We thank Tony Kulesa, Robin Kirkpatrick and Evangelos Gatzogiannis for assistance with initial instrumentation setup and testing. We further thank Tony Kulesa for significant help writing and evaluating the customized single particle tracking software. This work was supported by startup funding and a Career Award at the Scientific Interface (to PCB) from the Burroughs Wellcome Foundation.

1. Preparation of biotin- &#955;-DNA
NOTE: Biotin-&#955;-DNA molecules are prepared by ligating a biotin-labeled oligonucleotide, 5&#39;-AGGTCGCCGCCC(A)20-biotin-3&#39; to&#160;&#955;-DNA molecules20.
<ol>
	<li>Prepare 0.1 mM biotin labeled oligo in TE buffer.</li>
	<li>Heat 0.5 mg/mL &#955;-DNA stock at 65 &#176;C for 60 s, and plunge into wet ice right away. 
	NOTE: Fast quench cooling reduces &#955;-DNA concatemerization.</li>
	<li>Pipette 100 &#181;L of the &#955;-DNA solution into a microcentrifuge tube.</li>
	<li>Add 1 &#181;L of the 0.1 mM oligo into 9 &#181;L of TE buffer.</li>
	<li>Add 2 &#181;L of the oligo solution to the microcentrifuge tube containing the &#955;-DNA. Mix thoroughly. 
	NOTE: Oligo is present in approximately a 12-fold molar excess over the complementary &#955;-DNA end.</li>
	<li>Heat the &#955;-DNA/oligo mixture at 65 &#176;C for 60 s.</li>
	<li>Slowly cool the mixture to room temperature. 
	NOTE: This step allows the oligo to anneal to the complementary &#955;-DNA end.</li>
	<li>Place the mixture on ice. Add 11 &#181;L of T4 DNA ligase reaction buffer (final buffer concentration 1x). Mix gently. Then add 2 &#181;L (800 units) of T4 DNA ligase enzyme. Mix gently.</li>
	<li>Incubate the mixture for 2 h at 16 &#176;C or overnight at 4 &#176;C.</li>
	<li>Purify the product using centrifugal filter tubes with a nominal molecular weight limit of 100 kDa. Specifically, transfer the step 1.9 mixture into a centrifugal filter tube and centrifuge at 14,000 x g for 5 min, wash with 400 &#181;L TE buffer twice, and collect the purified product. 
	NOTE: The as-purified biotin-&#955;-DNAs in TE buffer are stable at -20 &#176;C for up to one year.</li>
</ol>
2. Coverslip functionalization
NOTE: Glass coverslips are functionalized by reacting with silane-PEG-biotin. This one-step reaction protocol is simpler and more reliable in our experience than the standard two-step reaction protocol20. Coverslip functionalization with silane-PEG-biotin creates binding sites for streptavidin and minimizes nonspecific DNA and protein adsorption to the coverslip surface.
<ol>
	<li>Sonicate five no. 1 coverslips in 95% ethanol inside a staining jar for 10 min and then rinse with ultrapure water for three times. 
	NOTE: The fifth coverslip provides a spare in case of breakage.</li>
	<li>Fill the staining jar with 1 M KOH, sonicate for 10 min and then rinse with ultrapure water three times.</li>
	<li>Repeat the 2.1-2.2 cycle twice.</li>
	<li>Dry the coverslips under clean dry N2 gas flow.</li>
	<li>Further clean and dry the coverslips by applying air plasma treatment at 900 mTorr pressure for 5 min.</li>
	<li>Incubate coverslips with 25 mg/mL silane-peg-biotin dissolved in 95% ethanol at room temperature for 2 h. Specifically, sandwich 50 &#181;L silane-peg-biotin solution between two clean dry coverslips, and incubate the coverslip &#34;sandwiches&#34; inside a closed pipette tip box with about 10 mL of water at the bottom (not in contact with the coverslips) to minimize solvent evaporation.</li>
	<li>Wash away excess PEG molecules with ultrapure water and dry the PEG coated coverslips under clean dry N2 gas flow. 
	NOTE: The functionalized coverslips can be stored up to three days under ambient conditions.</li>
</ol>
3. Flow cell construction
NOTE: Flow cells are constructed by sandwiching a double-adhesive tape with pre-cut channels between a PEG functionalized coverslip and a PDMS slab containing inlet and outlet holes. Multiple channels are integrated per flow cell.
<ol>
	<li>Design flow channels in a CAD software and cut the channels (the width, depth and length of each channels are 1, 0.13 and 12 mm, respectively) on a double-adhesive tape by using a tape cutter (typically eight channels are integrated per flow cell). Remove tape residuals to form the flow channels.</li>
	<li>To make PDMS slabs, thoroughly mix 45 g of PDMS with 5 g of crosslinking reagent (enough for making twelve 5-mm thick PMDS slabs which can be stored indefinitely under ambient conditions) using a mixer, pour the mixture into two 10 cm Petri dishes, and leave the dishes inside a vacuum chamber until essentially all air bubbles are gone (manually remove air bubbles if any remain when the dishes are removed from the vacuum chamber). Then transfer the dishes into an 80 &#176;C oven until PDMS solidifies (about 2 h).</li>
	<li>Cut out a PDMS slab that matches the size of the double-adhesive tape with pre-cut channels. Peel one protection film off the double-adhesive tape and adhere the film to a flat face of the PDMS slab. Punch the outlet and inlet holes on the PDMS slab using a biopsy hole-puncher with a 23 Gauge needle.</li>
	<li>Peel the second protection film off the double-adhesive tape and adhere the PDMS-tape assembly to the functionalized surface of the coverslip (make sure that the coverslip is flat. See a picture of a fully assembled flow cell in Figure 1a).</li>
</ol>
4. TIRF Imaging
NOTE: Biotin-&#955;-DNAs are tethered to a flow channel surface and laminar flow is applied to flow stretch the DNA molecules (Figure 1b). The &#62; 80% stretched DNA molecules serve as spatially extended templates for observing the binding and transport activity of fluorescently labeled molecules. The trajectories of single molecules are tracked by time-lapse TIRF imaging (Figure 1c &#38; e).
<ol>
	<li>Fix the flow cell on a microscope stage and load pre-degassed blank buffer (10 mM sodium phosphate, 2 mM NaCl, 50 &#181;M EDTA, 20 mM ethanol, 5% (v/v) glycerol, 0.01% Tween-20, 1% (v/v) &#946;-mercaptoethanol, pH 7.4), 0.2 mg/mL streptavidin solution, 1 mg/mL Bovine Serum Albumin (BSA) solution and 100 pM biotin-&#955;-DNA solution into four reservoirs that each have a Tygon tubing connected to a solenoid value and another Tygon tubing connected to a PEEK tubing by sleeving the Tygon tubing over the smaller PEEK tubing (prevents backflow of reagents). Degas small volumes of buffer by overnight exposure to vacuum or shorter exposure to vacuum with stirring (until bubbles are no longer visible).</li>
