Name: Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions (Video) | JoVE Journal
Uploaded: 2014-08-27T14:00:00.000+00:00
Duration: 14 min 43 s
Description: Learn to combine double optical tweezers and fluorescence microscopy for studying protein-DNA interactions at the single-molecule level. This method enables precise mechanical control and visualization of dynamic molecular binding events.

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5 minutes

We thank Gijs Wuite, Erwin J.G.&#160;Peterman, and Peter Gross for help with the microfluidics and Alessia Tempestini for help with sample preparation. This research was funded by the European Union Seventh Framework Programme (FP7/2007-2013) under grant agreement n&#176; 284464 and from the Italian Ministry for Education, University and Research FIRB 2011 RBAP11X42L006, Futuro in Ricerca 2013 RBFR13V4M2, and in the framework of the Flagship Project NANOMAX.

1. Optical Tweezers Setup with Nanometer Stability
The experimental setup must provide two optical tweezers with pointing stability at the nanometer level and intensity fluctuations of the trapping laser below 1%. Combination of these conditions will assure nanometer stability of the dumbbell under typical tension (1 pN - few tens of pN), trap stiffness (0.1 pN/nm) and measurement bandwidth (image acquisition rate 20 sec-1). A scheme of the experimental setup is depicted in Figure&#160;1.
<ol>
	<li>Optical tweezers design and construction15,17:
	<ol>
		<li>Mount the experimental apparatus on an optical table with active isolators to reduce mechanical vibrations. Mount the microscope structure on elastomeric isolators to absorb acoustic noise and mechanical resonances of the microscope structure.</li>
		<li>Insert an optical isolator in the path of the trapping laser, near the laser source, to reduce random amplitude fluctuations due to optical feedback.</li>
		<li>Reduce the path length of the trapping laser as much as possible and enclose the whole path in a sealed box to reduce laser pointing fluctuations due to air currents and turbulence.</li>
		<li>Create double optical tweezers by splitting a single near-infrared laser source (Nd:YAG laser, 1,064 nm wavelength) into two beams with orthogonal polarizations. Do not use time-shared traps because they cause oscillation of the dumbbell under tension12.</li>
		<li>Use acousto-optic deflectors (AODs) driven by Direct Digital Synthesizers (DDSs) to allow fine movements of (at least) one trap and precise regulation of DNA tension. DDSs allow a better trap stability than Voltage-Controlled Oscillators (VCOs).</li>
		<li>Insert two quadrant detector photodiodes (QDPs) in the back focal plane of the condenser to detect the position of the trapped beads with sub-nm accuracy.</li>
	</ol>
	</li>
	<li>Check that intensity fluctuations of the trapping laser at the microscope entrance are below 1% by using a photodiode with a bandwidth larger than the rate of image acquisition.</li>
	<li>Check pointing stability of the two trapping lasers:
	<ol>
		<li>Prepare silica beads in phosphate buffer: Dilute 20 &#956;l of silica beads (1.54 &#956;m, 10% solids) into 1 ml of acetone, sonicate 30 sec, vortex briefly, and centrifuge 2 min at 19,000 x g. Resuspend in 1 ml acetone and repeat wash. Resuspend in 1 ml of 50 mM phosphate buffer, wash 2 times, and finally resuspend in 100 &#956;l of 50 mM phosphate buffer.</li>
		<li>Calibrate optical tweezers with silica beads: prepare a flow cell by attaching a coverslip to a microscope slide using double-sticky tape (about 60 &#956;m thick). Flow 1 mg/ml BSA and wait 3 min. Flow silica beads diluted 1/1,000 in phosphate buffer and seal the flow chamber with silicon grease. Trap one bead in each trap and calibrate the traps using the power spectrum method 18.</li>
		<li>Prepare silica beads in pentyl acetate and nitrocellulose: Dilute 20 &#956;l of silica beads (1.54 &#956;m, 10% solids) into 1 ml of acetone, sonicate 30 sec, vortex briefly, and centrifuge 2 min at 19,000 x g. Resuspend in 1 ml acetone and repeat wash. Resuspend in 1 ml of pentyl acetate and wash 2 times. Finally resuspend them in pentyl acetate with 0.1% nitrocellulose dissolved in pentyl acetate.</li>
		<li>Spread 2 &#956;l of 1.54 &#956;m diameter silica beads dissolved in pentyl acetate + 0.1% nitrocellulose onto the surface of a coverslip (24 x 24 mm) using a second coverslip (24 x 60 mm). Attach the coverslip to a microscope slide by using two pieces of double-sided tape to create a flow cell. Fill the flow-cell with 50 mM phosphate buffer and seal it with silicon grease.</li>
		<li>Image one silica bead in brightfield microscopy at 2,000X magnification. The field of view should be 7 x 5 &#956;m acquired at 768 x 576 pixel, so that the bead image has a diameter of 190 pixels. Compensate thermal drifts with a feedback software that, as in ref. 17, calculates x-y position from the image centroid and z from the diffraction rings and corrects drifts moving a piezo stage with nm-accuracy or better.</li>
		<li>Overlap the center of left trap with the bead center (the x-y signal levels from the QDP must be the same measured during the calibration) and measure the position noise and its standard deviation from the QDP. Repeat the measurement for the right trap.</li>
	</ol>
	</li>
</ol>
2. Single Molecule Localization with Nanometer Accuracy
<ol>
	<li>Fluorescence microscopy design and construction15,19
	<ol>
		<li>Use a high quantum efficiency and electron-multiplied CCD to reach the high signal-to-noise ratios necessary for nanometer localization accuracy.</li>
		<li>Choose optics to get an image pixel size ~80 nm (for example, an image magnification ~200X with a camera pixel size of 16 &#956;m). This is sufficiently small to neglect localization error due to pixelation8, but sufficiently large to collect a considerable number of photons per pixel.</li>
		<li>As excitation source, use laser light that is unpolarized (by passing through a non-polarization-maintaining optical fiber) or circularly polarized (by passing through a &#955;/4 waveplate) to maximize excitation of single chromophores, regardless of their orientation.</li>
