We thank James Waters, Gable Wadsworth and Bo Broadwater for critically reading the manuscript. We also thank four anonymous reviewers for providing useful comments. We acknowledge financial support from Georgia Institute of Technology, the Burroughs Wellcome Fund Career Award at the Scientific Interface, and the student research network grant from NSF Physics of Living Systems.

1. dsDNA Sample Preparation
<ol>
	<li>Design globally curved DNAs by repeating a 10-mer sequence. For example, 5&#8217;-GTGCCAGCAACAGATAGC - (TTTATCATCCTTTATCATCCX)7 - TTTCATTCGAGCTCGTTGTTG-3&#8217; is a 186-bp curved DNA where X is a random extra base and the sequence flanking the repeating 10-mer sequence are adapter sequences. 
	NOTE: In this example two 10-mers with opposite preferences to nucleosome formation based on a large-scale nucleosome occupancy study by Kaplan et al.15 were chosen. Since the helical repeat of dsDNA is close to 10 bp, any net deflection of the helical axis of the 10-mer will accumulate to produce a shape like a circular arc (Figure 3A). Since the helical period is closer to 10.5 bp, an extra base is inserted after every two repeats to keep the curved structure as planar as possible. These sequences shorter than 200 bp can be ordered from companies that offer gene synthesis service. It is convenient to flank these sequences with common adapter sequences for subsequent steps. The procedure is schematized in Figure 3B.</li>
	<li>Perform PCR with Primer 1 (GTGCCAGCAACAGATAGC) and Primer 2 (/5Cy3/TAAATTCCTACAACAACGAGCTCGAATG). NOTE: Primer 2 is labeled with the FRET donor Cy3 at the 5&#39; end. A typical PCR recipe and cycling protocol are presented in Tables 1 and 2.</li>
	<li>Perform PCR with Primer 3 (/5BioTEG/GAAACATAG/iCy5/GAATTTACCGTGCCAGCAACAGATAGC) and Primer 4 (CAACAACGAGCTCGAATG). NOTE: Primer 3 is labeled with the FRET acceptor Cy5 through the backbone and the biotin-linker at the 5&#39; end. PCR recipe and cycling protocol are as above.</li>
	<li>Purify the PCR products using a PCR cleanup kit.</li>
	<li>Mix the Cy3-labeled product and the Cy5-labeled product in a buffer for strand exchange (100 mM NaCl, 10 mM Tris-HCl pH 7.0, 1 mM EDTA) at final concentrations of 0.4 &#181;M and 0.1 &#181;M, respectively. NOTE: The excess Cy3-DNA increases the concentration of the duplex carrying both Cy3 and Cy5 as a result of strand exchange.</li>
	<li>Exchange strands by incubating at 98.5 &#176;C for 2 min, gradually cooling to 5 &#176;C with ramp rate of 0.1 &#176;C/sec, and incubating at 5&#160;&#176;C for 2 hr.</li>
</ol>
2. Gel Electrophoresis to Detect dsDNA Curvature
<ol>
	<li>Pour the polyacrylamide gel16,17 by mixing acrylamide and bis-acrylamide solutions at 29.2:0.8 ratio, 5% (w/v) in 1X Tris/Borate/EDTA (TBE) buffer at pH 8.0. Note: 10 ml of gel solution contains: 1.217 ml 40% acrylamide, 0.667 ml 2% bis-acrylamide, 1 ml TBE 10X, 100 L ammonium persulfate (APS) 10%, 10 L TEMED, and the rest is dH2O. Full solidification of the gel takes about 30 min.</li>
	<li>Load the polyacrylamide gel with DNA samples and the DNA ladder in 1X loading buffer (5% glycerol, 0.03% (w/v) bromophenol blue) and run the gel at 5-8 V/cm at 4 &#176;C for 45 min or until the dye front approaches the end of the gel.</li>
	<li>Stain the gel using 1X TBE buffer that contains 0.5 g/ml ethidium bromide for 30 min. Identify the DNA bands under UV illumination. Compare band positions with the size marker (100 bp DNA ladder) to calculate the apparent sizes of the DNA molecules. NOTE: Curved DNAs generally move slower than straight DNAs.</li>
</ol>
3. Flow Cell Preparation
<ol>
	<li>Drill 6-7 pairs of holes along two opposite edges of a glass slide (3&#8217;&#8217; x 1&#8217;&#8217;) using a drill press and diamond drill bits. After drilling, rub the slide in flowing water to remove visible glass powder. NOTE: The holes serve as perfusion inlets and outlets. During drilling, cooling the slide with water is important to prevent cracking.</li>