	<li>Insert the PEEK tubing of the blank buffer reservoir into one inlet hole of the flow cell. Prime the channel by flowing in blank buffer, and insert three other PEEK tubing into inlet holes The air pressure driven flow of each reagent is controlled by solenoid valves and fully automated via a computer script. Be careful not to induce air bubble formation in the channel.</li>
	<li>Connect a Tygon tubing with a short PEEK tubing from the outlet hole to a waste container. The short PEEK tubing at the outlet helps suppress air bubble formation during infusion by maintaining positive pressure inside the channel.</li>
	<li>Tethering biotin- &#955;-DNA molecules to a flow channel surface.
	<ol>
		<li>Flow in 0.2 mg/mL streptavidin solution, incubate for 5 min and then flow in blank buffer to wash out the unbound streptavidin.</li>
		<li>Flow in 1 mg/mL BSA solution, incubate for 1 min, and then flow in blank buffer to wash out the unbound BSA. 
		NOTE: BSA further suppresses DNA and sample adsorption to the surface.</li>
		<li>Flow in 100 pM biotin-&#955;-DNA solution, incubate for 10 min and then flow in blank buffer to wash out the unbound DNAs. 
		NOTE: After the DNA is bound to the surface, avoid rapid flows to prevent damage to the tethered DNA molecules.</li>
	</ol>
	</li>
	<li>To check the quality of the functionalized coverslip, flow in DNA staining dye such as Sytox orange under the assay conditions to be used (read 4.6 below), and start fluorescence imaging under 532 nm laser illumination. 
	NOTE: The quality of the functionalized coverslip is typically good and the density of flow stretched DNA molecules will be high. It is essential to have a high enough density of flow stretched DNA molecules so that DNA binding and sliding/1D diffusion events can be observed.</li>
	<li>Fine tune the TIRF angle to achieve best signal to noise ratio of images of stretched DNA molecules (Figure 1d).</li>
	<li>To record trajectories of single molecules moving along DNA, repeat steps 4.2-4.4 inside a new channel (without DNA staining dye), then infuse pre-degassed sub-nanomolar fluorophore labeled molecules at a high enough flow rate by using a syringe pump and collect time-lapse fluorescence images with a frame rate of 100 Hz. 
	NOTE: A flow rate equivalent to a Weissenberg number (Wi: the dimensionless product of the longest polymer relaxation time - approximately 0.2 s for &#955;-DNA - and the shear rate - the change in flow velocity with distance from the coverslip surface) of 500 inside the channel is recommended for single molecule imaging with a frame rate of 100 Hz21.
	<ol>
		<li>Collect 10,000 frames in one Field-Of-View (FOV) to produce a movie, and collect similar movies from multiple (typically ten) FOVs per sample. 
		NOTE: The density of single particles in one FOV should be neither too high - which will complicate data analysis by making object assignment across frames ambiguous, nor too low - which might lead to low occurrence of single molecule events on DNA.</li>
		<li>Optimize the illumination intensity so that molecules bound to the coverslip surface are quickly photo-bleached to suppress background while molecules diffusing on DNA are not bleached too quickly so that longer trajectories can be detected. 
		NOTE: We typically use 200 - 800 W/cm2 power density in the FOV.</li>
	</ol>
	</li>
	<li>At the end of step 4.6, infuse DNA staining dye to confirm that DNA molecules can still be flow-stretched.</li>
	<li>Run both positive and negative control samples. 
	NOTE: A good positive control is a tetrapeptide, TetraMethylRhodamine(TMR)-KRRR (TMR is coupled to the N-terminal amine) which has a mean 1D diffusion constant of 10.5 &#177; 0.7 (standard error) M(bp2/s) at neutral pH11. A good negative control is to measure labeled samples of interest in assay buffer in a channel that has no DNA tethered, where no on-DNA diffusion activity should be detected.</li>
</ol>
5. Data Analysis:
NOTE:&#160;A custom single particle tracking software is used to identify trajectories of single molecules diffusing along DNA and estimate diffusion constants. This software first determines the centroid positions of single particles with high accuracy, identifies particles that are stuck on the coverslip surface, and then links particles in different frames to form time-lapse trajectories. From these trajectories, the 1D diffusion constants are estimated.
<ol>
	<li>To start data analysis, open a scripting software (see table of materials) and go to the directory of the single particle tracking software. Open the script named &#34;largedataprocess3.m&#34;. One important parameter to be defined is the threshold value for single particle detection.</li>
	<li>To determine the optimal threshold value, run the &#34;Determine_threshold_value.m&#34; script. This script will indicate how the threshold value affects single particle detection.After determining the optimal threshold value, run the &#34;largedataprocess3.m&#34; script.</li>
	<li>After completion of single particle tracking analysis, filter the raw trajectories by running the &#34;Trajectory_filtering.m&#34; script. A Graphic User Interface (GUI) is built in to visualize the filtering step.</li>
	<li>On the GUI panel, set the minimal displacement along DNA, MinXDisp, to 2 pixels; set the maximal displacement transverse to DNA, MaxYDisp, to 2 pixels; set the minimal number of frames, minFrames, to 10; set the minimal number of states of triplet, minTriplets, to 10; set the minimal diffusion constant along DNA, D_par, to 0; set the maximal estimated diffusion constant transverse to DNA, D_trans, to 10 M(bp2)S-1; set the minimal statistic parameter, Chi2_stat, to -5; set the minimal ratio of displacement along DNA to that transverse to DNA, MinX/Y, to 2. 
	NOTE: All raw trajectories that pass these filtering parameters are listed in Table 1, and those that do not are listed in Table 3.</li>
	<li>Click on a trajectory number in Table 1, the trajectory will be displaced in Graphics 1 &#38;2. Click on the &#34;Play&#34; button to play the raw fluorescence images. Click on the &#34;Add_to_Table2&#34; to identify the trajectory as a single molecule sliding trajectory. In the end, all trajectories added to Table2 will be saved when closing the GUI.</li>
	<li>To calculate the mean diffusion constants, run the script &#34;Calculate_mean_diffusion_constant.m&#34;. The mean diffusion constant is estimated from the set of trajectories that have passed all filtering steps and been added to Table 2.</li>
</ol>

