing the experimental workflow). In effect, those who choose this method trade in precise control of nucleosome positioning and density for greater workflow ease and low reagent requirement, the choice of which depends on each specific experiment. It is also noted that the outer turn of DNA is not resolved in the current protocol due to the long length of the tether (~48.5 kbps) and buffer conditions32. To obtain information about the outer turn of DNA as well as inter-nucleosome stacking, the user may choose to shorten the DNA length and load a higher density of nucleosomes onto the DNA and/or use the salt dialysis method to place nucleosomes on strong positioning sequences that are separated by regular linker lengths. Finally, non-specific binding of histone octamers to DNA is likely to occur; therefore, it is important to consider both fluorescence and force signals when interpreting the results.
One critical aspect of the current protocol is ensuring the user has achieved a high fluorescent labeling rate for histone octamers (if visualization is required) so that most, if not all, nucleosomes can be visually detected. Another critical aspect is confirming the user has chosen an appropriate buffer for the imaging channel that is compatible with nucleosomes and any other proteins of interest. User-specific modifications to the protocol may include fluorophore conjugation to a different histone (see reports33,34 for examples using this method to form nucleosomes with labeled H3 or H2B), choosing an alternative fluorophore, incorporating DNA tethers of different sequences and lengths, adding an oxygen-scavenging system to extend the lifetime of fluorophores, choosing a different bead size, and utilizing an alternative DNA-bead conjugation system (e.g., digoxigenin-anti-digoxigenin antibody linkage).
In summary, the described method provides smCFFM with a convenient and versatile approach to generate nucleosome-containing tethers that can be easily employed to study a wide array of chromatin-focused questions in biology. Applicable research areas include chromatin mechanics as regulated by histone variants and post-translational modifications, biophysical engagement and kinetics of chromatin-binding proteins, and nucleosome-directed higher-order assembly processes such as biomolecular condensation35,36,37,38.

The primary packaging unit of eukaryotic chromatin is the nucleosome, in which ~147 base pairs (bps) of DNA are wrapped around an octamer of core histones proteins1,2. In addition to genome packaging, the nucleosome architecture serves as another rich layer of biophysical regulation that can be harnessed by chromatin-binding proteins when performing their various functions3,4. Experimentally accessing and measuring the physical characteristics of nucleosomes has been technically challenging since these units perform at minuscule scales (e.g., nanometer lengths, piconewton forces), and thus, their probing requires sufficient sensitivity and precision to meaningfully inform function. Moreover, chromatin-binding proteins often engage with their substrates transiently, and most ensemble approaches lack appropriate temporal resolution to inform the kinetics of these interactions5. Fortunately, the advent of single-molecule techniques has made it possible to visualize and manipulate individual proteins and their interactions in real-time, revealing mechanistic information about these molecular events occurring at the nanoscale6. In particular, single-molecule correlative force and fluorescence microscopy (smCFFM) harnesses both high-resolution fluorescence detection and force manipulation tools to simultaneously resolve the mechanics, composition, and coordination of chromatin and chromatin-protein complexes7,8.
In smCFFM that specifically employs &#34;optical tweezers&#34; as the force manipulation method, a tightly focused laser is used to trap a dielectric object, typically a micron-sized plastic bead, in three dimensions9. A biopolymer such as a piece of DNA can be conjugated to the bead (via, e.g., streptavidin-biotin, digoxigenin-anti-digoxigenin antibody linkage), and the user can both apply a calibrated force and measure the force/displacement generated by the system10. The added fluorescence module enables simultaneous multicolor fluorescence imaging to track protein composition and behavior.
The mainstream method for reconstituting nucleosome templates for smCFFM involves assembling nucleosomes on custom DNA templates by salt dialysis11,12,13. In this procedure, purified histone octamers are incubated with DNA templates in a high-salt buffer (~1 M sodium chloride), and as the salt is slowly dialyzed away, nucleosomes spontaneously assemble on the DNA in a sequence-dependent positional manner14,15. As such, it is customary that strong nucleosome positioning sequences (e.g., Widom 60116, X. laevis 5S rDNA17) are incorporated within the DNA templates to generate custom nucleosome arrays. Although this well-established method offers precise control over the placement and number of nucleosomes across DNA, it suffers from several disadvantages: (1) incorporation of strong nucleosome positioning DNA sequences may generate artificially stable nucleosomes that bias the activity of chromatin-binding proteins, (2) the process of salt dialysis for nucleosome formation requires high amounts of DNA and octamers, and (3) the procedure takes one to several days of work as well as a considerable effort to titrate the correct DNA to octamer ratio for optimal nucleosome array density.
In this manuscript, we describe an alternative method to form nucleosomes along DNA in situ within a single-molecule correlative force and fluorescence microscope using recombinant histone chaperone Nap1, which resembles the physiological pathway of nucleosome formation in vivo18,19,20,21,22. This method allows users to assemble nucleosomes on non-specific DNA sequences, easily adjust nucleosome density, and utilize significantly fewer amounts of reagents, all within minutes. Additionally, the formation of nucleosomal DNA tethers in situ enables simpler experimental workflow and the convenience of built-in single-molecule visualization and manipulation. Thus, when uniform and specific nucleosome positioning is not a necessity, this protocol offers a useful option for the investigation of nucleosome mechanics or protein behavior on chromatin, for which we also describe example assays.