		<li>Choose emission bandpass filters that efficiently cut-off both excitation and trapping laser lights. Note: In our experiments we labeled our protein with Atto532 dye and used a bandpass emission filter centered at 600 nm with 100 nm bandwidth.</li>
	</ol>
	</li>
</ol>
3. Microfluidics
To achieve a precise control of buffer exchange and &#39;dumbbell&#39; assembly, i.e., the anchoring of a single DNA molecule between two optically trapped microspheres, a custom-built laminar multichannel flow system must be developed. This is composed by a flow chamber, a pressure-reservoir and a pressure control system (Figure&#160;2).
<ol>
	<li>Flow cell preparation 
	NOTE: The flow chamber used for single-molecule experiments on protein-DNA interaction has preferably 4 inlets and 1 outlet, to allow up to 4 parallel buffer flows, each one containing one of the different components required for the experiment: beads, DNA, fluorescently-labeled protein, and imaging buffer. Separation of the components in the flow-channels allows efficient assembly of the dumbbell. Moreover, the imaging channel is useful to avoid background fluorescence from diffusing proteins and attain the high signal-to-noise ratio required for the localization of the DNA-binding protein with a precision of the order of few nanometers.
	<ol>
		<li>Drill 1 mm diameter holes in the microscope slide (76 x 76 x 1 mm) with a diamond-tip, in order to create 4 inlets and one outlet.</li>
		<li>Clean the microscope slides with ethanol and center the port with the O-ring inserted and the adhesive ring in the slide surrounding the access holes. It is fundamental to wear gloves during this step to avoid fingerprints, which would disturb the gluing process.</li>
		<li>Clamp the NanoPorts to the slide, and place it in the oven preheated at 180 &#176;C for 1 hr to allow complete attachment.</li>
		<li>Draw the outline pattern of the flow cell on a piece of paper. The channels should be ~3 mm wide and match the holes position on the microscope slide.</li>
		<li>Place the drawing on top of two pieces of Parafilm, and with a scalpel make sharp and continuous cuts along the drawn outline. Remove any protuberance formed. Discard the lower Parafilm layer, which is just used to guarantee a clean support.</li>
		<li>Place the microscope slide containing the attached NanoPorts upside down in the metallic support.</li>
		<li>Carefully align the Parafilm chamber outline with the holes, making sure that the parafilm does not obtrude the inlets and outlet of the chamber.</li>
		<li>Cover with a cleaned microscope coverslip (62 x 56 x 0.15 mm) and carefully remove excess Parafilm surrounding the chamber.</li>
		<li>Place two heat blocks, previously heated at 120 &#176;C, on top of the flow cell to apply constant pressure on it. Let the Parafilm melt for approximately 25 min.</li>
	</ol>
	</li>
	<li>Pressure reservoir and buffer containers 
	The pressure reservoir should be made of Plexiglas (or similar) and allow accommodation of six buffer containers. The possibility of having at least two extra buffer containers improves the instrument flexibility. Note that transparent containers and tubes considerably assist in flow-system handling and troubleshooting of trapped air bubbles. Sterile and disposable luer lock-tip syringes (2.5 ml), from which the plungers have been previously removed, are good buffer containers and allow an easy connection with the inlet tubing. Make sure that the total fluid volume is small compared with the air volume inside the pressure reservoir, a condition essential to obtain smooth and stable flow.
	<ol>
		<li>Connect shut-off valves at the exit of the buffer containers for turning the buffer flow on or off during the experiments.</li>
		<li>Mount the flow cell on the microscope stage. Connect the shut-off valves to the flow chamber inlets with polyetheretherketone tubing (inner diameter ~150 &#181;m) and flangeless fittings. Add ~3 cm of fluorinated ethylene propylene tubing (inner diameter ~500 &#181;m) as a linkage between the tubing and the NanoPort assemblies of the flow cell. This step is essential to reduce the buffer flow at the chamber entrance to avoid breakage of the flow cell or NanoPort detachment from the glass. The use of large inner diameter tubing alone, on the other hand, would require huge volumes of biological samples.</li>
		<li>Connect the flow chamber outlet to an open Falcon tube at atmospheric pressure. This tube serves as disposal for the buffer flowing out of the flow chamber.</li>
	</ol>
	</li>
	<li>Pressure control system
	<ol>
		<li>Build a pressure control system capable of finely adjusting the air pressure inside the pressure reservoir. This can be done by using two computer-controlled solenoid valves, one connected to a +0.5 atm pressure source (compressor) and the second one to atmospheric pressure. Solenoid valves are repeatedly opened for approximately 7 msec to increase or decrease the pressure in the line until the desired value is reached. NOTE: this approach was adapted from Carlos Bustamante20, and it yields a smoother flow than using a stepping motor syringe pump21.</li>
		<li>Add a pressure transducer controlled by a custom-written software to monitor the effective pressure exerted on the pressure reservoir. Typical working pressures are on the order of 20 mBar for dumbbell assembly and 200 mBar for washing the flow system components.</li>
	</ol>
	</li>
</ol>
4. DNA Tailing with Biotinylated Deoxycytidine Triphosphate (biotin-dCTP)
The DNA molecule should be longer than 1 &#956;m to facilitate its extension under flow and anchoring to the trapped beads. Moreover, a long DNA will prevent the IR trapping light from illuminating the DNA binding protein. This is of special relevance because absorption of IR light from an excited chromophore results in increased photobleaching. NOTE: In the present study, the DNA molecule was 3.7 &#956;m long. The molecule contained three copies of the primary operator O1 and one copy of each of the two auxiliary operators, O2 and O3. These sequences have been inserted in the middle of the molecule, thus resulting approximately equidistant from the trapped beads and IR laser beams.
