This work was supported by Award Number K12GM074869 to CME from the National Institute of General Medical Sciences. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of General Medical Sciences or the National Institutes of Health.

1. General information
<ol>
	<li>Oligonucleotide design 
	NOTE: The 60 base-pair (bp)-long DNA substrate is formed from a pair of complementary oligonucleotides with a duplex melting temperature of 75 &#176;C in 100 mM NaCl.
	<ol>
		<li>Order one oligonucleotide synthesized with a single 5&#39; biotin modification and the other with a 5&#39; thiol modification (with a six-carbon spacer). Place the recognition site in the center of the duplex region. 
		NOTE: The oligonucleotide sequences for use with EcoRV are shown below (recognition site in bold). 
		5&#39;&#160;biotin - AAA ACC GAC ATG TTG ATT TCC TGA AAC GGG ATA TCA TCA AAG CCA TGA ACA AAG CAG CCG - 3&#39; 
		5&#39;&#160;thiol - CGG CTG CTT TGT TCA TGG CTT TGA TGA TAT CCC GTT TCA GGA AAT CAA CAT GTC GGT TTT - 3&#39;</li>
	</ol>
	</li>
	<li>Use ultrapure water with 18 MOhm resistivity for all steps.</li>
	<li>Protect all solutions containing quantum dots from light to prevent photobleaching.</li>
	<li>Use a compressed air source to complete this protocol.</li>
</ol>
2. Preparation of quantum-dot-labeled DNA substrate materials
NOTE: Besides the oligonucleotides described above, see the Table of Materials for other materials and Table 1 for buffers required for the preparation of quantum-dot-labeled DNA substrates.
<ol>
	<li>Reduce 5&#39; thiol groups on the oligonucleotide to be coupled to quantum dots.
	<ol>
		<li>Resuspend each thiolated oligonucleotide in water at a concentration of 100 &#181;M.</li>
		<li>Pipet 650 &#181;L of water into a size-exclusion spin column for each oligonucleotide sample, and vortex for ~15 s. Leave the column to pack for 30 min.</li>
		<li>Prepare fresh 100 mM dithiothreitol (DTT) solution immediately prior to each use as DTT degrades in solution rapidly. Carefully open one vial containing 7.7 mg of DTT, and pipet 500 &#181;L of sodium phosphate buffer into the vial; vortex to mix thoroughly.</li>
		<li>For each resuspended oligonucleotide, prepare a fresh tube, and add 50 &#181;L of the oligonucleotide and 50 &#181;L of DTT solution to the tube. Pipet up and down to mix. Incubate for 30 min at room temperature to reduce the disulfide bonds between the oxidized thiol groups on the 5&#39; end of the oligonucleotides.</li>
		<li>Remove the spin column caps, and place each column into the collection tube. Centrifuge the spin columns at 750 &#215; g for 2 min, and discard the eluate.</li>
		<li>Transfer the spin columns into fresh centrifuge tubes. Pipet the entire volume (100 &#181;L) of one sample of DNA/DTT mixture gently onto each prepared column. Centrifuge at 750 &#215; g for 2 min, and measure the absorbance of the eluate at 260 nm to confirm that the concentration is approximately 40 &#181;M.</li>
		<li>Store any samples that will not be used immediately at -20 &#176;C to prevent the oxidation of the thiol groups and the formation of disulfide bonds.</li>
	</ol>
	</li>
	<li>Coupling of DNA to quantum dots
	<ol>
		<li>Pipet one aliquot of 50 &#181;L of the quantum dot stock into a dialysis device for each construct to be made, making sure not to touch the membrane with the pipette tip. Dialyze against a volume of N-cyclohexyl-2-aminoethanesulfonic acid (CHES) buffer that is at least 1000x the sample volume with stirring at ~100 rpm for 15 min.</li>
		<li>Prepare a fresh 6 mM solution of sulfosuccinimidyl-4-(N-maleimidomethyl) cyclohexane-1-carboxylate (sulfo-SMCC) in CHES buffer immediately before each use. Carefully open one vial containing 2 mg of sulfo-SMCC, and pipet 800 &#181;L of CHES buffer into the vial. Vortex to mix thoroughly.</li>
		<li>Using a pipettor, carefully retrieve the suspended quantum dots from the dialysis device, and transfer into a fresh tube containing an equal volume of the sulfo-SMCC solution; pipet up and down to mix. Incubate for 1 h at room temperature with shaking at 1000 rpm to allow the sulfo-SMCC to react with the primary amines on the quantum dots.</li>
		<li>Using a pipettor, carefully transfer each sample into a fresh dialysis device. To remove excess sulfo-SMCC, dialyze against a volume of CHES buffer that is at least 1000x the volume contained in the dialysis device(s), with stirring, for 15 min. Exchange the buffer 2x, and perform dialysis with fresh CHES buffer 3x in total, allowing dialysis to proceed 15 min after each buffer exchange.</li>
		<li>To exchange the buffer in preparation for the second reaction, transfer the dialysis devices into a beaker containing a volume of phosphate-buffered saline (PBS) that is at least 1000x the volume contained in the device(s). Dialyze with stirring for 15 min, exchange the buffer 2x, and perform dialysis with fresh PBS 3x in total, allowing dialysis to proceed for 15 min after each buffer exchange.</li>
		<li>Using a pipettor, carefully recover the solution containing the quantum dots from the dialysis device(s), and transfer each sample into a fresh tube containing an equimolar amount of thiolated oligonucleotide diluted in PBS. Combine an equal volume of PBS and a 1/10th the volume of the reduced and purified oligonucleotide sample (approximately 40 &#181;M concentration) as the quantum dot concentration at this point is ~4 &#181;M. Incubate for 2 h at room temperature, with shaking at 1000 rpm.</li>
		<li>Add bovine serum albumin (BSA, 10 mg/mL) to each sample to obtain 0.5 mg/mL final BSA concentration. Using a pipettor, carefully transfer each sample into a fresh dialysis device, and dialyze against a volume of storage buffer that is at least 1000x the volume contained in the dialysis device(s). Exchange the buffer 2x, and perform dialysis with fresh storage buffer 3x in total, allowing dialysis to proceed for 15 min after each buffer exchange.</li>
		<li>Using a pipettor, carefully recover each sample, place it into a fresh tube, and store at 4 &#176;C. 
		NOTE: Do not store at -20 &#176;C as quantum dots are damaged by freezing.</li>
	</ol>
	</li>
	<li>Annealing of quantum-dot-labeled oligonucleotide to biotinylated oligonucleotide
	<ol>
		<li>Resuspend each biotinylated oligonucleotide in storage buffer at a concentration of 100 &#181;M. Store unused samples at -20 &#176;C.</li>