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The Broad Institute may elect to file patent applications that include aspects of the work presented here.

When switching from the traditional two-step coverslip functionalization reaction protocol to a one-step reaction protocol, we noticed that the reliability of coverslip functionalization is significantly improved. As the total hands-on time for coverslip preparation is less than thirty min, we recommend preparing fresh coverslips each day single molecule imaging is planned. To minimize deterioration of silane-PEG-biotin, the reagent should be aliquoted and stored under dry N2 gas at -20 &#176;C upon receipt. Here we do not use PEG molecules that produce -COO- or -CH3 terminal groups that have been used in the past20. The negative charges of -COO- groups and the hydrophobic -CH3 groups were utilized to prevent nonspecific adsorption of DNA molecules and specific protein samples to the surface, respectively. We observe very low surface adsorption by small peptide samples on the fully biotin-terminated PEG surface, while silane-PEG-COOH and/or silane-PEG-CH3 might be useful for assays of certain protein samples or when assay conditions (such as pH &#60;7.5) promote DNA-surface interactions.
The shift from glass slide flow cells to PDMS flow cells speeds up the flow cell&#160;construction and allows easy integration of multiple channels. We often engineer eight channels per flow cell to enable eight independent experiments per functionalized coverslip. To further increase throughput per coverslip, even larger numbers of micro-fabricated channels could be used.
It is advisable to prevent air bubble formation inside the channel at any point after initial filling. Usually, air bubbles do not cause loss of tethered DNAs (although this is possible), but rather disrupt flow and possibly cause DNAs to stick to the coverslip. Buffer degassing and the addition of small surfactant quantities are typically effective in preventing bubble formation within the tubing and flow channel. In cases where air bubbles are introduced to the tubing, try to flush the bubbles out under a slow flow and make sure that they do not flow through the region where DNAs are tethered.
A streamlined and efficient data analysis software is critical, due to the huge amount of data generated. We usually collect 700,000 high-resolution images from measurements on one flow cell, which can be processed using our custom single particle tracking software within one day on a desktop computer with a quad-core processor and 32 Gb memory. The false positive rate of detecting 1D diffusion trajectories by the software (step 5.5) is sample dependent and can be quite low given that: 1) The density of single particles in the FOV is low enough so that there is little ambiguity linking particles across frames (protein samples in general are more prone to nonspecific adsorption to the coverslip than peptide samples, and thus, the buffer and the power density for each protein sample needs to be optimized); 2) The signal to noise ratio of single particles is high enough so that the likelihood of false-identification of particles is low. Still, the software can make occasional mistakes, and we recommend manual inspection of the data using the provided GUI to evaluate the software performance. In an application particularly sensitive to false-positive trajectory detection, process parameters including the threshold value, minFrames, minTriplets, and Chi2_stat can be set more stringently at the cost of lower sensitivity for true binding events and potentially greater statistical bias in trajectory identification. We here share and describe our single particle tracking software with the hope of helping standardize data analysis methods and stimulate discussion about best practices.
In our protocol, which requires continuous flow during imaging to maintain the stretched state of the DNA template molecules, the fluid flow could bias the transport of some proteins along DNA templates if they diffuse by a hopping mechanism or if the hydrodynamic radius of the labeled fluorophore is large22,23. While such bias can be measured and corrected in most datasets, the transport of proteins labeled with a small fluorophore and diffusing by a sliding mechanism is not typically biased to a detectable extent by flow2,4,24. Notably, the impact of the flow velocity on the transport of proteins along DNA has been used to study the mechanisms of protein diffusing along DNA23.&#160;A closely related alternative approach that enables measurements in the absence of flow utilizes DNA templates with biotin at both termini that are transiently stretched in flow and tethered by both ends to the coverslip such that the DNA molecules remain in an extended configuration when the flow is turned off25,26,27,28.&#160;The double-tether approach has the advantage of allowing flow-free transport experiments but requires modifying both ends of the DNA template molecules, careful flow control during tethering, and characterizing the distances between the two tethered sites. In addition, the impact of heterogeneous tensions and DNA conformational dynamics across DNA molecules on the single-molecule assay must be assessed.