<table><tbody><tr><td>1x HR buffer</td><td>N/A</td><td>N/A</td><td>30 mM&amp;nbsp;tris&amp;nbsp;acetate pH 7.5, 20 mM magnesium acetate, 50 mM potassium chloride, 0.1 mg/mL BSA&amp;nbsp;</td></tr><tr><td>1x PBS (Phosphate-buffered saline)</td><td>N/A</td><td>N/A</td><td>137 mM sodium chloride, 2.7 mM potassium chloride, 10 mM sodium phosphate dibasic, 1.8 mM potassium phophate monobasic&amp;nbsp;</td></tr><tr><td>Acetic acid, glacial</td><td>Millipore Sigma</td><td>AX0074-6</td><td>Use to make tris acetate</td></tr><tr><td>Biotin-11-dUTP</td><td>Jena Bioscience</td><td>NU-803-BIOX-S</td><td></td></tr><tr><td>Biotin-14-dATP</td><td>Jena Bioscience</td><td>NU-835-BIO14-S</td><td></td></tr><tr><td>Biotin-14-dCTP</td><td>Jena Bioscience</td><td>NU-956-BIO14-S</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Millipore Sigma</td><td>A9418-50G</td><td>Dissolve in H2O and run through 0.22 um filter</td></tr><tr><td>Cy3 Maleimide Mono-Reactive Dye</td><td>Cytiva</td><td>PA23031</td><td>Maleimide functionalized Cy3 fluorophore</td></tr><tr><td>dGTP</td><td>New England Biolabs</td><td>N0442S</td><td></td></tr><tr><td>Eppendorf Centrifuge 5425 R</td><td>Fisher Scientific</td><td>05-414-051</td><td>Benchtop centrifuge with cooling</td></tr><tr><td>Ethyl alcohol, Pure</td><td>Millipore Sigma</td><td>459844</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Millipore Sigma</td><td>E9884</td><td>Dissolve in H2O to 0.5 M</td></tr><tr><td>Human histone octamer (H4, L50C; H2A, K119C)&amp;nbsp;</td><td>N/A</td><td>N/A</td><td>Recombinant histone proteins and those harboring labeling mutations were purified in-house as described previously (see refs. 33, 34, 35, 36). Briefly, recombinant histones were expressed in BL21 (De3) pLySS cells (Promega). Inclusion bodies were isolated after sonication, and histones were extracted under denaturing conditions. Histones were dialyzed into buffer A (7 M urea, 10 mM tris hydrochloride pH 8.0, 100 mM sodium chloride, 1 mM EDTA, and 5 mM 2-mercaptoethanol), and the solution was added to a gravity column loaded with Q Sepharose Fast Flow (Cytiva). The flow through was then added to a gravity column loaded SP Sepharose Fast Flow (Cytiva), and the histones were eluted from the column by adding buffer A supplemented with 600 mM sodium chloride. Histones harboring labeling mutations (H4, L50C; H2A, K119C) were purified and then conjugated to the desired fluorophore via maleimide-functionalized dyes (Cytiva, Lumidyne) using a 20:1 dye-to-protein molar ratio (see refs. 33, 34). Histone octamers were assembled by adding an equal molar ratio of each wild-type or fluorophore-labeled histone under denaturing conditions, dialyzed into a high-salt buffer containing 2 M sodium chloride, and then purified by size exclusion chromatography as described previously (see refs. 36, 37). Alternatively, individual histone proteins and ready-made histone octamers can be purchased commercially (e.g., Epicypher).