		<li>Combine a sample of quantum dot-labeled oligonucleotide with a 10-fold molar excess of biotinylated oligonucleotide. As the estimated concentration of DNA in the quantum-dot-labeled sample is 2 &#181;M, add 0.2 &#181;L of biotinylated oligonucleotide for every 10 &#181;L of this sample.</li>
		<li>Heat the mixture in a heat block to 75 &#176;C (the melting temperature for the complementary region). Hold at that temperature for 5 min. Turn off the heat block, and allow to cool slowly. Store the completed construct at 4 &#176;C; do not freeze.</li>
	</ol>
	</li>
</ol>
3. Surface functionalization of coverslips
NOTE: This process has been described previously in other JoVE video protocols16,17. This protocol describes an adapted version of the procedure with minor changes to accommodate a smaller glass slide. See the Table of Materials for other materials required for the surface functionalization of coverslips.
<ol>
	<li>Place 5 coverslips into each coverslip holder. Make sure to skip one space between each pair of coverslips so that they do not stick to one another. Place the coverslips in their holder into a jar with a cover, add ethanol to the jar until the coverslips are covered, and screw the top on securely. Place the entire jar into the water in the bath sonicator, without submerging it completely, and sonicate for 30 min.</li>
	<li>Fill a clean beaker or jar with ultrapure water. Use metal tweezers to remove the coverslips in their holder from the ethanol, and submerge them in water to rinse. Then, transfer the coverslips and holder into a jar containing a 1 M potassium hydroxide (KOH) solution. Screw the top on securely, place the jar in the sonicator bath, and sonicate for 30 min. 
	NOTE: Use care when handling the KOH solution as it is corrosive and an irritant.</li>
	<li>Repeat the sonication in ethanol and KOH solution, rinsing the coverslips in water as described above between each step.</li>
	<li>Transfer the coverslips in their holder into a clean jar filled with pure acetone. Finish the cleaning by sonicating the jar as described above for 30 min. Do not rinse with water after this step.</li>
	<li>Using metal tweezers, transfer the coverslips in their holder into a clean 100 mL beaker containing 80 mL of fresh acetone and a micro-scale stir bar. Place the beaker on a magnetic stir plate set to at least 1,000 rpm. While the acetone is being stirred vigorously, pipet 1.6 mL of 3-aminopropyltriethoxysilane (APTES) into the beaker to make a 2% v/v solution. 
	NOTE: Use care when pipetting APTES as it is corrosive.</li>
	<li>Allow the coverslips to incubate in the solution for 2 min, then use metal tweezers to transfer the coverslips in their holder into a beaker of water to quench the reaction. Rinse the coverslips two more times, replacing the water in the beaker.</li>
	<li>Cure the silanized glass in the oven at 120 &#176;C for 75 min. If not continuing to the next step immediately, store the coverslips under vacuum for maximum of a few days.</li>
	<li>Dissolve N-hydroxysuccinimide (NHS) ester-derivatized polyethyleneglycols (PEGs) in 100 mM sodium bicarbonate (pH 8.2). Use a 10:1 ratio of methoxy-terminated PEG to biotin-terminated PEG, with ~100 mg/mL of methoxy-terminated PEG.</li>
	<li>Pipet 100 &#181;L of the PEG solution onto half of the dry silanized coverslips, and cover each one with a second coverslip. Use small pieces of parafilm placed at each corner as spacers to prevent the coverslips from sticking to each other.</li>
	<li>Incubate the coverslips in a humid environment for 3.5 h. Separate the coverslip sandwiches. Use a squirt bottle to wash each coverslip with copious water and dry with compressed air.</li>
	<li>Store functionalized coverslips under vacuum. Be sure to keep the PEG-treated side up because the other side will not capture the DNA substrate.</li>
</ol>
4. Microfluidic device construction
NOTE: See the Table of Materials for other materials required for the construction of the microfluidic device.
<ol>
	<li>Using a handheld rotary multitool fitted with a tapered diamond point wheel bit, drill two holes in opposite corners of the quartz slide to serve as an inlet and outlet. Be sure to secure the slide in place, lubricate the bit, and slide constantly with water while drilling. After each experiment, recover the prepared quartz slide for reuse by soaking a used device in acetone to dissolve the adhesive and epoxy; discard the remaining components. 
	NOTE: The drilling can be carried out by hand, but the use of a press-style holder for the multitool makes the process easier.</li>
	<li>Combine 25 &#181;L of streptavidin solution with 80 &#181;L of PBS and 20 &#181;L of blocking buffer. Coat a PEG-treated coverslip with this mixture, and incubate in a humid environment for 30 min.</li>
	<li>During the incubation of the coverslip, prepare the imaging spacer to create a flow channel. Cut a 1-inch square piece of imaging spacer material, then mark a 2 mm wide channel aligned to the ends of the holes drilled in the quartz slide (Figure 2). Cut the channel out of the spacer material with a scalpel.</li>
	<li>Peel one side of the backing from the image spacer, and carefully place it onto the quartz slide, taking care not to cover the inlet and outlet holes. Be sure to clean the quartz slide thoroughly with acetone to remove any remnants of adhesive from prior experiments.</li>
	<li>Wash the coverslip with water, and dry with compressed air. Remove the other side of backing from the imaging spacer, and sandwich it between the quartz slide and the functionalized, streptavidin-coated coverslip, using the plastic tweezers to press the assembly together and remove air bubbles from the adhesive.</li>
	<li>Insert one 30 cm long polyethylene tubing into each hole in the quartz slide. Be sure to cut the ends of the tubing at an angle to ensure free flow of solution. Use a tube rack or other support to hold the tubes in place, and seal the tubes in place and the edges of the assembled device with epoxy. 
	NOTE: It works well to build the device on a piece of parafilm, which can easily be peeled off the bottom when the epoxy has set.</li>
	<li>Once the epoxy has set, insert the blunt needle on the empty syringe into the outlet tube of the device, and submerge the end of the inlet tube in a container filled with blocking buffer. Pull back on the syringe plunger to fill the device with blocking buffer. Leave the device to incubate for at least 30 min prior to use.</li>