Altogether, in addition to studying 1D diffusion of molecules along DNA, the as-reported single molecule assay can benefit many in vitro biochemical and biophysical studies at the single molecule level including DNA target search processes by proteins, enzymatic activities on DNA, and more.
Troubleshooting
Problem 1: The flow channels leak.
Solution: First, make sure that the design of flow channels allows at least 1.5 mm margin for sealing around and between channels; and then, during assembly of the flow cell, make sure that contact surfaces between the coverslip, the double-adhesive tape and the PDMS slab are flat, dry and free of debris, that the pressure applied to form the seal is uniform, and that the sealing operation is performed at room temperature.
Problem 2: The density of tethered DNAs is too low.
Solution: This problem arises when the PEG functionalization efficiency is low or the DNA molecules are not functionalized with biotin. From our experience, the density of tethered DNA should be high for &#62; 90% of the coverslips prepared from fresh or properly stored silane-PEG-biotin reagent. In cases when the density of tethered DNA is low with a proven batch of biotin-&#955;-DNA, try boosting concentrations of silane-PEG-biotin during PEG functionalization and concentrations of streptavidin and biotin-&#955;-DNA during DNA tethering. If this fails, replace the PEG and streptavidin reagents.
Problem 3: DNAs stick to the coverslip and cannot be stretched by flow.
Solution: This problem can also arise&#160;when the PEG functionalization efficiency is low, and is aggravated by low assay pH. Try flowing in more BSA or raising the pH (eg to 8.0) to help suppress DNA adsorption to the surface (step 4.3.2). If DNA adsorption is a persistent issue under a particular assay condition, try including up to 90% silane-PEG-COOH during PEG functionalization.
DATA ANALYSIS NOTES:
We use custom single particle tracking software to identify trajectories of single molecules diffusing along DNA, from which their 1D diffusion constants, D1 are estimated. To improve data analysis, we implemented the radial symmetry algorithm18&#160;for localizing particles and the covariance-based estimator19&#160;for estimating 1D diffusion constants, which dramatically sped up data analysis without compromising accuracy. We previously validated the custom software by analyzing simulated data11. The major steps are described below.
Particle identification:
This step is for detecting and segmenting particles - diffraction-limited spots - from background. First, spatially band-pass filter the images to enhance the signal of particles in the 3-5 pixel range (which corresponds to the size of the point-spread function on our system) and then threshold based on intensity (see script: spt2_SD_sandbox.m).
Centroid assignment
Segmenting the image only localizes molecules to pixel-level spatial resolution. To localize molecules to much higher precision, employ the Radial Symmetry super resolution algorithm18&#160;to images filtered as above. (This method is based on an analytical calculation and thus vastly reduces computational burden compared with other popular high-accuracy methods such as 2D Gaussian fitting (see script: spt2_SD_sandbox.m).)
Particle tracking: 
To track the trajectory of a particle through time, we must re-identify the same particles as they move between frames. We consider the initiation, blinking, and termination stages to be components of a trajectory. We constrain linkage of particles in current frames to those appearing in previous frames by the maximum displacement allowed per frame. In cases where multiple linkages are possible, the Jonker-Volgenant algorithm is used to yield globally optimal linkage sets (see script: traceTraj4_kx.3.m).
D1 estimation: 
To estimate D1 values from trajectories, we use the Covariance-Based Estimator (CVE)19.&#160;Based on an analytical calculation, this method is more computationally efficient and statistically rigorous than other commonly used methods such as regression of mean squared displacements (see script: runspt5_SD.m).
Trajectory Filtering:
Some trajectories contain artifacts such as linkage to surface-bound molecules and must be removed. We implemented several features such as minimum displacement parallel to DNA, maximum displacement transverse to DNA, minimal length of trajectory, minimal number of triplet consecutive detected positions, and minimum probability score (see Xiong et al11 for definition) that can be filtered to isolate a minimally biased set of trajectories likely to represent molecules diffusing on DNA (see script: sptViewFig2_update3x.m).