<ol>
	<li>Mix 1 pmol of linear DNA 3&#8217;-ends with 60 pmol of biotin-14-dCTP, 4 &#956;l of 5x Terminal deoxynucleotidyl Transferase (TdT) buffer (1&#160;M potassium cacodylate, 125&#160;mM Tris, 0.05%&#160;(v/v) Triton X-100, 5&#160;mM CoCl2 (pH&#160;7.2 at 25&#176;C)) and 1.5 &#956;l of TdT and ddH2O to a total volume of 20 &#956;l. Incubate the mixture at 37 &#176;C for 15 min. This will result in the addition of up to 60 nucleotides per DNA extremity. Note that DNA 3&#8217;-overhangs are tailed with higher efficiency than recessed or blunt ends.</li>
	<li>Purify tailed DNA with spin columns or phenol/chloroform extraction and subsequent ethanol precipitation to remove CoCl2 from the TdT reaction buffer.</li>
	<li>After purification, DNA can be stored at 4 &#176;C for several weeks or at -20 &#176;C for longer periods but avoid repeated thawing and refreezing.</li>
</ol>
5. Labeling Protein Thiol Groups with ATTO532
Labeling through modification of cysteine residues must not alter the protein activity. To overcome this problem, we used a LacI mutant, LacIQ231C, which carries just one cysteine per monomer at position 231. This mutant preserves the same characteristics of the wild type and chemical modifications at position 231 have been shown to not interfere with protein stability 22. NOTE: We labeled the protein with ATTO532 maleimide.
<ol>
	<li>Ensure that the protein is dissolved in a buffer with a pH ranging from 7.0 to 7.5 and with a concentration ranging from 2 to 10 mg/ml. NOTE: in these experiments, LacI was dissolved in potassium phosphate 0.3 M, 5% glucose, pH 7.5 (buffer A).</li>
	<li>Dissolve Tris-(2-carboxyethyl)phosphine hydrochloride (TCEP) to a final concentration of 100 mg/ml in H2O. TCEP solution must be prepared fresh prior to use.</li>
	<li>Mix 100 &#956;l of the protein with 3 &#956;l TCEP and vortex carefully.</li>
	<li>Dissolve the dye in N,N-dimethylformamide (DMF) to a final concentration of 10 mg/ml working at low light condition. Vortex until completely dissolved.</li>
	<li>Reaction mix: Add 33 &#956;l of dye to the protein+TCEP and vortex carefully. Fast spin to collect as much solution as possible.</li>
	<li>Incubate the reaction mix for 2 hr in a shaker (8,000 rpm) at 20 &#176;C (or at 4 &#176;C O/N), protected from light.</li>
	<li>Dissolve L-Glutathione reduced (GSH) to a final concentration of 100 mg/ml in H2O. Add 3 &#956;l to the reaction mixture and incubate on the shaker at 8,000 rpm for 15 min. GSH will stop the reaction by blocking the reactivity of thiol groups.</li>
	<li>Purify the labeled protein using standard gel filtration columns, dialysis or spin concentrators. NOTE: In these experiments, labeled LacI was purified using 10&#160;kDa cutoff spin concentrators.
	<ol>
		<li>Dilute the reaction mixture with up to 12 ml of buffer A, apply it to the concentrator device, and centrifuge it at 8,000 x g, 1 hr, 4 &#176;C. The labeled LacI is thus concentrated to a final volume of about 500 &#956;l and the excess dye passes the membrane in the flow-through. Discard the flow-through and repeat step 5.8.1 at least three times.</li>
	</ol>
	</li>
	<li>Divide the labeled protein into small aliquots and store at -80 &#176;C. Avoid repeated freezing and thawing.</li>
</ol>
6. Combined Optical Trapping and Single-molecule Fluorescence Imaging Experiments
<ol>
	<li>Add 1 ml of buffer A to the buffer containers and apply 200 mBar pressure to wash the connecting tubes and flow cell. To maximize diffusion of the binding protein on DNA, here and in the following steps, buffer A may be replaced with a low ionic strength buffer. NOTE: In these experiments, we used TBE 0.5x.</li>
	<li>Check for the presence of air-bubbles, which might disrupt the flow smoothness. A gentle flick on the outlet tube will help to remove any remaining bubbles.</li>
	<li>Decrease the pressure to 20 mBar and verify that all channels and tubes are clear from air bubbles and the buffer flows with similar velocities in all channels.</li>
	<li>Prepare the samples to a final volume of 300 &#181;l: streptavidin-coated polystyrene beads 1.87 &#181;m; 20 pM of linear DNA, biotinylated on both extremities according to section 4; 300 pM of LacI tetramer labeled with ATTO532 according to section 5.</li>
	<li>Add each sample to one of the syringe containers in the following order: beads in the first channel, DNA in the second, third channel with the imaging buffer only (buffer A + oxygen scavenger system) and the labeled protein in the fourth channel.</li>
	<li>Turn the flow on at 200 mBar for 3 min to let the samples arrive inside the flow channel. Then reduce the pressure to 20 mBar.</li>
	<li>While imaging the first channel under bright field illumination, switch on the two optical tweezers and catch a polystyrene bead in each trap.</li>
	<li>Move quickly to the second channel to avoid that other beads fall into the traps. Note that the arrival in the DNA channel is easily noticeable by the end of the bead flow.</li>
	<li>Using one AOD, move one trap (bead) back and forth in the proximity of the other bead to allow the attachment of the flow-extended DNA in between the two optically trapped beads. When a DNA dumbbell is formed, the static bead starts following the movement of the bead that is actively moved back and forth by the trap.</li>
	<li>Move quickly to the buffer channel and turn off the buffer flow. Perform a force-extension curve analysis4,23-25 to verify the presence of a single DNA molecule. If more than one molecule is present (i.e., the enthalpic, raising phase of the force-extension curve occurs at a length shorter than the DNA contour length), discard the dumbbell and start again from step 6.7.</li>
	<li>Once a single DNA dumbbell is obtained, turn on the flow again and move the dumbbell to the protein channel. Switch to wide field fluorescence microscopy to monitor the interaction of the labeled-LacI with DNA. Turn off the flow and start acquisition.</li>
</ol>