</ol>
5. Surface tethering of quantum-dot-labeled DNA substrate
NOTE: Besides the microfluidic device, DNA substrate, and blocking buffer described above, see the Table of Materials for other materials and Table 1 for buffers required for the surface tethering of quantum-dot-labeled DNA substrates.
<ol>
	<li>Attach the microfluidic device to the microscope stage plate with tape, bring the objective into contact with the bottom of the device, and position the objective so that the field of view is within the microfluidic channel.</li>
	<li>Flush the microfluidic channel with fresh blocking buffer by pulling back the syringe plunger after connecting the outlet tube to the syringe pump. Check to make sure there are no bubbles trapped in the tubing or in the channel. 
	NOTE: From this point on, make sure that the inlet tube is in liquid so that no air bubbles are introduced into the device.</li>
	<li>Dilute 1 &#181;L of the prepared DNA substrate into 1 mL of blocking buffer. Place the inlet tube into the diluted DNA substrate, and flow 800 &#181;L of the substrate solution through the channel at a rate of 200 &#181;L/min. Allow the DNA solution to incubate in the channel undisturbed for 15 min after the flow stops.</li>
	<li>Flow at least 800 &#181;L of blocking buffer through the channel at a rate of 200 &#181;L/min to rinse the unbound DNA out of the channel.</li>
	<li>Adjust the laser power, microscope focus, and TIRF angle so that the surface-tethered quantum dots are clearly visible. 
	NOTE: Although quantum dots do not bleach quickly, it is best to keep the power as low as possible to minimize blinking.</li>
</ol>
6. REase-mediated DNA cleavage
NOTE: See the Table of Materials for materials and Table 1 for buffers required for REase-mediated DNA cleavage.
<ol>
	<li>Make sure that the camera is cooled to optimal operating temperature and set up for high-speed streaming with an exposure time of 0.10 s. Be prepared to collect data for ~4 min.</li>
	<li>Flush the microfluidic device with 800 &#181;L of experimental buffer without magnesium at a flow rate of 200 &#181;L/min.</li>
	<li>Add the REase to an aliquot (1 mL) of experimental buffer without magnesium, and mix gently by pipetting. Use 4 &#181;L of 100,000 units/mL stock, which corresponds to 400 units/mL of EcoRV. Flow 800 &#181;L of the diluted enzyme through the channel at a flow rate of 200 &#181;L/min.</li>
	<li>Start the experiment by flowing experimental buffer containing magnesium and fluorescein at a flow rate of 200 &#181;L/min. Begin capturing data immediately after starting the syringe pump. After flowing 800 &#181;L of buffer, stop data acquisition.</li>
</ol>
7. Data analysis
NOTE: See the Table of Materials for the data analysis software used for this protocol, and make adjustments if using a different analysis platform.
<ol>
	<li>Zero time-point determination
	<ol>
		<li>Subtract background fluorescence from each frame of the experimental movie using the imtophat function, a built-in morphological top-hat filtering function in the Image Processing Toolbox. Select a disk with a radius of three pixels as the structuring element. Obtain the background intensity by subtracting the filtered image from the original image. 
		NOTE: The filter only retains image features brighter than the background and smaller than the structuring element.</li>
		<li>Average the subtracted background over each movie frame. Use a trajectory of this value to determine the zero time-point for the experiment (Figure 3A). Determine the start time by finding the first frame at which the rate of increase exceeds 3x the standard deviation of the rate of change during the dead volume flow. 
		NOTE: A sharp increase in the rate of change of the average value of the subtracted background for each movie frame indicates the time at which the final experimental buffer containing the tracer dye entered the flow cell (Figure 3B).</li>
	</ol>
	</li>
	<li>Quantum dot intensity trajectory calculation and analysis
	<ol>
		<li>Calculate a maximum-intensity projection for the set of background-corrected movie frames recorded prior to the arrival of the tracer dye. Determine the locations of quantum dots to one-pixel accuracy in this projection image using a peak-finding function such as pkfnd.</li>
		<li>Generate intensity trajectories for individual quantum dots by calculating the average intensity in each movie frame of a square region three pixels per side surrounding the location returned by the peak-finding function. 
		NOTE: The background correction described above removes the artefacts introduced by the tracer dye. The resulting intensity trajectories should be relatively flat, but still include naturally occurring intensity fluctuations (Figure 4).</li>
		<li>Identify quantum dot disappearance events by statistical analysis of the intensity trajectories. For each quantum dot intensity trajectory, calculate a threshold intensity that is above the minimum value by an appropriate fraction of the difference between the maximum and minimum values for that trajectory, depending on the noise observed in the trajectories. 
		NOTE: The intensity fluctuation for the quantum dot is typically much greater than the background fluctuation. An appropriate value can be determined by comparing these values: the threshold selected must be higher than the highest background value, but ideally be lower than the lowest value for quantum dot fluorescence. For the data presented, the threshold used was calculated to be above the minimum value for each trajectory plus one-third of the difference between the maximum and minimum values for that trajectory. A quantum dot can be deemed to disappear at the last time point that the observed intensity at its location was above this threshold.</li>
		<li>Confirm putative disappearance events. Only include events in the final analysis if the observed intensity is above the midpoint between the minimum and maximum intensity values for the trajectory for over half of the movie frames prior to the time of disappearance and the standard deviation of the intensity trajectory decreased after the event. 
		NOTE: Other statistical tests may be employed to ensure that only valid disappearance events are logged.</li>
	</ol>
	</li>
</ol>