A long-standing problem in biology is how endogenous proteins that act at specific sites in the genome can locate their DNA targets quickly enough for the organism to survive and respond effectively to its environment. Studies over the past forty years proposed and largely support the hypothesis that the kinetics of DNA target search by a protein can be accelerated by facilitated diffusion in which the protein alternates between 3D diffusion in the bulk and 1D diffusion (including sliding and hopping processes) along the DNA1. It is now known that many proteins involved in gene regulation, nucleic acid metabolism, and other processes are capable of sliding on DNA2,3,4,5,6,7,8,9. Furthermore, recent studies reported that even small peptides can bind to and slide on DNA, with the ability to carry a cargo; for example, a protein molecule or PCR primer, along DNA10,11,12,13,14.
Over the last 15 years, the single molecule flow-stretching assay has been widely used to study binding and diffusion of molecules along DNA2,15,16.&#160;In this type of assay, biotinylated double-stranded DNA molecules are immobilized to the surface and a laminar flow is applied to flow stretch the DNA. The &#62; 80% stretched DNAs serve as spatially extended templates for studying binding and transport activity of molecules labeled with fluorophores where the trajectories of single molecules along DNA are tracked by time-lapse fluorescence imaging. In our implementation, optimized for reproducibility and ease of use, this assay consists of five major steps: preparation of biotin-&#955;-DNA, coverslip functionalization, flow cell construction, fluorescence imaging and data analysis. In previous protocols17, glass coverslips were functionalized&#160;by first reacting with (3-Aminopropyl)triethoxysilane (APTES) and then with amine-reactive polyethylene glycol (PEG) reagents (e.g., NHS-PEG-biotin) to form a PEG layer that resists nonspecific adsorption of assay components to the coverslip surface. The quality of functionalized coverslips largely depended on the quality of PEG reagents and reaction conditions at each step. Our protocol describes a simplified functionalization protocol and multiplex flow cell construction which requires no liquid adhesive&#160;or curing time on the day flow cells are assembled. We also describe a streamlined and robust data analysis procedure11 that eliminates computationally intensive regression steps by applying a radial symmetry method for centroid localization18&#160;and a covariance-based diffusion constant estimator19.
Here, we report a simple, robust and high throughput single molecule flow stretching assay implementation with significant improvements made in coverslip functionalization, flow cell construction and data analysis. In particular, we developed a one-step coverslip functionalization protocol, in which clean dry coverslips react directly with silane-PEG-biotin. This protocol simplifies the coverslip preparation and improves the reliability of the quality of functionalized surfaces compared to the standard two-step reaction protocol. We describe the use of so-functionalized coverslips with multi-channel PDMS flow cells that enable robust tubing connections to be made without glue. These flow cells further include multiple computer-controlled inlets for each flow chamber which enable automated reagent flow to reduce hands-on time during setup and increased assay throughput.

<table><tbody><tr><td>Biotin labeled oligo, 5&amp;rsquo;-AGGTCGCCGCCC(A)&lt;sub&gt;20&lt;/sub&gt;-biotin-3&amp;rsquo;</td><td>IDT</td><td>Custom oligo synthesis</td><td>Standard desalting purification is sufficient</td></tr><tr><td>&amp;lambda;-DNA</td><td>New England Biolabs Inc.</td><td>N3011S</td><td>Make aliquots and minimize freeze-thaw cycle number</td></tr><tr><td>T4 DNA ligase and ligation reaction buffer</td><td>New England Biolabs Inc.</td><td>M0202S</td><td></td></tr><tr><td>Amicon centrifugal filter tubes with NMWL of 100k Dalton</td><td>EMD Millipore</td><td>UFC510008</td><td></td></tr><tr><td>No. 1 glass coverslips</td><td>VWR</td><td>48393-106</td><td></td></tr><tr><td>Pure ethanol</td><td>VWR</td><td>89125-186</td><td></td></tr><tr><td>Potassium hydroxide pellets</td><td>Fisher Scientific</td><td>P250-500</td><td></td></tr><tr><td>Plasma cleaner, model No. PDC-32G</td><td>Harrick Plasma</td><td></td><td></td></tr><tr><td>Silane-peg-biotin</td><td>Nanocs</td><td>PG2-BNSL-5k</td><td>Store at -20&lt;sup&gt;o&lt;/sup&gt;C in dry nitrogen gas</td></tr><tr><td>A CAD software</td><td>Autodesk</td><td>AutoCAD</td><td></td></tr><tr><td>Double-adhesive tape</td><td>Adhesive Applications</td><td>8512-2-DP/DL</td><td></td></tr><tr><td>Tape cutter, model No. CE5000-40-CRP</td><td>Graphtec America</td><td></td><td></td></tr><tr><td>PDMS and crosslink reagent kit</td><td>R.S. Hughes</td><td>RTV615 CLEAR 010</td><td></td></tr><tr><td>Degassing chamber, model No. F42010-0000</td><td>BEL-ART Products</td><td></td><td></td></tr><tr><td>Mixer, model No. ARV-310</td><td>Thinky</td><td></td><td></td></tr><tr><td>Hole puncher, Harris Uni-Cores, Size 0.5 mm</td><td>Harris Uni-Cores</td><td></td><td>This can be substituted with a manual biopsy punch with a similar needle size</td></tr><tr><td>Tygon tubing</td><td>Cole Parmer</td><td>EW-06420-02</td><td>0.020&quot; ID, 0.060&quot;OD</td></tr><tr><td>Peek tubing</td><td>Western Analytical</td><td>PK007-031-Y</td><td>0.007&quot; ID, 0.031&quot;OD</td></tr><tr><td>Streptavidin</td><td>New England Biolabs Inc.</td><td>N7021S</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>New England Biolabs Inc.</td><td>B9000S</td><td></td></tr><tr><td>Sytox orange nucleic acid stain</td><td>Invitrogen</td><td>S11368</td><td></td></tr><tr><td>Ti-E Microscope</td><td>Nikon</td><td></td><td>Microscope, camera, objective, dichroic mirror and bandpass filter can be substituted for similar equipment</td></tr><tr><td>100x Objective, NA/1.49, CFI Apo TIRF</td><td>Nikon</td><td></td><td></td></tr><tr><td>CMOS camera</td><td>Hammamatsu</td><td>Orca Flash 4.0</td><td></td></tr><tr><td>Dichroic mirror</td><td>Semrock</td><td>FF560/659-Di01-25x36</td><td></td></tr><tr><td>Bandpass filter</td><td>Semrock</td><td>FF01-577/690-25</td><td></td></tr><tr><td>532 nm laser</td><td>Rayscience Innovation Limited, Shanghai</td><td>OEMLL-FN-532nm-C</td><td>Select wavelength to correspond to stain of choice</td></tr><tr><td>Syringe pump</td><td>Chemxy</td><td>Fusion 200</td><td></td></tr><tr><td>TMR-KRRR, &gt;=85% purity</td><td>MIT Biopolymer lab</td><td></td><td></td></tr><tr><td>pVIc-Cy3B, HPLC purified</td><td>A gift from Dr. Walter Mangel at Brookhaven National lab</td><td></td><td></td></tr><tr><td>Script programming software</td><td>MathWorks</td><td>Matlab</td><td></td></tr><tr><td>* see parts list for solenoid valves, computer interface, and assembly instructions at: https://sites.google.com/site/rafaelsmicrofluidicspage/valve-controllers</td><td></td><td></td><td></td></tr></tbody></table>