<ol><li>Monico, C., Capitanio, M., Belcastro, G., Vanzi, F., Pavone, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Optical+Methods+to+Study+Protein-DNA+Interactions+in+Vitro+and+in+Living+Cells+at+the+Single-Molecule+Level%5BTitle%5D)+AND+%22International+journal+of+molecular+sciences%22%5BJournal%5D)">Optical Methods to Study Protein-DNA Interactions in Vitro and in Living Cells at the Single-Molecule Level.</a> International journal of molecular sciences. 14, 3961-3992 (2013).</li>
<li>Tinoco, I., Gonzalez, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Biological+mechanisms%2C+one+molecule+at+a+time%5BTitle%5D)+AND+%22Genes+%26+development%22%5BJournal%5D)">Biological mechanisms, one molecule at a time.</a> Genes & development. 25, 1205-1231 (2011).</li>
<li>Capitanio, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Exploring+molecular+motors+and+switches+at+the+single-molecule+level%5BTitle%5D)+AND+%22Micr.+Res.+Tech%22%5BJournal%5D)">Exploring molecular motors and switches at the single-molecule level.</a> Micr. Res. Tech. 65, 194-204 (2004).</li>
<li>Smith, S. B., Finzi, L., Bustamante, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Direct+mechanical+measurements+of+the+elasticity+of+single+DNA+molecules+by+using+magnetic+beads%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Direct mechanical measurements of the elasticity of single DNA molecules by using magnetic beads.</a> Science. 258, 1122-1126 (1992).</li>
<li>Block, S. M., Goldstein, L. S., Schnapp, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bead+movement+by+single+kinesin+molecules+studied+with+optical+tweezers%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Bead movement by single kinesin molecules studied with optical tweezers.</a> Nature. 348, 348-352 (1990).</li>
<li>Wang, M. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Force+and+velocity+measured+for+single+molecules+of+RNA+polymerase%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Force and velocity measured for single molecules of RNA polymerase.</a> Science. 282, 902-907 (1998).</li>
<li>Capitanio, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ultrafast+force-clamp+spectroscopy+of+single+molecules+reveals+load+dependence+of+myosin+working+stroke%5BTitle%5D)+AND+%22Nat+Methods%22%5BJournal%5D)">Ultrafast force-clamp spectroscopy of single molecules reveals load dependence of myosin working stroke.</a> Nat Methods. 9, 1013-1019 (2012).</li>
<li>Thompson, R. E., Larson, D. R., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Precise+nanometer+localization+analysis+for+individual+fluorescent+probes%5BTitle%5D)+AND+%22Biophys+J%22%5BJournal%5D)">Precise nanometer localization analysis for individual fluorescent probes.</a> Biophys J. 82, 2775-2783 (2002).</li>
<li>Yildiz, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Myosin+V+walks+hand-over-hand%3A+single+fluorophore+imaging+with+1.5-nm+localization%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Myosin V walks hand-over-hand: single fluorophore imaging with 1.5-nm localization.</a> Science. 300, 2061-2065 (2003).</li>
<li>Biebricher, A., Wende, W., Escude, C., Pingoud, A., Desbiolles, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tracking+of+single+quantum+dot+labeled+EcoRV+sliding+along+DNA+manipulated+by+double+optical+tweezers%5BTitle%5D)+AND+%22Biophys+J%22%5BJournal%5D)">Tracking of single quantum dot labeled EcoRV sliding along DNA manipulated by double optical tweezers.</a> Biophys J. 96, 50-52 (2009).</li>
<li>Candelli, A., Wuite, G. J., Peterman, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Combining+optical+trapping%2C+fluorescence+microscopy+and+micro-fluidics+for+single+molecule+studies+of+DNA-protein+interactions%5BTitle%5D)+AND+%22Physical+chemistry+chemical+physics+%3A+PCCP%22%5BJournal%5D)">Combining optical trapping, fluorescence microscopy and micro-fluidics for single molecule studies of DNA-protein interactions.</a> Physical chemistry chemical physics : PCCP. 13, 7263-7272 (2011).</li>
<li>Capitanio, M., Cicchi, R., Pavone, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Continuous+and+time-shared+multiple+optical+tweezers+for+the+study+of+single+motor+proteins%5BTitle%5D)+AND+%22Optics+and+Lasers+in+Engineering%22%5BJournal%5D)">Continuous and time-shared multiple optical tweezers for the study of single motor proteins.</a> Optics and Lasers in Engineering. 45, 450-457 (2007).</li>
<li>Harada, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Single-molecule+imaging+of+RNA+polymerase-DNA+interactions+in+real+time%5BTitle%5D)+AND+%22Biophys+J%22%5BJournal%5D)">Single-molecule imaging of RNA polymerase-DNA interactions in real time.</a> Biophys J. 76, 709-715 (1999).</li>
<li>van Mameren, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Counting+RAD51+proteins+disassembling+from+nucleoprotein+filaments+under+tension%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Counting RAD51 proteins disassembling from nucleoprotein filaments under tension.</a> Nature. 457, 745-748 (2009).</li>
<li>Capitanio, M., Maggi, D., Vanzi, F., Pavone, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fiona+in+the+trap%3A+the+advantages+of+combining+optical+tweezers+and+fluorescence%5BTitle%5D)+AND+%22J+Opt+A%3A+Pure+Appl+Opt%22%5BJournal%5D)">Fiona in the trap: the advantages of combining optical tweezers and fluorescence.</a> J Opt A: Pure Appl Opt. 9, s157(2007).</li>
<li>Dijk, M. A., Kapitein, L. C., Mameren, J., Schmidt, C. F., Peterman, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Combining+optical+trapping+and+single-molecule+fluorescence+spectroscopy%3A+enhanced+photobleaching+of+fluorophores%5BTitle%5D)+AND+%22J+Phys+Chem+B%22%5BJournal%5D)">Combining optical trapping and single-molecule fluorescence spectroscopy: enhanced photobleaching of fluorophores.</a> J Phys Chem B. 108, 6479-6484 (2004).</li>
<li>Capitanio, M., Cicchi, R., Pavone, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Position+control+and+optical+manipulation+for+nanotechnology+applications%5BTitle%5D)+AND+%22European+Physical+Journal+B%22%5BJournal%5D)">Position control and optical manipulation for nanotechnology applications.</a> European Physical Journal B. 46, 1-8 (2005).</li>
<li>Capitanio, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Calibration+of+optical+tweezers+with+differential+interference+contrast+signals%5BTitle%5D)+AND+%22Review+of+Scientific+Instruments%22%5BJournal%5D)">Calibration of optical tweezers with differential interference contrast signals.</a> Review of Scientific Instruments. 73, 1687-1696 (2002).</li>
<li>Elangovan, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+integrated+in+vitro+and+in+situ+study+of+kinetics+of+myosin+II+from+frog+skeletal+muscle%5BTitle%5D)+AND+%22J+Physiol%22%5BJournal%5D)">An integrated in vitro and in situ study of kinetics of myosin II from frog skeletal muscle.</a> J Physiol. 590, 1227-1242 (2012).</li>
<li>Wuite, G. J. L., Davenport, R. J., Rappaport, A., Bustamante, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+integrated+laser+trap%2Fflow+control+video+microscope+for+the+study+of+single+biomolecules%5BTitle%5D)+AND+%22Biophysical+Journal%22%5BJournal%5D)">An integrated laser trap/flow control video microscope for the study of single biomolecules.</a> Biophysical Journal. 79, 1155-1167 (2000).</li>
<li>Brewer, L. R., Bianco, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Laminar+flow+cells+for+single-molecule+studies+of+DNA-protein+interactions%5BTitle%5D)+AND+%22Nat+Methods%22%5BJournal%5D)">Laminar flow cells for single-molecule studies of DNA-protein interactions.</a> Nat Methods. 5, 517-525 (2008).</li>
<li>Rutkauskas, D., Zhan, H. L., Matthews, K. S., Pavone, F. S., Vanzi, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tetramer+opening+in+LacI-mediated+DNA+looping%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Tetramer opening in LacI-mediated DNA looping.</a> Proceedings of the National Academy of Sciences of the United States of America. 106, 16627-16632 (2009).</li>
<li>Marko, J. F., Siggia, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stretching+DNA%5BTitle%5D)+AND+%22Macromolecules%22%5BJournal%5D)">Stretching DNA.</a> Macromolecules. 28, 8759-8770 (1995).</li>
<li>van Mameren, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Unraveling+the+structure+of+DNA+during+overstretching+by+using+multicolor%2C+single-molecule+fluorescence+imaging%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">Unraveling the structure of DNA during overstretching by using multicolor, single-molecule fluorescence imaging.</a> Proc Natl Acad Sci U S A. 106, 18231-18236 (2009).</li>
<li>Wang, M. D., Yin, H., Landick, R., Gelles, J., Block, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stretching+DNA+with+optical+tweezers%5BTitle%5D)+AND+%22Biophys+J%22%5BJournal%5D)">Stretching DNA with optical tweezers.</a> Biophys J. 72, 1335-1346 (1997).</li>
<li>Ma, H., Long, F., Zeng, S., Huang, Z. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fast+and+precise+algorithm+based+on+maximum+radial+symmetry+for+single+molecule+localization%5BTitle%5D)+AND+%22Opt+Lett%22%5BJournal%5D)">Fast and precise algorithm based on maximum radial symmetry for single molecule localization.</a> Opt Lett. 37, 2481-2483 (2012).</li>
<li>Parthasarathy, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rapid%2C+accurate+particle+tracking+by+calculation+of+radial+symmetry+centers%5BTitle%5D)+AND+%22Nat+Methods%22%5BJournal%5D)">Rapid, accurate particle tracking by calculation of radial symmetry centers.</a> Nat Methods. 9, 724-726 (2012).</li>
</ol>