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The authors have no competing financial interests or other conflicts of interest

The DNA substrate for this assay is labeled with a quantum dot using a two-step reaction scheme using sulfo-SMCC. This bifunctional crosslinker consists of an NHS ester moiety that can react with a primary amine, and a maleimide moiety that can react with a sulfhydryl group20. The thiolated oligonucleotides used to prepare the substrate are shipped in their oxidized form. It is important to reduce and purify them, as described, before proceeding with the coupling procedure, or the efficiency of th...

Restriction endonucleases (REases) are enzymes that effect sequence-specific double-strand breaks in DNA. The discovery of REases in the 1970s led to the development of recombinant DNA technology, and these enzymes are now indispensable laboratory tools for genetic modification and manipulation1. Type II REases are the most widely used enzymes in this class as they cleave DNA at a fixed location either within or near their recognition sequence. However, there is a great deal of variation among the Type II REases, and they are divided into several subtypes based on particular enzymatic properties rather than being classified according to their evolutionary relationships. Among each subtype, there are frequent exceptions to the classification scheme, and many enzymes belong to multiple subtypes2. Thousands of Type II REases have been identified, and hundreds of them are commercially available.
However, in spite of the diversity among the Type II REases, very few REases have been studied in detail. According to REBASE, the restriction enzyme database established by Sir Richard Roberts in 19753, published kinetics data are available for fewer than 20 of these enzymes. Furthermore, while some REases have been directly observed at the single-molecule level while diffusing along the DNA prior to encountering and binding to their recognition sequence4,5,6,7, there are very few single-molecule studies of their cleavage reaction kinetics. The existing studies either do not report adequate statistics to undertake detailed analysis of the variation in the times at which single cleavage events take place8,9,10 or are not capable of capturing the full distribution of cleavage times11. This type of analysis can reveal the presence of relatively long-lived kinetic intermediates and could lead to better understanding of the mechanisms of REase-mediated DNA cleavage.
At the single-molecule level, biochemical processes are stochastic-the waiting time for a single instance of the process to occur, &#964;, is variable. However, many measurements of &#964; can be expected to obey a probability distribution, p(&#964;), that is indicative of the type of process taking place. For instance, a single-step process, such as the release of a product molecule from an enzyme, will obey Poisson statistics, and p(&#964;) will take the form of a negative exponential distribution:
<img alt="Equation 1" src="/files/ftp_upload/62112/62112eq1v3.png" />
where &#946; is the mean waiting time. Note that the rate of the process, k, will be equal to 1/&#946;, the inverse of the mean waiting time. For processes that require more than one step, p(&#964;) will be the convolution of the single-exponential distributions for each of the individual steps. A general solution for the convolution of N single-exponential decay functions with identical mean waiting times, &#946;, is the gamma probability distribution:
<img alt="Equation 2" src="/files/ftp_upload/62112/62112eq2v3.png" />
where &#915;(N) is the gamma function, which describes the interpolation of the factorial of N-1 to non-integer values of N. Although this general solution can be used as an approximation when the mean waiting times of individual steps are similar, it must be understood that the presence of relatively fast steps will be masked by steps with significantly longer waiting times. In other words, the value of N represents a lower limit on the number of steps12. With an adequate number of waiting-time measurements, the parameters &#946; and N can be estimated by binning the events and fitting the gamma distribution to the resulting histogram or by using a maximum-likelihood estimation approach. This type of analysis can therefore reveal the presence of kinetic steps that cannot be easily resolved in ensemble assays and requires a large number of observations to estimate parameters accurately12,13.
This paper describes a method to use quantum-dot-labeled DNA and total internal reflection fluorescence (TIRF) microscopy to observe hundreds of individual REase-mediated DNA cleavage events in parallel. The design of the assay makes it possible to pool the results of several experiments and can create dwell-time distributions containing thousands of events. The high photostability and brightness of quantum dots permit a 10 Hz time resolution without sacrificing the ability to observe cleavage events occurring even many minutes after the start of the experiment. Good temporal resolution and a broad dynamic range, combined with the ability to collect a large data set, allow accurate characterization of the dwell-time distributions to uncover the presence of multiple kinetic steps in the cleavage pathways of REases, which have turnover rates in the 1 min-1 range. In the case of EcoRV, three kinetic steps can be resolved, all of which have been identified through other means, confirming that the assay is sensitive to the presence of such steps.