a simple, robust, and high throughput single molecule flow stretching assay implementation for studying transport of molecules along dna

We describe a simple, robust and high throughput single molecule flow-stretching assay for studying 1D diffusion of molecules along DNA. In this assay, glass coverslips are functionalized in a one-step reaction with silane-PEG-biotin. Flow cells are constructed by sandwiching an adhesive tape with pre-cut channels between a functionalized coverslip and a PDMS slab containing inlet and outlet holes. Multiple channels are integrated into one flow cell and the flow of reagents into each channel can be fully automated, which significantly increases the assay throughput and reduces hands-on time per assay. Inside each channel, biotin-&#955;-DNAs are immobilized on the surface and a laminar flow is applied to flow-stretch the DNAs. The DNA molecules are stretched to &#62;80% of their contour length and serve as spatially extended templates for studying the binding and transport activity of fluorescently labeled molecules. The trajectories of single molecules are tracked by time-lapse Total Internal Reflection Fluorescence (TIRF) imaging. Raw images are analyzed using streamlined custom single particle tracking software to automatically identify trajectories of single molecules diffusing along DNA and estimate their 1D diffusion constants.

Figure 2a shows the detected trajectories of pVIc-Cy3B (pVIc: GVQSLKRRRCF; Cy3B is conjugated to the cysteine residue) molecules diffusing on DNA in 2 mM NaCl buffer at pH 6.5. The diffusion of pVIc is bi-directional and driven by ambient thermal energy. From these trajectories, the 1D diffusion constants (D1 values) are estimated for each trajectory (see the histogram in Figure 2b). The mean D1 value of this pVIc conjugate is calculated to be 22.2 &#177; 0.9 (standard error) M(bp2/s) by averaging each D1 value across trajectories weighted by the number of triplet consecutive position calls contained in each trajectory.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55923/55923fig1.jpg" /> 
Figure 1: Apparatus for tracking motion of single molecules along DNA. 
a) An eight-channel PDMS microfluidic chip adhered to a functionalized coverslip to which DNA can be tethered (with USA quarter coin for scale); b) Flow cell zoom schematic showing flow-stretched &#955;-DNA molecules; c) Schematic of TIRF microscope and flow system for stretching DNA; d) TIRF image of a flow-stretched &#955;-DNA; e) TIRF image of pVIc-Cy3B (pVIc: GVQSLKRRRCF; Cy3B is conjugated to the cysteine residue) molecules bound to a flow-stretched &#955;-DNA. This figure has been modified with permission2,12. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55923/55923fig1large.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/55923/55923fig2.jpg" /> 
Figure 2: One-dimensional diffusion of pVIc (GVQSLKRRRCF) along DNA. 
a). Diffusion of pVIc along flow stretched &#955;-DNAs (137 trajectories) in 2 mM NaCl buffer at pH 6.5. x(t) and y(t) are the displacements along and transverse to &#955;-DNA, respectively. Dotted horizontal lines indicate missing points resulting from dye blinking. b). Histogram of the diffusion constant, D1 for pVIc diffusing along &#955;-DNAs. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55923/55923fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Simple, Robust, and High Throughput Single Molecule Flow Stretching Assay Implementation for Studying Transport of Molecules Along DNA at JoVE.com

A Simple, Robust, and High Throughput Single Molecule Flow Stretching Assay Implementation for Studying Transport of Molecules Along DNA

This protocol demonstrates a simple, robust and high throughput single molecule flow-stretching assay for studying one-dimensional (1D) diffusion of molecules along DNA.

