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.

<table><tbody><tr><td>Small DNA FRAG Extract Kit-100PR</td><td>VWR</td><td>97060-558</td><td></td></tr><tr><td>Acrylamide 40% solution 500 ml</td><td>VWR</td><td>97064-522</td><td></td></tr><tr><td>Bis-acrylamide 2% (w/v) solution 500 ml</td><td>VWR</td><td>97063-948</td><td></td></tr><tr><td>GeneRuler 100 bp DNA Ladder, 100-1,000 bp</td><td>Fermentas</td><td>SM0241</td><td></td></tr><tr><td>Mini Vertical PAGE System</td><td>VWR</td><td>89032-300</td><td></td></tr><tr><td>Syringe filter 0.2 &amp;mu;m CS50</td><td>VWR</td><td>A2666</td><td></td></tr><tr><td>Trolox</td><td>Sigma-Aldrich</td><td>238813-1G</td><td>triplet state quencher</td></tr><tr><td>Protocatechuic acid (PCA)</td><td>Sigma-Aldrich</td><td>08992-50MG</td><td>oxygen scavenging system</td></tr><tr><td>Protocatechuate 3,4-Dioxygenase (PCD)</td><td>Sigma-Aldrich</td><td>P8279-25UN</td><td>oxygen scavenging system</td></tr><tr><td>mPEG-silane, MW 2,000 1 g</td><td>Laysan Bio</td><td>MPEG-SIL-2000-1g</td><td></td></tr><tr><td>Biotin-PEG-Silane, MW 3,400</td><td>Laysan Bio</td><td>Biotin-PEG-SIL-3400-1g</td><td></td></tr><tr><td>Avidin, NeutrAvidin Biotin-binding Protein</td><td>Invitrogen</td><td>A2666</td><td></td></tr><tr><td>Phusion Hot Start High-Fidelity DNA Polymerase</td><td>New England Biolabs</td><td>F-540L</td><td></td></tr><tr><td>Gel/PCR DNA Fragments Extraction Kit</td><td>IBI Scientific</td><td>IB47020</td><td></td></tr><tr><td>Premium plain glass microscope slides</td><td>Fisher Scientific</td><td>12-544-1</td><td></td></tr><tr><td>VWR micro cover glass, rectangular, no. 1</td><td>VWR</td><td>48404-456</td><td></td></tr><tr><td>Fisher Scientific Isotemp 1006S Recirculating Chiller/Heater</td><td>Fisher Scientific</td><td></td><td>temperature control</td></tr><tr><td>Objective Cooling Collar</td><td>Bioptechs</td><td>150303</td><td>temperature control</td></tr><tr><td>KMI53 Biological Micrometer Measuring Stage</td><td>Semprex</td><td>KMI53</td><td></td></tr><tr><td>High Performance DPSS Laser 532 nm 50 mW</td><td>Edmund optics</td><td>NT66-968</td><td>Cy3 excitation</td></tr><tr><td>CUBE Fiber Pigtailed 640 nm, 30 mW, Fiber, FC/APC Connector</td><td>Coherent</td><td>1139604</td><td>Cy5 excitation</td></tr><tr><td>650 nm BrightLine Dichroic Beamsplitter</td><td>Semrock</td><td>FF650-Di01-25x36</td><td>splitting dichroic</td></tr><tr><td>LaserMUX Beam Combiner, reflects 514.5, 532, &amp;amp; 543.5 nm lasers, 25 mm</td><td>Semrock</td><td>LM01-552-25</td><td>combining dichroic</td></tr><tr><td>Brightline Fluorescence Filter 593/40</td><td>Semrock</td><td>FF01-593/40-25</td><td>Cy3 emission filter</td></tr><tr><td>635 nm EdgeBasic LWP longpass Filter, 25 mm</td><td>Semrock</td><td>BLP01-635R-25</td><td>Cy5 emission filter</td></tr><tr><td>EMCCD iXon+</td><td>Andor Technology</td><td>DU-897E-CS0-#BV</td><td></td></tr><tr><td>IX51 inverted microscope frame</td><td>Olympus</td><td></td><td></td></tr><tr><td>Objective UApo N 100X/1.49 Oil TIRF</td><td>Olympus</td><td></td><td></td></tr><tr><td>Immersion oil type-F for fluorescence microscopy</td><td>Olympus</td><td>IMMOIL-F30CC</td><td></td></tr><tr><td>2 mm Diameter 45&amp;deg; Rod Lens Aluminum Coated&amp;nbsp;</td><td>Edmund optics</td><td>54-092</td><td>miniature mirror</td></tr><tr><td>1/4&quot; Travel Single-Axis Translation Stage</td><td>Thorlabs</td><td>MS-1</td><td>translation of miniature mirror</td></tr><tr><td>&amp;Oslash;1&quot; Achromatic Doublet, ARC: 400-700 nm, f=200 mm</td><td>Thorlabs</td><td>AC254-200-A</td><td>focusing lens</td></tr><tr><td>Adjustable Mechanical Slit</td><td>Thorlabs</td><td>VA100</td><td></td></tr><tr><td>Dielectric Mirror</td><td>Thorlabs</td><td>BB1-E02</td><td></td></tr><tr><td>&amp;Oslash;1&quot; Achromatic Doublet, f = 100 mm</td><td>Thorlabs</td><td>AC254-100-A</td><td>relay lens</td></tr><tr><td>Lens Mount for &amp;Oslash;1&quot; Optics</td><td>Thorlabs</td><td>LMR1</td><td></td></tr><tr><td>Dichroic Filter Mount</td><td>Thorlabs</td><td>FFM1</td><td></td></tr><tr><td>Fixed Cage Cube Platform</td><td>Thorlabs</td><td>B3C</td><td></td></tr><tr><td>Kinematic Mount for &amp;Oslash;1&quot; Optics</td><td>Thorlabs</td><td>KM100</td><td></td></tr><tr><td>N-BK7 Plano-Convex Lens, &amp;Oslash;1&quot;, f = 40 mm</td><td>Thorlabs</td><td>LA1422-A</td><td>collimating lens</td></tr><tr><td>N-BK7 Plano-Convex Lense, &amp;Oslash;6.0 mm, f = 15 mm</td><td>Thorlabs</td><td>LA1222-A</td><td>telescope lens</td></tr><tr><td>N-BK7 Plano-Convex Lense, &amp;Oslash;6.0 mm, f = 150 mm</td><td>Thorlabs</td><td>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 ...