Duplex DNA substrates containing the recognition sequence of interest are produced by annealing a biotinylated oligonucleotide to a complementary strand labeled with a single, covalently attached semiconductor nanocrystal quantum dot. These substrates are introduced into a flow channel built on top of a glass coverslip with a lawn of high molecular weight polyethylene glycol (PEG) molecules covalently attached to its surface. The DNA substrates are captured via a biotin-streptavidin-biotin linkage by a fraction of the PEG molecules that have a biotin at their free end. In TIRF microscopy, an evanescent wave that decays exponentially with distance from the glass-liquid interface provides illumination; the penetration depth is on the order of the wavelength of the light used. Under these conditions, only quantum dots that are tethered to the surface by a DNA molecule that has been captured on the functionalized glass surface will be excited. Quantum dots that are free in solution will not be constrained within the illuminated region and therefore will not luminesce. If the DNA tethering a quantum dot to the surface is cleaved, that quantum dot will be free to diffuse away from the surface, and it will disappear from the fluorescence image.
Although many Type II REases are known to bind DNA in the absence of magnesium14, all require magnesium to mediate DNA cleavage15. These REases can bind the surface-immobilized DNA in the absence of magnesium. When magnesium-containing buffer is flowed through a channel with REase prebound to the DNA, cleavage begins immediately, as indicated by the disappearance of quantum dots. The synchronization achieved by prebinding the REase molecules, and then initiating DNA cleavage by introducing magnesium, facilitates measurement of the lag time to the completion of DNA cleavage independently for each molecule in the population of enzymes observed in an experiment. Fluorescein is included as a tracer dye in the magnesium-containing buffer to indicate the arrival of magnesium into the field of view. As no enzyme is included in the magnesium-containing buffer, the lag time from the arrival of magnesium-containing buffer to the disappearance of each quantum dot indicates the time it takes for an REase that is already bound to the DNA to cleave the DNA and release the quantum dot from the glass surface. Quantum dot disappearance happens quickly and results in a sharp decrease in the intensity trajectory, providing a clear indication of the time at which a given DNA molecule is cleaved. The determination of event times is accomplished by mathematical analysis of intensity trajectories, and a typical experiment results in hundreds of identifiable cleavage events. The results of multiple experiments can be pooled to provide adequate statistics to allow estimation of the parameters, N and &#946;, by either nonlinear least-squares or maximum-likelihood analysis.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3-aminopropyltriethoxysilane (APTS)</td>
			<td>Sigma Aldrich</td>
			<td>440140-100ML</td>
			<td>Store in dessicator box</td>
		</tr>
		<tr>
			<td>5 Minute Epoxy</td>
			<td>Devcon</td>
			<td>20845</td>
			<td>use for sealing the microfluidic device</td>
		</tr>
		<tr>
			<td>acetone</td>
			<td>Pharmco</td>
			<td>329000ACS</td>
			<td>use for cleaning coverslips</td>
		</tr>
		<tr>
			<td>bath sonicator</td>
			<td>Fisher Scientific</td>
			<td>CPXH Model 2800</td>
			<td>catalog number 15-337-410</td>
		</tr>
		<tr>
			<td>Beaker, glass, 100 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Benchtop centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Biotin-PEG-Succinimidyl Valerate (MW 5,000)</td>
			<td>Laysan Bio</td>
			<td>BIO-SVA-5K</td>
			<td>Succinimidyl valerate has a longer half-life than succinimidyl carbonate</td>
		</tr>
		<tr>
			<td>biotinylated oligonucleotide</td>
			<td>Integrated DNA Technologies</td>
			<td>custom - see protocol for design considerations</td>
			<td>Request 5&#39; Biotin modification and HPLC purification</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>VWR</td>
			<td>0903-5G</td>
			<td>prepare a 10 mg/mL solution (aq) and heat to 95 &#176;C before using</td>
		</tr>
		<tr>
			<td>Centri-Spin-10 Size Exclusion Spin Columns</td>
			<td>Princeton Separations</td>
			<td>CS-100 or CS-101</td>
			<td>used to purify thiolated oligonucleotides after reducing the disulfide bond</td>
		</tr>
		<tr>
			<td>Centrifuge tubes 1.5. mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips, 1-inch square glass</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>coverslip holders</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>diamond point wheel</td>
			<td>Dremel</td>
			<td>7134</td>
			<td>use for drilling holes in quartz flow cell topper</td>
		</tr>
		<tr>
			<td>dithiothreitol (DTT)</td>
			<td>Thermo Scientific</td>
			<td>A39255</td>
			<td>No-Weigh Format, 7.7 mg/vial</td>
		</tr>
		<tr>
			<td>drill press rotary tool workstation stand</td>
			<td>Dremel</td>
			<td>220-01</td>
			<td>facilitates quartz drilling</td>
		</tr>
		<tr>
			<td>EcoRV (REase used to generate example data)</td>
			<td>New England Biolabs</td>
			<td>R0195T or R0195M</td>
			<td>Use 100,000 units/mL stock to avoid adding excess glycerol Check REBASE for suppliers of other REases</td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>various</td>
			<td>CAS 64-17-5</td>
			<td>denatured or 95% are acceptable, use for cleaning coverslips</td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid (EDTA)</td>
			<td>Sigma Aldrich</td>
			<td>EDS</td>
			<td>BioUltra, anhydrous, store in dessicator box</td>
		</tr>
		<tr>
			<td>Flea Micro Spinbar</td>
			<td>Fisherbrand</td>
			<td>14-513-65</td>
			<td>3 mm x 10 mm size to fit beneath coverslip rack</td>
		</tr>
		<tr>
			<td>fluorescein</td>
			<td>Acros Organics</td>