&amp;nbsp;</td></tr><tr><td>Image buffer</td><td>N/A</td><td>N/A</td><td>20 mM tris hydrochloride pH 8.0, 100 mM sodium chloride</td></tr><tr><td>Klenow Fragment (3&#039; to 5&#039; exo-)</td><td>New England Biolabs</td><td>M0212S</td><td></td></tr><tr><td>Lambda DNA (dam-, dcm-)</td><td>Thermo Fisher Scientific</td><td>SD0021</td><td>Methylation-free &amp;lambda; DNA</td></tr><tr><td>LD655-MAL</td><td>Lumidyne Technologies</td><td>9</td><td>Maleimide functionalized LD655 fluorophore</td></tr><tr><td>Linker histone H1.4 (A4C)</td><td>N/A</td><td>N/A</td><td>Purified recombinant protein made in-house (see ref. 38)</td></tr><tr><td>LUMICKS C-Trap Dymo</td><td>LUMICKS</td><td>N/A</td><td>Dual-trap configuration; standard materials for instrument provided by manufacturer</td></tr><tr><td>Magnesium acetate solution</td><td>Millipore Sigma</td><td>63052-100ML</td><td>Use to make HR buffer</td></tr><tr><td>NEBuffer 2</td><td>New England Biolabs</td><td>B7002S</td><td>Included with Klenow Fragment kit</td></tr><tr><td>Pluronic F-127</td><td>Millipore Sigma</td><td>P2443-250G</td><td>Dissolve in H2O and run through 0.22 um filter</td></tr><tr><td>Potassium chloride</td><td>Millipore Sigma</td><td>P3911</td><td>Use to make PBS and HR buffer</td></tr><tr><td>Potassium phosphate monobasic</td><td>Millipore Sigma</td><td>P0662</td><td>Use to make PBS</td></tr><tr><td>&lt;em&gt;S. cerevisiae&lt;/em&gt; Nap1</td><td>N/A</td><td>N/A</td><td>Nap1 was purified in-house as previously described (see refs. 33, 34). Alternatively, Nap1 can be purchased commercially (e.g., Active Motif).</td></tr><tr><td>Sodium acetate</td><td>Millipore Sigma</td><td>241245</td><td>Dissolve in H2O to 3 M</td></tr><tr><td>Sodium chloride</td><td>Millipore Sigma</td><td>S9888</td><td>Use to make PBS and image buffer</td></tr><tr><td>Sodium phosphate dibasic</td><td>Millipore Sigma</td><td>S9763</td><td>Use to make PBS</td></tr><tr><td>SPHERO Biotin Coated Particles (3.0-3.4 &amp;micro;m)</td><td>Spherotech</td><td>TP-30-5</td><td></td></tr><tr><td>Thermo Scientific NanoDrop 2000/2000c Spectrophotometer</td><td>Fisher Scientific</td><td>ND2000</td><td>NanoDrop Spectrophotometer</td></tr><tr><td>Tris Base</td><td>Fisher Scientific</td><td>BP152-500</td><td>Dissolve in H2O and adjust to appropriate pH; use to make image buffer and tris acetate</td></tr></tbody></table>