The authors have nothing to disclose.

In the last decade, single molecule manipulation and imaging techniques have seen great progress in terms of spatial and temporal resolution. The combination of manipulation and imaging techniques is at the base of powerful instruments that now allow the control of the mechanical conditions of a single biological polymer, such as DNA, RNA or cytoskeletal filaments, and the simultaneous localization of single proteins interacting with the same polymer. Controlling the mechanical conditions of the trapped polymer is of par...

Single molecule (SM) techniques have greatly developed over the past thirty years to respond to the need of overcoming some of the limitations of traditional, bulk solution measurements 1-3. The manipulation of single biological molecules has created the opportunity to measure mechanical properties of biopolymers4 and control the mechanical parameters of protein-protein5 and protein-DNA interactions6,7. SM fluorescence detection, on the other hand, represents an incredibly versatile tool for studying protein activity in vitro and in vivo, leading to the possibility of localizing and tracking single molecules with nanometer precision. Through fitting of the instrument point-spread-function to the SM image, in fact, one can accomplish localization with a precision depending mainly on signal-to-noise ratio (SNR) and reaching a limit of about one nanometer8,9. These methodologies find powerful applications in the study of the dynamics of motor proteins, as well as of the diffusion processes underlying target search in DNA-binding proteins. The capability of determining diffusion constants as a function of the DNA sequence, residence time on the target and accurately measuring the DNA length explored during one-dimensional diffusion events, represent a powerful tool for the study of protein-DNA interaction dynamics and for the investigation of the mechanisms of specific target search.
Recently, the combination of these two techniques has produced a new generation of experimental setups10-14 enabling the simultaneous manipulation of a biological substrate (for example an actin filament or a DNA molecule) and detection/localization of an interacting partner enzyme (for example myosin or a DNA-binding protein). The advantages of these techniques mainly rest on the possibility of exerting mechanical control over the trapped polymer, thus enabling the study of interaction dynamics versus forces or torques. Also, the methodology allows measuring biochemical reactions far from the surface, avoiding one of the main limitations of classic SM methods, i.e., the need for immobilization of the molecules under study on a surface (glass slide or microspheres).
The combination of two single molecules techniques requires overcoming several technical difficulties, mainly arising from the requirements of mechanical stability and adequate SNR (especially when requiring localization with nm precision)15. Particularly, when coupling SM fluorescence detection with optical tweezers, the reduction of noise and photobleaching from the trapping infrared lasers16 and the control of biochemical buffers for assembly of the biological complexes and performance of the experimental measurements11 are of paramount importance. Here, we describe the methods for performing successful measurements in a dual trapping/SM Fluorescence localization setup. The methodology is illustrated with the example of lactose repressor protein (LacI) fluorescently labeled (with Atto532) and detected as it binds to a DNA molecule (trapped between two optical tweezers) containing specific LacI binding sequences (i.e., operators). We demonstrate the effectiveness of the method in detecting binding of LacI to DNA and diffusion along its contour in the target search process. The method is applicable to any combination of DNA sequence and DNA-binding protein, as well as to other systems (microtubules or actin filaments and the motor proteins interacting with them).