	<li>Place the slides upright in a glass jar and fill it with water. Sonicate for 15 min and transfer them into another glass jar dedicated for acetone cleaning. Fill it with acetone and sonicate for 15 min. Rinse the slides with ethanol using a spray bottle and then with water. Place them in a polypropylene jar, fill it with 5 M potassium hydroxide, and sonicate for 15 min. Finally, sonicate the slides in water for 15 min. Clean the coverslips (no. 1, 24 x 40 mm) using the same protocol. NOTE: Cleaned slides and coverslips can be stored in dH2O for long term use.</li>
	<li>Mix 1 mg of biotin-PEG-silane (MW 3400) with 80 mg of mPEG-silane (MW 2000) in 340 L of 0.1 M sodium bicarbonate solution. Mix well and centrifuge the mixture briefly to get rid of bubbles. NOTE: The functionalization of the surface with polyethylene glycol (PEG) helps reduce nonspecific binding of DNA to the surface.</li>
	<li>Put 80 L of the PEG solution on each slide and gently lower a coverslip over it. Wait for 45 min. Separate the coverslip from the slide with tweezers, rinse them with copious amount of dH2O and let them dry in open air.</li>
	<li>Place thin strips of double-stick tape across the slide to form channels. Align a coverslip over it and firmly press the coverslip against the slide to form liquid-tight channels. Use 5 minute epoxy to seal the edges of the channels.</li>
</ol>
4. Preparation of Trolox Solution
<ol>
	<li>Put ~30 mg Trolox and 10 ml dH2O in a flask and use a magnet stir bar to stir the solution in open air for 18 hr.</li>
	<li>Filter the solution using a 0.2 &#181;m filter and adjust the pH to 7 by adding ~6 &#181;l of 1 M Tris base (pH 11). Note: Trolox is an anti-blinking reagent which is commonly used in single-molecule studies18. The antifading action of Trolox comes from its oxidized derivative which is present in partially degraded Trolox solution19. Quickly dissolving Trolox in methanol or high pH Tris solution should be avoided because of inefficient oxidation.</li>
</ol>
5. Single-molecule Imaging
<ol>
	<li>Inject 15 &#181;l of NeutrAvidin solution (0.5 mg/ml) into the channel and wait for 2 min before rinsing with 100 &#181;l of T50 buffer (10 mM Tris-HCl, 50 mM NaCl, pH 7.0).</li>
	<li>Inject 50 &#181;l of DNA sample (50-100 pM) into the channel. Wait for 5 min and rinse the unbound DNA away with 100 L of T50 buffer. NOTE: DNA molecules will specifically bind to the surface through the NeutrAvidin-biotin interaction.</li>
	<li>Fill the channel with the imaging buffer that contains an oxygen scavenging system20 (100 mM PCD, 5 mM PCA, 1 mM Trolox, and 500 mM NaCl).</li>
	<li>Set the EMCCD in frame-transfer mode to stream 2 x 2 binned images (256 x 256) to the computer at 25 frames per sec.</li>
	<li>Put immersion oil on the microscope objective, and fix the flow cell on the microscope stage using specimen clips. Coarse-adjust the focus by looking at the laser reflection pattern on the wall. Fine-adjust the focus using the live view of fluorescence images on the monitor.</li>
	<li>Begin data acquisition with the 532 nm laser on. Check the live view and adjust the focus if necessary. Stop data acquisition when most molecules have photobleached. 
	NOTE: In this facility, a lab-written C program to control the microscope and display live images on the monitor as they are saved to the hard disk is used.</li>
</ol>
6. Image Processing and Data Analysis
NOTE: A time series of 256 x 256 images are processed by a MATLAB code to generate single-molecule time traces of Cy3 and Cy5 intensities. To pair pixels between the donor channel and the acceptor channel of the split-view image, 6-7 pairs of Cy3 and Cy5 spots, each pair from the same molecule evenly dispersed across the field of view, are manually picked, and an affine transformation is calculated using the coordinates of these spots as anchor points.
<ol>
	<li>Using a MATLAB script, look through all single-molecule time traces that show multiple transitions between low and high FRET signals. Identify the looped and unlooped states. 