in situ nucleosome assembly for single-molecule correlative force and fluorescence microscopy

Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding their biophysical properties and interactions with chromatin-binding proteins. Nucleosome reconstitution on DNA for these studies typically involves a salt dialysis procedure that provides precise control over the placement and number of nucleosomes formed along a DNA tether. However, this protocol is time-consuming and requires a substantial amount of DNA and histone octamers as inputs. To offer an alternative strategy, an in situ nucleosome reconstitution method for single-molecule force and fluorescence microscopy that utilizes the histone chaperone Nap1 is described. This method enables users to assemble nucleosomes on any DNA template without the need for strong nucleosome positioning sequences, adjust nucleosome density on demand, and use fewer reagents. In situ nucleosome formation occurs within seconds, offering a simpler experimental workflow and a convenient transition into single-molecule measurements. Examples of two downstream assays for probing nucleosome mechanics and visualizing the behavior of individual proteins on chromatin are further described.

Using the setup described in step 3 (Figure 1A), nucleosome formation along the DNA tether was visualized as the appearance of red fluorescent foci on a 2D scan (Figures 1B, left) or trajectories over time on a kymograph (Figure 1B, right). Properly wrapped nucleosomes yielded fluorescence trajectories that were stationary over time within the diffraction limit of confocal detection (~300 nm). Notably, multiple nucleosomes formed near each other under the diffraction limit cannot be spatially resolved and appear as brighter foci or trajectories. Nevertheless, the photobleaching steps of individual nucleosomal trajectories were analyzed and frequently showed 1-step (Figure 1C) or 2-step (Figure 1D) photobleaching events, which indicates that foci mostly contained mononucleosomes given the approximately 70% labeling rate of the H4 protein in this sample. Moreover, based on the average photon count for a single LD655 fluorophore in this experimental condition, the number of labeled H4 copies in each nucleosome trajectory was estimated and showed that the nucleosome loci predominately (&#62;75%) contained 1 or 2 labeled H2A copies (Figure 1E), again consistent with the notion that they were primarily loaded as spatially separated mononucleosomes. The other brighter foci contained more than 3 copies of labeled H4, likely representing multiple nucleosomes within the diffraction limit.
Nucleosome formation was also monitored by an increase in the tension along the tether when the positions of the OTs remained constant as more DNA was wrapped around histone octamers (Figure 1B, bottom). In the absence of Nap1, histone octamers can still bind DNA but remain mostly unwrapped, as the associated tension when the OT positions were held remained constant (Figure 1F). It is possible that non-specific binding of histone octamers to DNA occurs even in the presence of Nap1. In another experiment, the same protocol to form nucleosomes containing Cy3-labeled H2A was performed, which yielded similar results as those performed with LD655-labeled H4 (Figure 1G), indicating the assembly of intact octamers into nucleosomes.
Using the setup described in step 4 (Figure 2A), nucleosome unwrapping was visualized over time on a kymograph as the tether distance increased (Figure 2B). It was previously reported that histone octamers often remain bound to the DNA after being unwrapped30,31, and therefore, fluorescent trajectories often persist even at high tensions. Nucleosome unwrapping was also monitored on the associated FD curve (Figure 2C). Quantifiable features of the FD curve include the force at which nucleosome(s) unwrap (see &#34;Transition F&#34; in Figure 2C inset) as well as the length of DNA that was released from the histone octamer (see &#34;&#916;L&#34; in Figure 2C inset), which exists in multiples of ~24 nm, consistent with independent unwrapping of the inner turns of DNA of individual nucleosomes27,30. These parameters can be used to estimate the number of nucleosomes formed on each tether or for additional downstream analysis of nucleosome mechanics. As such, the number of inner-turn ruptures obtained from mechanical pulling experiments was compared to the estimated number of labeled H4 copies at each nucleosome trajectory based on the monomer intensity, which showed that the numbers are largely consistent (examples shown in Figures 2D-G). However, it was noted that they do not perfectly match due to the ~70% labeled efficiency of the H4 protein in the sample and any residual Nap1-independent histone binding to DNA, as shown in Figure 1F. Finally, as expected, stretching Nap1-bound bare DNA tethers (without histones) do not generate force-induced transitions (Figure 2H), indicating that Nap1 alone does not significantly distort DNA.