<table><tbody><tr><td>Elastomeric isolators</td><td>Newport</td><td>Newdamp</td><td>Choose the appropriate Newdamp elastomer depending on the microscope weight and resonance frequencies</td></tr><tr><td>Optical isolator</td><td>Optics for Research</td><td>IO-3-YAG-VHP</td><td></td></tr><tr><td>Nd:YAG laser, 1,064 nm wavelength</td><td>Spectra-Physics</td><td>Millennia IR</td><td></td></tr><tr><td>Acousto-optic deflectors (AODs)</td><td>A&amp;amp;A optoelectronic</td><td>DTS-XY 250</td><td></td></tr><tr><td>Direct Digital Synthesizers</td><td>Analog Devices</td><td></td><td></td></tr><tr><td>Quadrant detector photodiodes</td><td>OSI optoelectronics</td><td>SPOT-15-YAG</td><td></td></tr><tr><td>DIO and FPGA board</td><td>National Instruments</td><td>NI-PCI-7830R</td><td></td></tr><tr><td>Halogen lamp</td><td>Schott</td><td>KL 1500 LCD</td><td></td></tr><tr><td>Condenser</td><td>Olympus</td><td>U-AAC 1.4NA Aplanat Apchromat</td><td></td></tr><tr><td>Objective</td><td>Nikon</td><td>CFI Plan Apochromat 60x 1.2NA water immersion</td><td></td></tr><tr><td>532 nm laser</td><td>Coherent</td><td>Sapphire</td><td></td></tr><tr><td>CCD 200X and 2000X</td><td>Hamamatsu&amp;nbsp;</td><td>XC-ST70 CE</td><td></td></tr><tr><td>Electron-multiplied CCD</td><td>Hamamatsu&amp;nbsp;</td><td>C9100-13</td><td></td></tr><tr><td>Piezo stage with nm-accuracy</td><td>Physik Instrumente</td><td>P-527.2CL&amp;nbsp;</td><td></td></tr><tr><td>Emission Filter</td><td>Chroma Technologies</td><td>600/100m</td><td></td></tr><tr><td>Silica beads (1.54 mm)</td><td>Bangs Laboratories</td><td>SS04N/5303</td><td></td></tr><tr><td>Albumin from bovine serum (BSA)</td><td>Sigma Aldrich</td><td>B4287</td><td></td></tr><tr><td>Pentyl acetate&amp;nbsp;</td><td>Sigma Aldrich</td><td>46022</td><td>Flammable liquid and vapor (No 1272/2008)</td></tr><tr><td>Nitrocellulose</td><td>Sigma Aldrich</td><td>N8267-5EA</td><td>Flammable solid&amp;nbsp; (No 1272/2008)</td></tr><tr><td>Heat block</td><td>MPM Instruments Srl</td><td>M502-HBD</td><td>with 2 removable blocks; preheated at 120 &amp;deg;C</td></tr><tr><td>NanoPort assemblies</td><td>Upchurch Scientific Inc.</td><td>N-333</td><td></td></tr><tr><td>Polyetheretherketone tubing&amp;nbsp;</td><td>Upchurch Scientific Inc.</td><td>1535</td><td></td></tr><tr><td>Home-made metallic holder for the assembly of the flow-chamber pressure reservoir made of Plexiglass</td><td></td><td></td><td></td></tr><tr><td>Luer lock-tip syringes 2.5 ml</td><td>Terumo</td><td>SS 02LZ1</td><td></td></tr><tr><td>Shut-off valves&amp;nbsp;</td><td>Upchurch Scientific, Inc.</td><td>P-732</td><td></td></tr><tr><td>Flangeless fittings&amp;nbsp;</td><td>Upchurch Scientific, Inc.</td><td>LT-111</td><td></td></tr><tr><td>Fluorinated ethylene propylene tubing&amp;nbsp;</td><td>Upchurch Scientific, Inc.</td><td>1549</td><td></td></tr><tr><td>Two computer-controlled solenoid valves</td><td>Clippard, Cincinnati, USA</td><td>ET-2-H-M5</td><td></td></tr><tr><td>Pressure transducer</td><td>Druck LTD</td><td>PTX 1400</td><td></td></tr><tr><td>biotin-14-dCTP&amp;nbsp;</td><td>Life Technologies</td><td>19518-018</td><td></td></tr><tr><td>Terminal deoxynucleotidyl Transferase (TdT)</td><td>Thermoscientific</td><td>EP0161</td><td></td></tr><tr><td>ATTO532 maleimide</td><td>Sigma Aldrich</td><td>68499</td><td></td></tr><tr><td>&lt;em&gt;N,N&lt;/em&gt;-dimethylformamide (DMF)&amp;nbsp;</td><td>Sigma Aldrich</td><td>227056</td><td>Combustible Liquid, Harmful by skin absorption. Irritant, Teratogen. H226; H303; H312; H316; H319; H331; H360; P201; P261; P280;P305; P351; P338; P311</td></tr><tr><td>Tris-(2-carboxyethyl)phosphine hydrochloride (TCEP)&amp;nbsp;</td><td>Sigma Aldrich</td><td>C4706</td><td></td></tr><tr><td>L-Glutathione reduced (GSH)</td><td>Sigma Aldrich</td><td>G4251</td><td>Acute toxicity, Oral (Category 5), H303</td></tr><tr><td>Amicon Ultra-15, PLQK Ultracel-PL Membrane, 10&amp;nbsp;kDa cutoff spin concentrators</td><td>Merck Millipore</td><td>UFC901024</td><td></td></tr><tr><td>Streptavidin-coated polystyrene beads 1.87 &amp;micro;m</td><td>Spherotech, Inc.</td><td>SVP-15-5</td><td></td></tr></tbody></table>

combining single-molecule manipulation and imaging for the study of protein-dna interactions

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.

In a successful experiment, one (or more) labeled proteins undergo binding/unbinding and/or monodimensional diffusion along the DNA molecule (Figure 3A). Localization of proteins along the DNA molecule allows the quantification of kinetic parameters as a function of the DNA sequence. When buffer conditions causing 1D diffusion are applied, it is possible to follow protein trajectories, and determine, for example, the diffusion coefficient D1D.
The precise position o...