	NOTE: The FRET signal is defined as the Cy5 intensity ( Ia ) divided by the sum of Cy3 and Cy5 intensities ( Ia + Id ). Looped state has high FRET value while unlooped state has low FRET value. The histogram of FRET signals from a single molecule is bimodally distributed because of reversible looping and unlooping.</li>
	<li>Find the threshold that separates the two distributions by determining the intersection between the two fitted Gaussian curves.</li>
	<li>Calculate the FRET efficiency as <img fo:content-width="0.6in" src="/files/ftp_upload/51667/eq7.jpg" /> and assign the looped states with high FRET values and the unlooped states with low FRET values.</li>
	<li>Using a MATLAB script, analyze the cumulative number of molecules ( N(t) ) that looped (or reached the high FRET state) at different time lapses since the start of data acquisition. NOTE: Since a dsDNA molecule can start in any conformation of the unlooped state at the start of data acquisition, the rate of increase in the looped population reflects the mean looping rate averaged over initial conformations. Extract the looping rate kloop by fitting N(t) with an exponential function: <img fo:content-width="1.8in" src="/files/ftp_upload/51667/eq1.jpg" /> If increases biphasically, it can be fitted with a double exponential function: <img fo:content-width="2.2in" src="/files/ftp_upload/51667/eq2.jpg" /> In this case, kloop is obtained from: <img fo:content-width="1.7in" src="/files/ftp_upload/51667/eq3.jpg" /> NOTE: Theoretically, the survival probability that a polymer has not looped at time t is not a single or double exponential function12. Exponential fitting is used as a practical means to extract the average looping time.</li>
</ol>
7. Determining the J Factor
NOTE: The J factor represents how concentrated one end of a dsDNA is around the other end. It can be determined by interpolating the concentration of one end segment of the DNA that would produce the same reaction rate with the other end segment as the measured looping rate. Experimentally, one end segment is immobilized on the surface, and the other end segment is introduced at a certain concentration c. If the measured annealing rate between the two ends is kanneal, then the J factor21 is given by <img fo:content-width="0.6in" src="/files/ftp_upload/51667/eq4.jpg" />. The annealing rate constant ( kanneal = kanneal / C ) is independent of the probe concentration.
<ol>
	<li>Flow 20 &#181;l of 30-50 pM biotin-Cy5 oligo (Primer 3) into a NeutrAvidin-coated channel. Rinse the channel with 100 &#181;l T50 to wash away unbound oligos.</li>
	<li>Prepare the imaging buffer as described in part 5 with the addition of Cy3 oligo (Primer 2) at a final concentration of 50 nM. Flow this imaging buffer into the channel.</li>
	<li>While keeping the 532 nm laser on, briefly turn the 640 nm laser on to identify locations of surface-bound Cy5 oligos. Turn off the 640 nm laser and start monitoring the FRET signal. 
	NOTE: Upon hybridization of a Cy3 oligo to a surface-bound Cy5 oligo, Cy5 fluorescence will arise from FRET.</li>
	<li>Using a MATLAB script, analyze the number of molecules which start in the unbound state (low Cy5 intensity) but later turn into the annealed state (high Cy5 intensity) as a function of time from the Cy5 intensity traces.</li>
	<li>Plot this number of annealed molecules vs. time. Fit this curve with a single exponential function (<img fo:content-width="1.8in" src="/files/ftp_upload/51667/eq5.jpg" />) to obtain the annealing rate ( kanneal ).</li>
	<li>Repeat this experiment at different Cy3 oligo concentrations (60, 100, and 180 nM) to confirm linearity between the annealing rate and the reactant concentration. Extract the second-order annealing rate constant ( kanneal ) from the slope.</li>
	<li>Calculate the J factor from <img fo:content-width="0.6in" src="/files/ftp_upload/51667/eq6.jpg" />, where kloop is the looping rate measured in the same buffer condition.</li>
</ol>