Employing the setup described in step 5 (Figure 3A), the binding of linker histone H1 to chromatin was visualized as the appearance of green fluorescent foci along the chromatin tether (Figure 3B). Notably, H1-bound nucleosomes generated dual-color trajectories (see white arrows in Figure 3B) that appeared stationary over time. In addition, diffusing H1 trajectories across nucleosome-free DNA regions were observed (see green stars in Figure 3B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66579/66579fig01.jpg" /> 
Figure 1: Nap1-mediated in situ nucleosome formation. (A) Schematic of the experimental setup described in the Nap1-mediated in situ nucleosome formation protocol (step 3). A pair of beads were optically trapped in ch. 1, and a single &#955; DNA molecule was tethered between the beads through biotin-streptavidin linkage in ch. 2. The tether was then moved to ch. 4, containing LD655-labeled histone octamers (red) and Nap1 for real-time formation of nucleosomes across the DNA.&#160;(B) Representative 2D scan (left) and kymograph (right) of an LD655-H4-labeled nucleosome-containing DNA tether. The associated force reading (bottom) shows an increase in force immediately as the tether is moved to ch. 4 when the OT positions are held constant, indicating nucleosome wrapping. Brighter trajectories signify multiple adjacent nucleosomes located within the diffraction limit of ~300 nm. (C) Example kymograph showing a 1-step photobleaching event (white arrow) of a nucleosome containing LD655-H4. (D) Example kymograph showing a 2-step photobleaching event (white arrows) of a nucleosome containing LD655-H4. (E) Relative frequency of the number of LD655-labeled H4 per trajectory estimated from their fluorescence intensity. (F) Representative kymograph of a tether bound with histone octamers in the absence of Nap1. The associated force reading (bottom) remains unchanged even as histone octamers bind the DNA. (G) Representative kymograph of a Cy3-H2A-labeled nucleosome-containing DNA tether. The associated force reading (bottom) shows an increase in force immediately as the tether is moved to ch. 4 (containing octamers and Nap1), as nucleosomes are formed when the OT positions are held constant. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66579/66579fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66579/66579fig02.jpg" /> 
Figure 2: Nucleosome unwrapping assay. (A) Schematic of the experimental setup described in the unwrapping nucleosomes protocol (step 4). The DNA tether containing nucleosomes was moved to ch. 3 containing a protein-free image buffer where the inter-bead distance was increased and the nucleosomes were unwrapped. (B) Representative kymograph of a nucleosome-containing DNA tether pulled to high forces by gradually increasing the inter-bead distance. The decrease in fluorescence intensity toward the bottom bead was likely due to the DNA tilting slightly out of line scanning focus. (C) The associated FD curve showing force-induced transitions of inner turn unwrapping occurring between 8-37 pN. The inset shows a zoom-in view of an example transition for which the distance unwrapped (&#916;L = ~24 nm for the inner turn of DNA of individual nucleosomes) and the transition force (Transition F) can be recorded and further analyzed. For this tether, 26 nucleosomes were unwrapped, and the tether ruptured at ~65 pN, as denoted by the white dotted line in the kymograph. (D,E) Example LD655-H4 nucleosome-containing tether pulled to high forces where the number of H4 per trajectory is estimated via fluorescence intensity (indicated in the kymograph, D) and the number of wrapped nucleosomes is estimated via the size of force-induced distance change (indicated in the associated force-distance curve, E). (F,G) Another example as in (D,E). (H) DNA tethers (n = 5) pulled to high forces after being incubated in a channel containing Nap1 only. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66579/66579fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66579/66579fig03.jpg" /> 
Figure 3: Visualizing protein-chromatin interaction. (A) Schematic of the experimental setup described in the visualizing protein binding to chromatin protocol (step 5). The DNA tether containing nucleosomes was moved to ch. 5, containing 10 nM of Cy3-labeled linker histone H1 (green). (B) Representative two-color kymograph of H1 binding to a nucleosome-containing DNA tether. H1-nucleosome colocalization events are denoted with white arrows. Unbound nucleosome trajectory is denoted with the magenta arrow. H1 molecules diffusing on bare DNA are denoted with green stars. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66579/66579fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Efficient Nucleosome Reconstitution for Single-Molecule Techniques

This article presents a detailed experimental procedure for reconstituting nucleosome-containing DNA tethers for single-molecule correlative force and fluorescence microscopy. It further describes several downstream experiments that can be conducted to visualize the binding behavior of chromatin-interacting proteins and analyze changes in the physical properties of nucleosomes.