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 ...

The overall goal of this procedure is to observe the dynamic interaction between a single DNA binding protein and a DNA molecule. This is achieved by combining in a custom built microscope, two single molecule techniques, double optical tweezers and fluorescence microscopy. This setup is also equipped with bright field illumination for visualization and nanometer stabilization of the sample.As a second step, a multi-channel flow cell integrated in a micro flow system is built to allow up to four laminar flows to efficiently recruit and assemble all the components required for the experiment. Next, the single molecule construct is assembled in order to perform the experiments. First two strip titin coated microspheres are caught with the dual optical tweezers in the first channel.The two trapped beads are then moved to the second channel to allow the attachment of the DNA template labeled with biotin on both extremities. The presence of a single DNA molecule is checked in the third channel by A DNA force extension analysis. Lastly, the bead DNA bead construct named DNA dumbbell is moved to the protein channel where the fluorescently labeled lactose repressor protein is present.Results are obtained that show diffusion and specific binding of single LAC eye molecules on a single stretched DNA molecule and the quantification of the kinetic parameters of LAC eye motion. The main advantages of this technique over existing single molecule methods are the possibility to control tension applied to the new molecule, the possibility to precisely track movements of the binding proteins over an nanometer stable spot, as well as the ability to operate far from the cover sleep surface. This method can help answer key questions on protein DNA interactions, especially on the mechanics of gene expression regulation, as well as the dynamics of protein diffusion along the DNA in the mechanisms of target search.These demonstration of this method is very useful to show the practical aspects of the construction of the flow cell sample handling and assembly of the beat DNA bead construct. This video will first demonstrate how to check the pointing stability of the two trapping lasers using micro beads stuck on a cover slip surface as a reference. Details on the design and construction of the optical tweezer setup can be found in the accompanying protocol text.Use a 24 by 60 millimeter cover slip to spread two microliters of silica beads onto the surface of a 24 by 32 millimeter cover slip. Using two pieces of double-sided tape. Attach the 24 by 32 millimeter cover slip onto a microscope slide to create a flow cell.Fill the flow cell with 50 millimolar of phosphate buffer and seal it with silicon grease image. One silica bead in brightfield microscopy at approximately 2000 times magnification. The field of view should be approximately seven by five micrometers acquired at 7 68 by 5 76 pixels so that the bead image has a diameter of a approximately 190 pixels compensate thermal drifts with a feedback software that calculates XY position from the image centro and Z from the diffraction rings and corrects drifts.Moving a pizo stage with nanometer accuracy or better overlap the center of the left trap with the bead center and measure the position noise and its standard deviation from the quadrant detector photo diode or QDP. Repeat the measurement for the right trap. The standard deviation of the position noise should be within a few nanometers to allow precise localization of DNA binding proteins to make the flow cell.A diamond tip is first used to drill one millimeter diameter holes in a 76 by 76 by one millimeter microscope. Slide to create four inlets and one outlet. This will allow up to four parallel buffer flows.Each containing one of the different components required for the experiment beads, DNA fluorescently labeled protein and imaging buffer. Clean the microscope slide with ethanol and center the port with the O-ring inserted and the adhesive ring in the slide surrounding the access holes. It is essential to wear gloves during this step to avoid fingerprints, which would disturb the gluing process.With the help of a second slide clamp the nano ports to the slide and place it in an oven preheated at 180 degrees Celsius for one hour to allow complete attachment. In the meantime, draw the outline pattern of the flow cell on a piece of paper. The channel should be approximately three millimeters wide and match the whole positions on the microscope slide.Place the drawing on top of two pieces of perfil and with a scalpel, make sharp and continuous cuts along the drawn outline. Remove any protuberances formed, discard the lower parfum layer, which is just used to guarantee a clean support. Next place the microscope slide containing the attached nano ports upside down in the metallic support.Carefully align the paraform chamber outline with the holes, making sure that the param does not extrude the inlets and outlet of the chamber cover with a cleaned microscope cover slip and carefully remove excess params surrounding the chamber. Place two heat blocks previously heated at 120 degrees Celsius on top of the flow cell to apply homogenous pressure on it. Let the para film melt for approximately 25 minutes.The next step is to prepare the pressure reservoir and buffer containers. The pressure reservoir is made of plexiglass and allows accommodation of up to six buffer containers. First, connect the shutoff valves at the exit of the buffer containers for turning the buffer flow on with the valve lever in the vertical position or off in the clockwise rotated position.During the experiments, mount the flow cell on the microscope stage. Then connect the shutoff valves through the poly ether, ether ketone, or peak tubing, and the flangeless spinnings to the flow chamber inlets. The peak tubing has an inner diameter of approximately 150 micrometers kilometers, approximately three centimeters of fluorinated ethylene propylene tubing with an inner diameter of approximately 500 micrometers are used as a linkage between the peak tubing and the nano port assemblies of the flow cell.This is essential to reduce the buffer flow at the chamber entrance to avoid breakage of the flow cell or nano port detachment from the glass. The use of large inner diameter tubing alone on the other hand, would require large volumes of biological samples. Connect the flow chamber outlet to an open Falcon tube at atmospheric pressure.This tube serves as disposal for the buffer flowing out of the flow chamber. To begin this procedure, add one milliliter of buffer A to each of the buffer containers and apply 200 millibars of pressure to wash the connecting tubes and flow cell. A pressure control system is capable.Finally, adjusting the air pressure inside the pressure reservoir. Check for the presence of air bubbles, which might disrupt the flow smoothness. A gentle flick on the outlet tube will help to remove any remaining bubbles, decrease the pressure to 20 millibars, and verify that all channels and tubes are clear from air bubbles and that the buffer flows with similar velocities in all channels.Prepare the samples to a final volume of 300 microliters. This experiment utilizes strept coated polystyrene beads, linear DNA biotinylated on both extremities that contain specific target sequences for lactose repressor binding and fluorescently labeled lactose repressor protein. Add each sample to one of the syringe containers in the following order beads in the first channel DNA in the second, the imaging buffer in the third and the labeled protein in the fourth channel.Turn the flow on at 20 millibars and open the shutoff valves to let the samples arrive inside the flow channel. This is easily detectable in the bead channel. While imaging the first channel under brightfield illumination, switch on the two optical tweezers and catch a polystyrene bead in each trap.Move quickly to the second channel to avoid the other beads falling into the traps. Note that the arrival in the DNA channel is easily noticeable by the end of the bead flow. Using one AO optic deflector or a OD move, one trap bead back and forth in the proximity of the other bead to allow the attachment of the flow extended DNA in between the two optically trapped beads.When a DNA dumbbell is formed, the static bead starts following the movement of the bead that is actively moved back and forth by the trap. Move quickly to the buffer channel and turn off the buffer flow. Perform a force extension curve analysis to verify the presence of a single DNA molecule.This is accomplished by separating the two beads apart while measuring the DNA tension. Once a single DNA dumbbell is obtained, turn on the flow again and move the dumbbell to the protein channel. Switch to wide field fluorescence microscopy to monitor the interaction of the labeled lactose repressor protein with DNA, turn off the flow and start acquisition.This method enables the detection of single lactose repressor or lac eye molecules interacting with a stretched DNA molecule. In this representative CH agram, a single lac eye molecule diffuses along non-specific DNA sequences, while another lac eye molecule is specifically bound to one operator located in the center of the DNA molecule. This plot shows the position of the two lac eye molecules along the direction connecting the centers of the two traps.As a function of time, the positions of the molecules were determined by the radial symmetry center algorithm shown here is a plot of the mean square displacement or MSD versus time for a single lac eye molecule diffusing along a stretched DNA molecule. The diffusion coefficient of the protein can be readily obtained from linear regression of the data. The combination of manipulation and imaging techniques is a powerful method, allowing mechanical control on a single biological polymer such as D-N-A-R-N-A or a cytoskeletal filament, and the simultaneous localization of single protein molecules interacting with the polymer itself.While attempting this Procedure, it's important to remember that combining single molecule manipulation and imaging techniques require careful design of the experimental setup to ensure nanometer level stability of the suspended polymer. On the other end, proper design of the biological probe is needed to guarantee special separation between the trapping G laser and the fluorescent probe. Please refer to the accompanying protocol text for details.After watching this video, you should have a good understanding of how to assemble the DNA molecule in between the two optical rapid beats the assembly of this construct. It's better obtained with the multichannel flow cell for which we provided the detailed description. We also indicated how to optimize buffer flow and to remove the air bubbles, which otherwise could compromise your experiments.In the combining text protocol, you can find the protocols to label the DNA with bioTE, which is necessary for the assembly with the streptavidin coated beats as well as the protocols to labeled the protein. Finally, we illustrated the step-by-step operations to quantify the binding and the diffusion of a single lactose repressive protein on a stretch DNA molecule.