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The authors declare no conflicts of interest.

A simple single-molecule assay based on FRET was used to study looping kinetics of DNAs of different intrinsic shapes. Curved DNAs can be prepared by repeating a 10-mer sequence in phase with the helical period of 10.5 bp, and their curvatures can be estimated using PAGE. These dsDNAs are designed with sticky ends to allow transient loop stabilization. We extracted the looping rate from the exponential rise in the number of looped molecules over time. The annealing rate constant between the disconnected sticky end segmen...

Understanding the mechanical properties of dsDNA is of fundamental importance in basic sciences and engineering applications. The structure of dsDNA is more complicated than a straight helical ladder because roll, tilt, and twist angles between successive base pairs can vary with sequence. Thermal fluctuations can cause dsDNA to undergo diverse modes of conformational fluctuations such as bending, twisting and stretching. Transitions such as melting and kinking can also occur in extreme conditions.
Among these motions, dsDNA bending has the most noticeable biological impact1. dsDNA bending is associated with gene repression or activation by bringing two distant sites close to each other. It also plays an important role in DNA packaging inside the cell nucleus or a viral capsid. Bending deformation of dsDNA can be visualized experimentally by high-resolution microscopy (AFM2 and TEM3), and the thermodynamics and kinetics can be studied by looping assays, which chemically link juxtaposed sites of the dsDNA.
One such assay is ligase-dependent cyclization4. In this assay, dsDNA molecules with &#8216;sticky&#8217; (cohesive) ends are circularized or dimerized by DNA ligase. By comparing the rates of circle and dimer formation, one can obtain an effective molar concentration of one end of the DNA in the vicinity of the other end, which is known as the J factor. This J factor is dimensionally equivalent to the probability density of finding one end of the DNA at a short distance from the other end, and thus reflects the flexibility of the DNA. Measuring the J factor as a function of DNA length reveals many characteristics about DNA mechanics including the persistence length4,5.
The worm-like chain (WLC) model has been widely regarded as the canonical polymer model for dsDNA mechanics based on its success in explaining the force-extension curves obtained in DNA pulling experiments6, and correctly predicting the J factors of dsDNAs longer than 200 bp7. However, using the cyclization assay on dsDNA molecules as short as 100 bp, Cloutier and Widom measured the J factors to be several orders of magnitude higher than the WLC model prediction8. A year later, Du et al. produced J factors in agreement with the WLC model using the cyclization assay with lower concentrations of ligase and attributed the anomalous result from the Widom group to high ligase concentrations used9. This controversy exemplifies the unavoidable influence of DNA ligase on cyclization kinetics when using the conventional assay9. Moreover, DNA ligase can also affect DNA structure and stiffness through nonspecific binding10,11.
To eliminate the technical concerns of protein-dependent looping assays, we recently demonstrated a protein-free looping assay based on Fluorescence Resonance Energy Transfer (FRET)12. In this method, looped conformations are detected by FRET between the donor and acceptor attached near the sticky ends of a DNA molecule. An objective-type total internal reflection fluorescence microscope (TIRFM) is used to record trajectories of reversible looping and unlooping events from surface-immobilized single DNA molecules for a prolonged period of time. This method features PCR-based assembly of DNA molecules to generate mismatch-free DNA molecules, which is a crucial improvement over a similar method by Vafabakhsh and Ha13. The single-molecule aspect of this protocol allows measurement of distributions in addition to ensemble averages while the FRET aspect allows one to measure DNA looping dynamics repeatedly from the same molecule, even in conditions that can impair ligase activity.
The TIRFM setup is shown in Figure 1. A custom-designed specimen stage is placed over an Olympus IX61 microscope body. 532 nm and 640 nm lasers are introduced from the side and are reflected by tiny elliptical mirrors14 into the high NA objective to achieve critical angle of incidence at the coverslip-water interface. We note that more widespread through-objective TIR using dichroic mirrors or prism-based TIR setups can also be used for this FRET application. The fluorescence image formed by the microscope is split into donor and acceptor images by a dichroic mirror. They are then re-imaged onto two halves of an EMCCD. Additional long-pass emission filters are used to reduce background signal.
Temperature control is essential for acquiring reproducible kinetic data. For temperature control, the objective is separated from the nosepiece of the microscope body to minimize heat transfer, and water from a temperature controlled chiller/heater is circulated through a brass collar that tightly fits around the interior metal beneath the objective jacket. This setup is able to achieve robust temperature control at the coverslip surface between 15 and 50 &#176;C (Figure 2). In this work, the sample temperature was maintained at 24 &#176;C.
The following protocol presents the step-by-step procedure for DNA construction, estimation of DNA shape, single-molecule experiment, and J factor determination.