			<td>17324</td>
			<td>use to make experimental buffers</td>
		</tr>
		<tr>
			<td>gravity convection oven</td>
			<td>Binder</td>
			<td>9010-0131</td>
			<td></td>
		</tr>
		<tr>
			<td>handheld rotary multitool</td>
			<td>Dremel</td>
			<td>8220</td>
			<td>use for drilling holes in quartz flow cell topper</td>
		</tr>
		<tr>
			<td>ImagEM X2 EM-CCD Camera</td>
			<td>Hamamatsu</td>
			<td>C9100-23B</td>
			<td>air cooling is adequate for this experiment, use HCImage software or similar to control</td>
		</tr>
		<tr>
			<td>Imaging spacer, double-sided, adhesive</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Jar, glass with screw cap, (approximately 50 mm diameter by 50 mm high)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>magnesium chloride hexahydrate</td>
			<td>Fisher Bioreagents</td>
			<td>BP214-500</td>
			<td>use to make experimental buffer with magnesium</td>
		</tr>
		<tr>
			<td>MATLAB software</td>
			<td></td>
			<td></td>
			<td>Data analysis</td>
		</tr>
		<tr>
			<td>metal tweezers</td>
			<td>Fisher Brand</td>
			<td>16-100-110</td>
			<td></td>
		</tr>
		<tr>
			<td>methoxy-PEG-Succinimidyl Valerate (MW 5,000)</td>
			<td>Laysan Bio</td>
			<td>M-SVA-5K</td>
			<td>Both PEGs should have the same NHS ester so that the rate of reaction is consistent</td>
		</tr>
		<tr>
			<td>microcentrifuge</td>
			<td>Eppendorf</td>
			<td>5424</td>
			<td></td>
		</tr>
		<tr>
			<td>multiposition magnetic stirrer</td>
			<td>VWR</td>
			<td>12621-022</td>
			<td></td>
		</tr>
		<tr>
			<td>N-cyclohexyl-2-aminoethanesulfonic acid (CHES)</td>
			<td>Acros Organics</td>
			<td>AC20818</td>
			<td>CAS 103-47-9, use to make CHES buffer</td>
		</tr>
		<tr>
			<td>orbital shaker and heater for microcentrifuge tubes</td>
			<td>Q Instruments</td>
			<td>1808-0506</td>
			<td>with 1808-1061 adaptor for 24 x 2.0 mL or 15 x 0.5 mL tubes</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PE60 Polyethylene tubing (inner diameter 0.76 mm, outer diameter 1.22 mm)</td>
			<td>Intramedic</td>
			<td>6258917</td>
			<td>22 G blunt needles are a good fit for this tubing size</td>
		</tr>
		<tr>
			<td>Phosphate-Buffered Saline (PBS) 10x</td>
			<td>Sigma Aldrich</td>
			<td>P7059</td>
			<td>Use at 1x strength</td>
		</tr>
		<tr>
			<td>potassium hydroxide</td>
			<td>VWR Chemicals BDH</td>
			<td>BDH9262</td>
			<td>use a 1 M solution to clean coverslips</td>
		</tr>
		<tr>
			<td>Qdot 655 ITK Amino (PEG) Quantum Dots</td>
			<td>Invitrogen</td>
			<td>Q21521MP</td>
			<td></td>
		</tr>
		<tr>
			<td>Quartz Slide, 1 inch square, 1 mm thick</td>
			<td>Electron Microscopy Sciences</td>
			<td>72250-10</td>
			<td>holes must be drilled in the corners for inlet and outlet tubing insertion</td>
		</tr>
		<tr>
			<td>reinforced plastic tweezers</td>
			<td>Rubis</td>
			<td>K35a</td>
			<td>use for handling coverslips and building microfluidic device</td>
		</tr>
		<tr>
			<td>SecureSeal Adhesive Sheets</td>
			<td>Grace Biolabs</td>
			<td>SA-S-1L</td>
			<td>cut to form spacer for microfluidic device</td>
		</tr>
		<tr>
			<td>Single channel syringe pump for microfluidics</td>
			<td>New Era Pump Systems</td>
			<td>NE-1002X-US</td>
			<td>fitted with a 50 mL syringe and a 22 G blunt needle</td>
		</tr>
		<tr>
			<td>Slide-a-Lyzer MINI Dialysis Devices, 10 kDa MWCO, 0.1 mL</td>
			<td>Thermo Scientific</td>
			<td>69570 or 69572</td>
			<td>used for buffer exchange during quantum dot coupling to DNA</td>
		</tr>
		<tr>
			<td>sodium bicarbonate</td>
			<td>EMD Millipore</td>
			<td>SX0320</td>
			<td>use to make buffer for surface functionalization; 100 mM, pH 8</td>
		</tr>
		<tr>
			<td>sodium chloride</td>
			<td>Macron</td>
			<td>7581-12</td>
			<td>use to make experimental buffers</td>
		</tr>
		<tr>
			<td>Sodium phosphate dibasic solution (BioUltra, 0.5 M in water)</td>
			<td>Sigma Aldrich</td>
			<td>94046</td>
			<td>use to make 100 mM sodium phosphate buffer</td>
		</tr>
		<tr>
			<td>Sodium phosphate monobasic solution (BioUltra, 5M in water)</td>
			<td>Sigma Aldrich</td>
			<td>74092</td>
			<td>use to adjust pH of 100 mM sodium phosphate buffer</td>
		</tr>
		<tr>
			<td>Streptavidin from Streptomyces avidinii</td>
			<td>Sigma Aldrich</td>
			<td>S4762</td>
			<td>dissolve at 1 mg/mL and store 25 mL aliqouts at -20 &#8451;</td>
		</tr>
		<tr>
			<td>Sulfosuccinimidyl-4-(N-maleimidomethyl) cyclohexane-1-carboxylate (sulfo-SMCC)</td>
			<td>Thermo Scientific</td>
			<td>A39268</td>
			<td>No-Weigh Format, 2 mg/vial</td>
		</tr>
		<tr>
			<td>Syringe fitted with blunt 21 G needle</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>thiolated oligonucleotide</td>
			<td>Integrated DNA Technologies</td>
			<td>custom - see protocol for design considerations</td>
			<td>Request 5&#39; Thiol Modifier C6 S-S and HPLC purificaiton</td>
		</tr>
		<tr>
			<td>TIRF imaging system with 488 nm laser illumination</td>
			<td>various</td>
			<td></td>
			<td>custom built</td>
		</tr>
		<tr>
			<td>Tris -HCl</td>
			<td>Research Products International</td>
			<td>T60050</td>
			<td>use to make experimental buffers</td>
		</tr>
		<tr>
			<td>Tris base</td>
			<td>JT Baker</td>
			<td>4101</td>
			<td>use to make experimental buffers</td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Sigma</td>
			<td>P7949</td>
			<td>use to make blocking buffer</td>
		</tr>
		<tr>
			<td>Ultrapure water</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>vortex mixer</td>
			<td>VWR</td>
			<td>10153-842</td>
			<td></td>
		</tr>
		<tr>
			<td>Wash-N-Dry Coverslip Rack</td>
			<td>Electron Microscopy Sciences</td>
			<td>70366-16</td>
			<td>used for surface functionalization of coverslips</td>
		</tr>
	</tbody>
</table>