We describe a simple, robust and high throughput single molecule flow-stretching assay for studying 1D diffusion of molecules along DNA. In this assay, ...

a-simple-robust-high-throughput-single-molecule-flow-stretching-assay

Nanyang Technological University

Research

JoVE Journal

Biochemistry

8.1K Views.  Massachusetts Institute of Technology and Harvard Medical School. Dit protocol demonstreert een eenvoudig, robuust en hoge doorvoer enkel molecuul stroom-stretching assay voor het bestuderen van de eendimensionale (1D) verspreiding van moleculen langs DNA.

Video: A Simple, Robust, and High Throughput Single Molecule Flow Stretching Assay Implementation for Studying Transport of Molecules Along DNA

Stretching Short Sequences of DNA with Constant Force Axial Optical Tweezers

Single-molecule techniques for stretching DNA of contour lengths less than a kilobase are fraught with experimental difficulties. However, many interesting biological events such as histone binding and protein-mediated looping of DNA1,2, occur on this length scale. In recent years, the mechanical properties of DNA have been shown to play a significant role in fundamental cellular processes like the packaging of DNA into compact nucleosomes and chromatin fibers3,4. Clearly, it is then important to understand the mechanical properties of short stretches of DNA. In this paper, we provide a practical guide to a single-molecule optical tweezing technique that we have developed to study the mechanical behavior of DNA with contour lengths as short as a few hundred basepairs.
The major hurdle in stretching short segments of DNA is that conventional optical tweezers are generally designed to apply force in a direction lateral to the stage5,6, (see Fig. 1). In this geometry, the angle between the bead and the coverslip, to which the DNA is tethered, becomes very steep for submicron length DNA. The axial position must now be accounted for, which can be a challenge, and, since the extension drags the microsphere closer to the coverslip, steric effects are enhanced. Furthermore, as a result of the asymmetry of the microspheres, lateral extensions will generate varying levels of torque due to rotation of the microsphere within the optical trap since the direction of the reactive force changes during the extension.
Alternate methods for stretching submicron DNA run up against their own unique hurdles. For instance, a dual-beam optical trap is limited to stretching DNA of around a wavelength, at which point interference effects between the two traps and from light scattering between the microspheres begin to pose a significant problem. Replacing one of the traps with a micropipette would most likely suffer from similar challenges. While one could directly use the axial potential to stretch the DNA, an active feedback scheme would be needed to apply a constant force and the bandwidth of this will be quite limited, especially at low forces.
We circumvent these fundamental problems by directly pulling the DNA away from the coverslip by using a constant force axial optical tweezers7,8. This is achieved by trapping the bead in a linear region of the optical potential, where the optical force is constant-the strength of which can be tuned by adjusting the laser power. Trapping within the linear region also serves as an all optical force-clamp on the DNA that extends for nearly 350 nm in the axial direction. We simultaneously compensate for thermal and mechanical drift by finely adjusting the position of the stage so that a reference microsphere stuck to the coverslip remains at the same position and focus, allowing for a virtually limitless observation period.

We illustrate the use of a constant force axial optical tweezers to explore the mechanical properties of short DNA molecules. By stretching DNA axially, we minimize steric hindrances and artifacts arising in conventional lateral manipulation, allowing us to study DNA molecules as short as ~100 nm.

Single-molecule techniques for stretching DNA of contour lengths  less than a kilobase are fraught with experimental difficulties. However, many ...

Bioengineering

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Biology

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. Linearized &lambda; DNA is modified to have a biotin on the end of one strand, and a digoxigenin moiety on the other end of the same strand. The biotinylated end is attached to a functionalized glass coverslip and the digoxigeninated end to a small bead. The assembly of these DNA-bead tethers on the surface of a flow cell allows a laminar flow to be applied to exert a drag force on the bead. As a result, the DNA is stretched close to and parallel to the surface of the coverslip at a force that is determined by the flow rate (Figure 1). The length of the DNA is measured by monitoring the position of the bead. Length differences between single- and double-stranded DNA are utilized to obtain real-time information on the activity of the replication proteins at the fork. Measuring the position of the bead allows precise determination of the rates and processivities of DNA unwinding and polymerization (Figure 2).

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system.

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. ...

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

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the averaging that takes place in bulk-phase studies. To achieve visualization, dual optical tweezers, where one trap is fixed and the other is mobile, are focused into one channel of a multi-stream microfluidic chamber positioned on the stage of an inverted fluorescence microscope. The optical tweezers trap single molecules of fluorescently labeled DNA and fluid flow through the chamber and past the trapped beads, stretches the DNA to B-form (under minimal force, i.e., 0 pN) with the nucleic acid being observed as a white string against a black background. DNA molecules are moved from one stream to the next, by translating the stage perpendicular to the flow to enable the initiation of reactions in a controlled manner. To achieve success, microfluidic devices with optically clear channels are mated to glass syringes held in place in a syringe pump. Optimal results use connectors permanently bonded to the flow cell, tubing that is mechanically rigid and chemically resistant, and which is connected to switching valves that eliminate bubbles that prohibit laminar flow.

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.

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the ...