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			<td height="17" style="height:17px;width:351px;">Small DNA FRAG Extract Kit-100PR</td>
			<td style="width:124px;">VWR</td>
			<td style="width:135px;">97060-558</td>
			<td style="width:183px;"></td>
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		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Acrylamide 40% solution 500 mL</td>
			<td style="width:124px;">VWR</td>
			<td style="width:135px;">97064-522</td>
			<td style="width:183px;"></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Bis-acrylamide 2% (w/v) solution 500 mL</td>
			<td style="width:124px;">VWR</td>
			<td style="width:135px;">97063-948</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">GeneRuler 100 bp DNA Ladder, 100-1000 bp</td>
			<td style="width:124px;">Fermentas</td>
			<td>SM0241</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Mini Vertical PAGE System</td>
			<td style="width:124px;">VWR</td>
			<td style="width:135px;">89032-300</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Syringe filter 0.2um CS50</td>
			<td style="width:124px;">VWR</td>
			<td style="width:135px;">A2666</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Trolox</td>
			<td style="width:124px;">Sigma-Aldrich</td>
			<td style="width:135px;">238813-1G</td>
			<td>triplet state quencher</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Protocatechuic acid (PCA)</td>
			<td style="width:124px;">Sigma-Aldrich</td>
			<td style="width:135px;">08992-50MG</td>
			<td>oxygen scavenging system</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Protocatechuate 3,4-Dioxygenase (PCD)</td>
			<td style="width:124px;">Sigma-Aldrich</td>
			<td style="width:135px;">P8279-25UN</td>
			<td>oxygen scavenging system</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">mPEG-silane, MW 2000 1g</td>
			<td style="width:124px;">Laysan Bio</td>
			<td style="width:135px;">MPEG-SIL-2000-1g</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Biotin-PEG-Silane, MW 3400</td>
			<td style="width:124px;">Laysan Bio</td>
			<td style="width:135px;">Biotin-PEG-SIL-3400-1g</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Avidin, NeutrAvidin Biotin-binding Protein</td>
			<td style="width:124px;">Invitrogen</td>
			<td style="width:135px;">A2666</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Phusion Hot Start High-Fidelity DNA Polymerase</td>
			<td style="width:124px;">New England Biolabs</td>
			<td style="width:135px;">F-540L</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Gel/PCR DNA Fragments Extraction Kit</td>
			<td>IBI Scientific</td>
			<td>IB47020</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Premium plain glass microscope slides</td>
			<td style="width:124px;">Fisher Scientific</td>
			<td style="width:135px;">12-544-1</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">VWR micro cover glass, rectangular, no. 1</td>
			<td style="width:124px;">VWR</td>
			<td style="width:135px;">48404-456</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Fisher Scientific Isotemp 1006s Recirculating Chiller/Heater</td>
			<td style="width:124px;">Fisher Scientific</td>
			<td style="width:135px;"></td>
			<td>temperature control</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Objective Cooling Collar</td>
			<td style="width:124px;">Bioptechs</td>
			<td style="width:135px;">150303</td>
			<td>temperature control</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">KMI53 Biological Micrometer Measuring Stage</td>
			<td style="width:124px;">Semprex</td>
			<td style="width:135px;">KMI53</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">High Performance DPSS Laser 532nm 50mW&#160;</td>
			<td style="width:124px;">Edmund optics</td>
			<td style="width:135px;">NT66-968</td>
			<td>Cy3 excitation</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:351px;">CUBE Fiber Pigtailed 640 nm, 30mW, Fiber, FC/APC Connector</td>
			<td style="width:124px;">Coherent</td>