In Situ Nucleosome Assembly for Single-Molecule Correlative Force and Fluorescence Microscopy

Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding ...

in-situ-nucleosome-assembly-for-single-molecule-correlative-force

Nanyang Technological University

Research

JoVE Journal

Biochemistry

974 Views.  The Rockefeller University. Technieken met één molecuul zijn krachtige hulpmiddelen om de mechanica, coördinatie en samenstelling van chromatinesystemen te bestuderen. En daarom zijn onderzoekers altijd op zoek naar betere methoden om nucleosoomsubstraten te genereren. Hier beschrijven we een protocol om nucleosomen te vormen in DNA in C2 in een enkele molecuul correlatieve kracht en fluorescentiemicroscoop.Doorgaans worden nucleosoomsubstraten gemaakt door nucleosomen te asse...

Video: Auteur in de schijnwerpers: Efficiënte nucleosoomreconstitutie voor technieken met één molecuul

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

Biology

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Developmental Biology

Assembly of Nucleosomal Arrays from Recombinant Core Histones and Nucleosome Positioning DNA

Core histone octamers that are repetitively spaced along a DNA molecule are called nucleosomal arrays. Nucleosomal arrays are obtained in one of two ways: purification from in vivo sources, or reconstitution in vitro from recombinant core histones and tandemly repeated nucleosome positioning DNA. The latter method has the benefit of allowing for the assembly of a more compositionally uniform and precisely positioned nucleosomal array. Sedimentation velocity experiments in the analytical ultracentrifuge yield information about the size and shape of macromolecules by analyzing the rate at which they migrate through solution under centrifugal force. This technique, along with atomic force microscopy, can be used for quality control, ensuring that the majority of DNA templates are saturated with nucleosomes after reconstitution. Here we describe the protocols necessary to reconstitute milligram quantities of length and compositionally defined nucleosomal arrays suitable for biochemical and biophysical studies of chromatin structure and function.

A method is presented for the reconstitution of model nucleosomal arrays from recombinant core histones and tandemly repeated nucleosome positioning DNA. We also describe how sedimentation velocity experiments in the analytical ultracentrifuge, and atomic force microscopy (AFM) are used to monitor the extent of nucleosomal array saturation after reconstitution.

Core histone octamers that are repetitively spaced along a DNA molecule  are called nucleosomal arrays. Nucleosomal arrays are obtained in one of two  ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. This approach is particularly effective at mapping highly repetitive sequences to heterochromatic regions, where computational approaches face prohibitive challenges. Here we describe a streamlined protocol for DNA ISH that circumvents formamide washes that are standard steps in other DNA ISH protocols. Our protocol is optimized for hybridization with short single strand DNA probes that carry fluorescent dyes, which effectively mark repetitive DNA sequences within heterochromatic chromosomal regions across a number of different insect tissue types. However, applications may be extended to use with larger probes and visualization of single copy (non-repetitive) DNA sequences. We demonstrate this method by mapping several different repetitive sequences to squashed chromosomes from Drosophila melanogaster neural cells and Nasonia vitripennis spermatocytes. We show hybridization patterns for both small, commercially synthesized probes and for a larger probe for comparison. This procedure uses simple laboratory supplies and reagents, and is ideal for investigators who have little experience with performing DNA ISH.

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

Here, we present a simple method for performing fluorescence DNA in situ hybridization (DNA ISH) to visualize repetitive heterochromatic sequences on slide-mounted chromosomes. The method requires minimal reagents and it is versatile for use with short or long probes, different tissues, and detection with fluorescence or non-fluorescence-based signals.

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. This approach is particularly ...