CMG-Mediated DNA Unwinding Observed by ssDNA-RPA Tract Growth

Single-Molecule Real-Time Visualization of DNA Unwinding by CMG Helicase

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a dynamic complex of proteins termed the replisome. At the heart of the replisome is the&#160;CDC45-MCM2-7-GINS (CMG) helicase, which separates the two strands of the DNA double helix such that DNA polymerases can copy each strand. During genome duplication, replisomes must overcome a plethora of obstacles and challenges. Each of these threatens genome stability, as failure to replicate DNA completely and accurately can lead to mutations, diseases, or cell death. Therefore, it is of great interest to understand how CMG functions in the replisome during both normal replication and replication stress. Here, we describe a total internal reflection fluorescence (TIRF) microscopy assay using recombinant purified proteins, which allows for real-time visualization of surface-tethered stretched DNA molecules by individual CMG complexes. This assay provides a powerful platform to investigate CMG behavior at the single-molecule level, allowing helicase dynamics to be directly observed with real-time control over reaction conditions.

Author Spotlight: Unraveling the Dynamics of Eukaryotic DNA Replication Through Single-Molecule Visualization

This protocol demonstrates performing a single-molecule assay for live visualization of DNA unwinding by CMG helicase. It describes (1) preparing a DNA substrate, (2) purifying fluorescently labeled Drosophila melanogaster CMG helicase, (3) preparing a microfluidic flow cell for total internal reflection fluorescence (TIRF) microscopy, and (4) the single-molecule DNA unwinding assay.

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a ...

Characterizing Single-Molecule Conformational Changes Under Shear Flow with Fluorescence Microscopy

Single-molecule behavior under mechanical perturbation has been characterized widely to understand many biological processes. However, methods such as atomic force microscopy have limited temporal resolution, while F&#246;rster resonance energy transfer (FRET) only allow conformations to be inferred. Fluorescence microscopy, on the other hand, allows real-time in situ visualization of single molecules in various flow conditions. Our protocol describes the steps to capture conformational changes of single biomolecules under different shear flow environments using fluorescence microscopy. The shear flow is created inside microfluidic channels and controlled by a syringe pump. As demonstrations of the method, von Willebrand factor (VWF) and lambda DNA are labeled with biotin and fluorophore and then immobilized on the channel surface. Their conformations are continuously monitored under variable shear flow using total internal reflection (TIRF) and confocal fluorescence microscopy. The reversible unraveling dynamics of VWF are useful for understanding how its function is regulated in human blood, while the conformation of lambda DNA offers insights into the biophysics of macromolecules. The protocol can also be widely applied to study the behavior of polymers, especially biopolymers, in varying flow conditions and to investigate the rheology of complex fluids.

We present a protocol for immobilizing single macromolecules in microfluidic devices and quantifying changes in their conformations under shear flow. This protocol is useful for characterizing the biomechanical and functional properties of biomolecules such as proteins and DNA in a flow environment.

Single-molecule behavior under mechanical perturbation has been characterized widely to understand many biological processes. However, methods such as ...

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 ...

Studying DNA Looping by Single-Molecule FRET

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into nucleosomes. Thermodynamics of dsDNA bending has been studied by a method called cyclization which relies on DNA ligase to covalently join short sticky ends of a dsDNA. However, ligation efficiency can be affected by many factors that are not related to dsDNA looping such as the DNA structure surrounding the joined sticky ends, and ligase can also affect the apparent looping rate through mechanisms such as nonspecific binding. Here, we show how to measure dsDNA looping kinetics without ligase by detecting transient DNA loop formation by FRET (Fluorescence Resonance Energy Transfer). dsDNA molecules are constructed using a simple PCR-based protocol with a FRET pair and a biotin linker. The looping probability density known as the J factor is extracted from the looping rate and the annealing rate between two disconnected sticky ends. By testing two dsDNAs with different intrinsic curvatures, we show that the J factor is sensitive to the intrinsic shape of the dsDNA.

This study presents a detailed experimental procedure to measure looping dynamics of double-stranded DNA using single-molecule Fluorescence Resonance Energy Transfer (FRET). The protocol also describes how to extract the looping probability density called the J factor.

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into ...

Visualization of Surface-tethered Large DNA Molecules with a Fluorescent Protein DNA Binding Peptide

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating interactions between DNA and its binding proteins. Here, we report a method that uses fluorescent microscopy for visualizing large DNA molecules tethered on the surface. First, glass coverslips are biotinylated and passivated by coating with biotinylated polyethylene glycol, which specifically binds biotinylated DNA via avidin protein linkers and significantly reduces undesirable binding from non-specific interactions of proteins or DNA molecules on the surface. Second, the DNA molecules are biotinylated by two different methods depending on their terminals. The blunt ended DNA is tagged with biotinylated dUTP at its 3' hydroxyl terminus, by terminal transferase, while the sticky ended DNA is hybridized with biotinylated complimentary oligonucleotides by DNA ligase. Finally, a microfluidic shear flow makes single DNA molecules stretch to their full contour lengths after being stained with fluorescent protein-DNA binding peptide (FP-DBP).

We present an approach for visualizing fluorescent protein DNA binding peptide (FP-DBP)-stained large DNA molecules tethered on the polyethylene glycol (PEG) and avidin-coated glass surface and stretched with microfluidic shear flows.

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating ...

A Simple, Robust, and High Throughput Single Molecule Flow Stretching Assay Implementation for Studying Transport of Molecules Along DNA

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