			<td style="width:135px;">1139604</td>
			<td>Cy5 excitation</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">650 nm BrightLine Dichroic Beamsplitter</td>
			<td style="width:124px;">Semrock</td>
			<td style="width:135px;">FF650-Di01-25x36</td>
			<td>splitting dichroic</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:351px;">LaserMUX Beam Combiner, reflects 514.5, 532, &#38; 543.5 nm lasers, 25 mm</td>
			<td style="width:124px;">Semrock</td>
			<td style="width:135px;">LM01-552-25</td>
			<td>combining dichroic</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Brightline Fluorescence Filter 593/40</td>
			<td style="width:124px;">Semrock</td>
			<td>FF01-593/40-25</td>
			<td>Cy3 emission filter</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">635 nm EdgeBasic LWP longpass Filter, 25 mm</td>
			<td style="width:124px;">Semrock</td>
			<td style="width:135px;">BLP01-635R-25</td>
			<td>Cy5 emission filter</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">EMCCD iXon+</td>
			<td style="width:124px;">Andor Technology</td>
			<td style="width:135px;">DU-897E-CS0-#BV</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">IX51 inverted microscope frame</td>
			<td style="width:124px;">Olympus</td>
			<td style="width:135px;"></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Objective UApo N 100x/1.49 Oil TIRF</td>
			<td style="width:124px;">Olympus</td>
			<td style="width:135px;"></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Immersion oil type-F for fluorescence microscopy</td>
			<td style="width:124px;">Olympus</td>
			<td style="width:135px;">IMMOIL-F30CC</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">2mm Diameter 45&#176; Rod Lens Aluminum Coated&#160;</td>
			<td style="width:124px;">Edmund optics</td>
			<td style="width:135px;">54-092</td>
			<td>miniature mirror</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">1/4&#34; Travel Single-Axis Translation Stage</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">MS-1</td>
			<td>translation of miniature mirror</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">&#216;1&#34; Achromatic Doublet, ARC: 400-700 nm, f=200 mm</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">AC254-200-A</td>
			<td>focusing lens</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Adjustable Mechanical Slit</td>
			<td style="width:124px;">Thorlabs</td>
			<td>VA100</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Dielectric Mirror</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">BB1-E02</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">&#216;1&#34; Achromatic Doublet, f = 100 mm</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">AC254-100-A</td>
			<td>relay lens</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Lens Mount for &#216;1&#34; Optics</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">LMR1</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Dichroic Filter Mount</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">FFM1</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Fixed Cage Cube Platform</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">B3C</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">Kinematic Mount for &#216;1&#34; Optics</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">KM100</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">N-BK7 Plano-Convex Lens, &#216;1&#34;, f = 40 mm</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">LA1422-A</td>
			<td>collimating lens</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">N-BK7 Plano-Convex Lense, &#216;6.0 mm, f = 15 mm</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">LA1222-A</td>
			<td>telescope lens</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:351px;">N-BK7 Plano-Convex Lense, &#216;6.0 mm, f = 150 mm</td>
			<td style="width:124px;">Thorlabs</td>
			<td style="width:135px;">LA1433-A</td>
			<td>telescope lens</td>
		</tr>
	</tbody>
</table>