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Research

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Biology

15.4K 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

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Biochemistry

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

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

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

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

Single-Molecule F&ouml;rster Resonance Energy Transfer Methods for Real-Time Investigation of the Holliday Junction Resolution by GEN1

Bulk methods measure the ensemble behavior of molecules, in which individual reaction rates of the underlying steps are averaged throughout the population. Single-molecule F&#246;rster resonance energy transfer (smFRET) provides a recording of the conformational changes taking place by individual molecules in real-time. Therefore, smFRET is powerful in measuring structural changes in the enzyme or substrate during binding and catalysis. This work presents a protocol for single-molecule imaging of the interaction of a four-way Holliday junction (HJ) and gap endonuclease I (GEN1), a cytosolic homologous recombination enzyme. Also presented are single-color and two-color alternating excitation (ALEX) smFRET experimental protocols to follow the resolution of the HJ by GEN1 in real-time. The kinetics of GEN1 dimerization are determined at the HJ, which has been suggested to play a key role in the resolution of the HJ and has remained elusive until now. The techniques described here can be widely applied to obtain valuable mechanistic insights of many enzyme-DNA systems.

Single-Molecule F&#246;rster Resonance Energy Transfer Methods for Real-Time Investigation of the Holliday Junction Resolution by GEN1

Presented here is a protocol for performing single-molecule F&#246;rster resonance energy transfer to study HJ resolution. Two-color alternating excitation is used for determining the dissociation constants. Single-color time lapse smFRET is then applied in real-time cleavage assays to obtain the dwell time distribution prior to HJ resolution.

Bulk methods measure the ensemble behavior of molecules, in which individual reaction rates of the underlying steps are averaged throughout the ...

Genetics

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, ...

Making Precise and Accurate Single-Molecule FRET Measurements using the Open-Source smfBox

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes measurements on freely diffusing biomolecules more accessible. This overview includes a step-by-step protocol for using this instrument to make measurements of precise FRET efficiencies in duplex DNA samples, including details of the sample preparation, instrument setup and alignment, data acquisition, and complete analysis routines. The presented approach, which includes how to determine all the correction factors required for accurate FRET-derived distance measurements, builds on a large body of recent collaborative work across the FRET Community, which aims to establish standard protocols and analysis approaches. This protocol, which is easily adaptable to a range of biomolecular systems, adds to the growing efforts in democratising smFRET for the wider scientific community.

This article provides step-by-step instructions for making fully-corrected accurate FRET measurements on individual, freely diffusing biomolecules using the open-source, inexpensive smfBox, from switch on, through alignment and focusing, to data collection and analysis.

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes ...

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. ...

Automated Two-dimensional Spatiotemporal Analysis of Mobile Single-molecule FRET Probes

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. It has been used in a wide range of biophysical and molecular biological experiments, including the measurement of molecular forces, characterization of conformational dynamics of biomolecules, observation of intracellular colocalization of proteins, and determination of receptor-ligand interaction times. In a widefield microscopy configuration, experiments are typically performed using surface-immobilized probes. Here, a method combining single-molecule tracking with alternating excitation (ALEX) smFRET experiments is presented, permitting the acquisition of smFRET time traces of surface-bound, yet mobile probes in plasma membranes or glass-supported lipid bilayers. For the analysis of recorded data, an automated, open-source software collection was developed supporting (i) the localization of fluorescent signals, (ii) single-particle tracking, (iii) determination of FRET-related quantities including correction factors, (iv) stringent verification of smFRET traces, and (v) intuitive presentation of the results. The generated data can conveniently be used as input for further exploration via specialized software, e.g., for the assessment of the diffusional behavior of probes or the investigation of FRET transitions.

This article presents a method for spatiotemporal analysis of mobile, single-molecule F&#246;rster resonance energy transfer (smFRET)-based probes using widefield fluorescence microscopy. The newly developed software toolkit allows the determination of smFRET time traces of moving probes, including the correct FRET efficiency and the molecular positions, as functions of time.

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. ...

Bioengineering

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

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

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

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

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