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as specialized arrays during mitosis to regulate chromosome segregation. Proteins that interact with microtubules include motors such as kinesins and dynein, which can generate active forces and directional motion, as well as non-motor proteins that crosslink filaments into higher-order networks or regulate filament dynamics. To date, biophysical studies of microtubule-associated proteins have overwhelmingly focused on the role of single motor proteins needed for vesicle transport, and significant progress has been made in elucidating the force-generating properties and mechanochemical regulation of kinesins and dyneins. However, for processes in which microtubules act both as cargo and track, such as during filament sliding within the mitotic spindle, much less is understood about the biophysical regulation of ensembles of the crosslinking proteins involved. Here, we detail our methodology for directly probing force generation and response within crosslinked microtubule minimal networks reconstituted from purified microtubules and mitotic proteins. Microtubule pairs are crosslinked by proteins of interest, one microtubule is immobilized to a microscope coverslip, and the second microtubule is manipulated by an optical trap. Simultaneous total internal reflection fluorescence microscopy allows for multichannel visualization of all the components of this microtubule network as the filaments slide apart to generate force. We also demonstrate how these techniques can be used to probe pushing forces exerted by kinesin-5 ensembles and how viscous braking forces arise between sliding microtubule pairs crosslinked by the mitotic MAP PRC1. These assays provide insights into the mechanisms of spindle assembly and function and can be more broadly adapted to study dense microtubule network mechanics in diverse contexts, such as the axon and dendrites of neurons and polar epithelial cells.

Here, we present a protocol for reconstituting microtubule bundles in vitro and directly quantifying the forces exerted within them using simultaneous optical trapping and total internal reflection fluorescence microscopy. This assay allows for nanoscale-level measurement of the forces and displacements generated by protein ensembles within active microtubule networks.

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as ...

High-Speed Time-Lapse Atomic Force Microscopy to Capture Nucleosome Dynamics

Source: Stumme-Diers, M. P., et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/58820">Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging.</a>&#160;J. Vis. Exp.&#160;(2019)
In this video, we demonstrate high-speed time-lapse atomic force microscopy, AFM, imaging techniques to study the dynamics of nucleosome complexes. As the AFM tip scans across the surface of the nucleosome, it detects alterations in the nucleosome height and records the DNA unwrapping over time.

Source: Stumme-Diers, M. P., et al., Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging.&#160;J. Vis. ...

JoVE Encyclopedia of Experiments

Biological Techniques

Microscopy Techniques

Single-Molecule Imaging to Visualize Fusion Protein Condensates Assembly on DNA Curtains

To image the EWS-FLI1 condensation formation on DNA curtains, open the imaging software and find and mark the positions of the nine zig-zag patterns under bright-field. Then, turn on the flow at 0.2 milliliters per minute to stain the DNA with double-stranded DNA dye for 10 minutes.
Next, dilute the mCherry-EWS-FLI1 protein with the imaging buffer at a concentration of 100 nanomoles and 100 microliters. Then, load the protein sample through the valve with a 100-microliter glass syringe, and change the flow rate to 0.4 milliliters per minute.
After turning on the 488-nanometer laser, pre-scan each region to check the DNA distribution state, and select the region in which the DNA molecules distribute evenly. Then, set the laser power to 10% for the 488-nanometer laser and 20% for the 561-nanometer laser, and using the power meter, measure the real laser power near the prism.
Start acquiring images at two-second intervals with both 488 and 561-nanometer lasers simultaneously. Then, change the valve from the manual mode to injection mode to let the imaging buffer flush the protein sample into the flow cell after 60 seconds.
To remove the free EWS-FLI1, keep washing the flow cell with the imaging buffer for five minutes with only the 561-nanometer laser switched on. Then, stop the flow and incubate at 37 degrees Celsius for 10 minutes.
After 10 minutes, turn on the flow at 0.4 milliliters per minute to let the DNA extend, and acquire images at two-second intervals between different frames to obtain high-throughput data of EWS-FLI1 condensate formation.

In Situ Nucleosome Assembly for Single-Molecule Correlative Force and Fluorescence Microscopy

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