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.

DNA molecules used for the looping study consist of a duplex region of variable sequence and length and single-stranded overhangs that are complementary to each other. The overhangs, which are 7-base long, can anneal to each other to capture the looped state. Each overhang contains either Cy3 or Cy5 that is linked in the DNA backbone through amidite chemistry. The Cy5-overhang is also linked with biotin-TEG (15-atom Tetra-Ethylene Glycol spacer) for surface immobilization (see Figure 4A). All these modif...

Watch this Scientific Journal Video about Studying DNA Looping by Single-Molecule FRET at JoVE.com

Studying DNA Looping by Single-Molecule FRET

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 ...

studying-dna-looping-by-single-molecule-fret

Research

JoVE Journal

Biology

15.3K Views.  Georgia Institute of Technology. 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.

Video: Studying DNA Looping by Single-Molecule FRET

Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function.   The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid.  Hence the gramicidin channel  current is a reporter of altered properties of the bilayer due to certain compounds.  

Membrane protein function is regulated by the cell membrane lipid composition. This video-article details how to form a patch using bilayer patch electrodes, as well as how to use gramicidin channels as reporters of altered membrane properties.

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein ...

Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically functional inside the cell, nucleic acid-based therapeutics require an efficient intracellular delivery system. One widely adopted approach is to complex DNA with a gene carrier to form nanocomplexes via electrostatic self-assembly, facilitating cellular uptake of DNA while protecting it against degradation. The challenge lies in the rational design of efficient gene carriers, since premature dissociation or overly stable binding would be detrimental to the cellular uptake and therapeutic efficacy. Nanocomplexes synthesized by bulk mixing showed a diverse range of intracellular unpacking and trafficking behavior, which was attributed to the heterogeneity in size and stability of nanocomplexes. Such heterogeneity hinders the accurate assessment of the self-assembly kinetics and adds to the difficulty in correlating their physical properties to transfection efficiencies or bioactivities. We present a novel convergence of nanophotonics (i.e. QD-FRET) and microfluidics to characterize the real-time kinetics of the nanocomplex self-assembly under laminar flow. QD-FRET provides a highly sensitive indication of the onset of molecular interactions and quantitative measure throughout the synthesis process, whereas microfluidics offers a well-controlled microenvironment to spatially analyze the process with high temporal resolution (~milliseconds). For the model system of polymeric nanocomplexes, two distinct stages in the self-assembly process were captured by this analytic platform. The kinetic aspect of the self-assembly process obtained at the microscale would be particularly valuable for microreactor-based reactions which are relevant to many micro- and nano-scale applications. Further, nanocomplexes may be customized through proper design of microfludic devices, and the resulting QD-FRET polymeric DNA nanocomplexes could be readily applied for establishing structure-function relationships. 

We present a novel and powerful integration of nanophotonics (QD-FRET) and microfluidics to investigate the formation of polyelectrolyte polyplexes, which is expected to provide better control and synthesis of uniform and customizable polyplexes for future nucleic acid-based therapeutics.

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically ...

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 ...

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

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. ...

Single-Molecule Imaging of Nuclear Transport

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

Single molecule microscopy approch provided novel insights into nuclear transport.

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

Genomic MRI - a Public Resource for Studying Sequence Patterns within Genomic DNA

Non-coding genomic regions in complex eukaryotes, including intergenic areas, introns, and untranslated segments of exons, are profoundly non-random in their nucleotide composition and consist of a complex mosaic of sequence patterns. These patterns include so-called Mid-Range Inhomogeneity (MRI) regions -- sequences 30-10000 nucleotides in length that are enriched by a particular base or combination of bases (e.g. (G+T)-rich, purine-rich, etc.). MRI regions are associated with unusual (non-B-form) DNA structures that are often involved in regulation of gene expression, recombination, and other genetic processes (Fedorova &amp; Fedorov 2010). The existence of a strong fixation bias within MRI regions against mutations that tend to reduce their sequence inhomogeneity additionally supports the functionality and importance of these genomic sequences (Prakash et al. 2009).
Here we demonstrate a freely available Internet resource -- the Genomic MRI program package -- designed for computational analysis of genomic sequences in order to find and characterize various MRI patterns within them (Bechtel et al. 2008). This package also allows generation of randomized sequences with various properties and level of correspondence to the natural input DNA sequences. The main goal of this resource is to facilitate examination of vast regions of non-coding DNA that are still scarcely investigated and await thorough exploration and recognition.

We present a public computational web site for the analysis of genomic sequences. It detects DNA sequence patterns with various non-random nucleotide compositions. This resource also generates randomized sequences with diverse levels of complexity.

Non-coding genomic regions in complex eukaryotes, including intergenic areas, introns, and untranslated segments of exons, are profoundly non-random in ...

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method makes it possible to count the numbers of protein molecules produced in one cell during sequential, five-minute time windows. It requires a fluorescence microscope with laser excitation power density of ~0.5 to 1 kW/cm2, which is sufficiently sensitive to detect single fluorescent protein molecules in live cells. The fluorescent reporter used in this method consists of three parts: a membrane targeting sequence, a fast-maturing, yellow fluorescent protein and a protease recognition sequence. The reporter is translationally fused to the N-terminus of a protein of interest. Cells are grown on a temperature-controlled microscope stage. Every five minutes, fluorescent molecules within cells are imaged (and later counted by analyzing fluorescence images) and subsequently photobleached so that only newly translated proteins are counted in the next measurement.
Fluorescence images resulting from this method can be analyzed by detecting fluorescent spots in each image, assigning them to individual cells and then assigning cells to cell lineages. The number of proteins produced within a time window in a given cell is calculated by dividing the integrated fluorescence intensity of spots by the average intensity of single fluorescent molecules. We used this method to measure expression levels in the range of 0-45 molecules in single 5 min time windows. This method enabled us to measure noise in the expression of the &lambda; repressor CI, and has many other potential applications in systems biology.

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage CoTrAC

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC), to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method has been used to follow the stochastic expression dynamics of a transcription factor, the &lambda; repressor CI 1.

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells ...

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 ...