single-molecule dwell-time analysis of restriction endonuclease-mediated dna cleavage

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for hundreds of individual restriction endonuclease (REase) molecules in one experiment. This assay does not require protein labeling and can be carried out with a single imaging channel. In addition, the results of multiple individual experiments can be pooled to generate well-populated dwell-time distributions. Analysis of the resulting dwell-time distributions can help elucidate the DNA cleavage mechanism by revealing the presence of kinetic steps that cannot be directly observed. Example data collected using this assay with the well-studied REase, EcoRV - a dimeric Type IIP restriction endonuclease that cleaves the palindromic sequence GAT&#8595;ATC (where &#8595; is the cut site) - are in agreement with prior studies. These results suggest that there are at least three steps in the pathway to DNA cleavage that is initiated by introducing magnesium after EcoRV binds DNA in its absence, with an average rate of 0.17 s-1 for each step.

The flow cell is directly coupled to a high numerical aperture oil-immersion 60x magnification objective on an inverted microscope equipped with laser illumination for through-objective TIRF imaging (Figure 5A). After introducing the DNA substrate and washing away excess DNA and quantum dots, there are typically thousands of individual quantum dots in a field of view (Figure 5B). These quantum dots are stably attached to the glass surface, and they do not underg...

Watch this Scientific Journal Video about Single-Molecule Dwell-Time Analysis of Restriction Endonuclease-Mediated DNA Cleavage at JoVE.com

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

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.

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for ...

single-molecule-dwell-time-analysis-restriction-endonuclease-mediated

Research

JoVE Journal

Biochemistry

1.8K Views.  Tufts University. 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.

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

Determining if DNA Stained with a Cyanine Dye Can Be Digested with Restriction Enzymes

Visualization of DNA for fluorescence microscopy utilizes a variety of dyes such as cyanine dyes. These dyes are utilized due to their high affinity and sensitivity for DNA. In order to determine if the DNA molecules are full length after the completion of the experiment, a method is required to determine if the stained molecules are full length by digesting DNA with restriction enzymes. However, stained DNA may inhibit the enzymes, so a method is needed to determine what enzymes one could use for fluorochrome stained DNA. In this method, DNA is stained with a cyanine dye overnight to allow the dye and DNA to equilibrate. Next, stained DNA is digested with a restriction enzyme, loaded into a gel and electrophoresed. The experimental DNA digest bands are compared to an in silico digest to determine the restriction enzyme activity. If there is the same number of bands as expected, then the reaction is complete. More bands than expected indicate partial digestion and less bands indicate incomplete digestion. The advantage of this method is its simplicity and it uses equipment that a scientist would need for a restriction enzyme assay and gel electrophoresis. A limitation of this method is that the enzymes available to most scientists are commercially available enzymes; however, any restriction enzymes could be used.

Staining DNA molecules for fluorescence microscopy allows a scientist to view them during an experiment. In the method presented here, DNA molecules are pre-stained with fluorescent dyes and digested with methylation and non-methylation sensitive restriction enzymes.

Visualization of DNA for fluorescence microscopy utilizes a variety of dyes such as cyanine dyes. These dyes are utilized due to their high affinity and ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed extensive studies of cap-dependent translation and its regulation, high resolution kinetic characterization of this translation pathway is still lacking. Recently, we developed an in vitro assay to measure cap-dependent translation kinetics with single-molecule resolution. The assay is based on fluorescently labeled antibody binding to nascent epitope-tagged polypeptide. By imaging the binding and dissociation of antibodies to and from nascent peptide&#8211;ribosome&#8211;mRNA complexes, the translation progression on individual mRNAs can be tracked. Here, we present a protocol for establishing this assay, including mRNA and PEGylated slide preparations, real-time imaging of translation, and analysis of single molecule trajectories. This assay enables tracking of individual cap-dependent translation events and resolves key translation kinetics, such as initiation and elongation rates. The assay can be widely applied to distinct translation systems and should broadly benefit in vitro studies of cap-dependent translation kinetics and translational control mechanisms.