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Research

JoVE Journal

Biology

A tapping mode atomic force microscope (AFM) method for the visualization of plasmid DNA, cytoplasmic proteins, and DNA-protein complexes is described. The method includes alternate approaches for preparing samples for AFM imaging following biochemical manipulation. DNA containing specific protein interacting regions are observed in near-physiologic buffer conditions.

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

This video describes a method to visualize the assembly of fusion protein condensates on DNA molecules at specific loci using single-molecule imaging. These condensates are formed due to LCD-LCD and LCD-DBD interactions, which allow for the recruitment of RNA polymerase II and enhanced gene transcription.

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

Photoactivated localization microscopy (PALM) combined with single-molecule tracking allows direct observation and quantification of protein-DNA interactions in live Escherichia coli cells.

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Here, we present a protocol to study DNA-protein interactions by total internal reflection fluorescence microscopy (TIRFM) using a site-specifically modified λ DNA substrate and a Quantum-dot labeled protein.

Visualizing the Interaction Between the Qdot-labeled Protein and Site-specifically Modified &#955; DNA at the Single Molecule Level

Using quantum-dot-labeled DNA and total internal reflection fluorescence microscopy, we can investigate the reaction mechanism of restriction endonucleases while using unlabeled protein. This single-molecule technique allows for massively multiplexed observation of individual protein-DNA interactions, and data can be pooled to generate well-populated dwell-time distributions.

Single-Molecule Dwell-Time Analysis of Restriction Endonuclease-Mediated DNA Cleavage

Here, we describe the use of the single-molecule imaging method, DNA Curtains, to study the biophysical mechanism of EWS-FLI1 condensates assembling on DNA.

Single-Molecule Imaging of EWS-FLI1 Condensates Assembling on DNA

DNA curtain, a high-throughput single-molecule imaging technique, provides a platform for real-time visualization of diverse protein-DNA interactions. The present protocol utilizes the DNA curtain technique to investigate the biological role and molecular mechanism of Abo1, a Schizosaccharomyces pombe bromodomain-containing AAA+ ATPase.

Deciphering Molecular Mechanism of Histone Assembly by DNA Curtain Technique

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

Many intrinsically disordered proteins have been shown to participate in the formation of highly dynamic biomolecular condensates, a behavior important for numerous cellular processes. Here, we present a single-molecule imaging-based method for quantifying the dynamics by which proteins interact with each other in biomolecular condensates in live cells.

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates

We report a hybrid ensemble and single-molecule assay to directly image and quantify the motion of fluorescently labeled, fully reconstituted Cdc45/Mcm2-7/GINS (CMG) helicase on linear DNA molecules held in place in an optical trap.

Hybrid Ensemble and Single-molecule Assay to Image the Motion of Fully Reconstituted CMG

This protocol outlines the procedure for performing deep, whole-cell single-molecule localization microscopy (SMLM) using a spinning disk confocal microscope with optical photon reassignment, thereby enabling DNA-PAINT imaging without the need for custom optics or complex illumination.

Whole-cell Super-Resolution Imaging via DNA-PAINT on a Spinning Disk Confocal with Optical Photon Reassignment

Learn to combine double optical tweezers and fluorescence microscopy for studying protein-DNA interactions at the single-molecule level. This method enables precise mechanical control and visualization of dynamic molecular binding events.

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

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