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Tracking individual translation events allows for high-resolution kinetic studies of cap-dependent translation mechanisms. Here we demonstrate an in vitro single-molecule assay based on imaging interactions between fluorescently labeled antibodies and epitope-tagged nascent peptides. This method enables single-molecule characterization of initiation and peptide elongation kinetics during active in vitro cap-dependent translation.

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed ...

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...

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

The fusion genes resulting from chromosomal translocation have been found in many solid tumors or leukemia. EWS-FLI1, which belongs to the FUS/EWS/TAF15 (FET) family of fusion oncoproteins, is one of the most frequently involved fusion genes in Ewing sarcoma. These FET family fusion proteins typically harbor a low-complexity domain (LCD) of FET protein at their N-terminus and a DNA-binding domain (DBD) at their C-terminus. EWS-FLI1 has been confirmed to form biomolecular condensates at its target binding loci due to LCD-LCD and LCD-DBD interactions, and these condensates can recruit RNA polymerase II to enhance gene transcription. However, how these condensates are assembled at their binding sites remains unclear. Recently, a single-molecule biophysics method-DNA Curtains-was applied to visualize these assembling processes of EWS-FLI1 condensates. Here, the detailed experimental protocol and data analysis approaches are discussed for the application of DNA Curtains in studying the biomolecular condensates assembling on target DNA.

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.

The fusion genes resulting from chromosomal translocation have been found in many solid tumors or leukemia. EWS-FLI1, which belongs to the FUS/EWS/TAF15 ...

Single-Molecule Analysis of Sf9 Purified Superprocessive Kinesin-3 Family Motors

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved single-molecule studies and has gained immense popularity in detecting and understanding the dynamic behavior of fluorescent-tagged molecules. Here, we describe a detailed method for&#160;in vitro single-molecule studies of kinesin-3 family motors using Total Internal Reflection Fluorescence (TIRF) microscopy. Kinesin-3 is a large family that plays critical roles in cellular and physiological functions ranging from intracellular cargo transport to cell division to development. We have shown previously that constitutively active dimeric kinesin-3 motors exhibit fast and superprocessive motility with high microtubule affinity at the single-molecule level using cell lysates prepared by expressing motor in mammalian cells. Our lab studies kinesin-3 motors and their regulatory mechanisms using cellular, biochemical and biophysical approaches, and such studies demand purified proteins at a large scale. Expression and purification of these motors using mammalian cells would be expensive and time-consuming, whereas expression in a prokaryotic expression system resulted in significantly aggregated and&#160;inactive protein. To overcome the limitations posed by bacterial purification systems and mammalian cell lysate, we have established a robust Sf9-baculovirus expression system to express and purify these motors. The kinesin-3 motors are C-terminally tagged with 3-tandem fluorescent proteins (3xmCitirine or 3xmCit) that provide enhanced signals and decreased photobleaching. In vitro single-molecule and multi-motor gliding analysis of Sf9 purified proteins demonstrate that kinesin-3 motors are fast and superprocessive akin to our previous studies using mammalian cell lysates. Other applications using these assays include detailed knowledge of oligomer conditions of motors, specific binding partners paralleling biochemical studies, and their kinetic state.

This study details purification of KIF1A(1-393LZ), a member of kinesin-3 family, using Sf9-baculovirus expression system. In vitro single-molecule and multi-motor gliding analysis of these purified motors exhibited robust motility properties comparable to motors from mammalian cell lysate. Thus, Sf9-baculovirus system is amenable to express and purify motor protein of interest.

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved ...

Studying TRF1/2 Interactions with Telomeric DNA Using Single-Molecule Assay

Analyzing Telomeric Protein-DNA Interactions Using Single-Molecule Magnetic Tweezers

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper function depends on tightly regulated processes of replication, elongation, and damage response. The shelterin complex, especially Telomere Repeat-binding Factor 1 (TRF1) and TRF2, plays a pivotal role in telomere protection and has emerged as a potential anti-cancer target for drug discovery. These proteins bind to the repetitive telomeric DNA motif TTAGGG, facilitating the formation of protective structures and recruitment of other telomeric proteins. Structural methods and advanced imaging techniques have provided insights into telomeric protein-DNA interactions, but probing the dynamic processes requires single-molecule approaches. Tools like magnetic tweezers, optical tweezers, and atomic force microscopy (AFM) have been employed to study telomeric protein-DNA interactions, revealing important details such as TRF2-dependent DNA distortion and telomerase catalysis. However, the preparation of single-molecule constructs with telomeric repetitive motifs continues to be a challenging task, potentially limiting the breadth of studies utilizing single-molecule mechanical methods. To address this, we developed a method to study interactions using full-length human telomeric DNA with magnetic tweezers. This protocol describes how to express and purify TRF2, prepare telomeric DNA, set up single-molecule mechanical assays, and analyze data. This detailed guide will benefit researchers in telomere biology and telomere-targeted drug discovery.

Author Spotlight: Advanced Single-Molecule Techniques for Investigating Telomeric Protein-DNA Interactions

This protocol demonstrates using single-molecule magnetic tweezers to study interactions between telomeric DNA-binding proteins (Telomere Repeat-binding Factor 1 [TRF1] and TRF2) and long telomeres extracted from human cells. It describes the preparatory steps for telomeres and telomeric repeat-binding factors, the execution of single-molecule experiments, and the data collection and analysis methods.

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper ...

In Situ Nucleosome Reconstitution for Single-Molecule Studies

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

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

Author Spotlight: Efficient Nucleosome Reconstitution for Single-Molecule Techniques

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

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

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 ...