Studying DNA Looping by Single-Molecule FRET

The overall goal of the following experiment is to investigate how the looping dynamics of double stranded DNA are affected by the intrinsic shape of DNA without the use of DNA binding proteins. This is achieved by incorporating dye molecules into double stranded DNA fragments of different curvatures so that DNA looping can be monitored by fluorescence resonance, energy transfer, or fret. The reversible looping and un looping events from the single DNA molecules can then be observed by total internal reflection microscopy.

The looping rate of the DNA can then be extrapolated from the time trajectories of each single molecule fluorescence. Ultimately, the looping probability of the DNA in relation to the intrinsic shape of the fragment can be measured. The main advantage of this DNA looping assay is that it doesn't rely on A DNA binding protein, which might affect the looping kinetics of A DNA.

The simple protocol is about using a technique called fluorescence resonance energy transfer, and PCR-based DNA synthesis. To measure the looking probability of DNA Begin by designing globally curved DNAs by repeating a 10 MER sequence. For example, this representative sequence is a 186 base pair curved DNA, where X is a random extra base and the sequences flanking the repeating 10 mers sequence are adapter sequences.

Next, perform PCR with primers one and two with fret donor SI three labeling of primer two at the five prime end. Then perform PCR with primers three and four with fret acceptor SCI five labeling through the backbone and the biotin linker at the five prime end of primer three. After purifying the PCR products using a PCR cleanup kit, mix the SCI three labeled and the SCI five labeled products in a buffer for strand exchange at final concentrations of 0.4 micromolar and 0.1 micromolar respectively.

Then incubate the strands at 98.5 degrees Celsius for two minutes, gradually cooling to five degrees Celsius with a ramp rate of 0.1 degrees Celsius per second, and then incubating at five degrees Celsius for two hours. To exchange the strands to prepare the flow cell, use a drill press and diamond drill bits to create six to seven pairs of holes along the two opposite edges of a three inch by one inch glass slide. When the holes have all been drilled, rub the slide under flowing water to remove any visible glass powder.

Place the slides upright in a glass jar and fill it with distilled water. Sonicate the jar for 15 minutes and then transfer the slides into a glass jar dedicated for acetone cleaning. Fill the jar with acetone and sonicate the slides in acetone for another 15 minutes.

Next, spray the slides with ethanol and then with water to rinse them and then transfer the slides to a polypropylene jar. Fill the polypropylene jar with five molar potassium hydroxide and then sonicate the slides for 15 more minutes after the last wash, rinse the slides in distilled water, followed by another 15 minutes. Sonication then after cleaning cover slips.

In the same way, dispense 80 microliters of freshly prepared PEG solution onto each slide, and then gently lower a cover slip over the peg. After 45 minutes, use tweezers to remove the cover slips from the slides, rinse the slides and the cover slips with copious amount of distilled water. Then dry them in a desiccate when the cover slips and slides are dry, place thin strips of double stick tape across the slide to form channels, a line a cover slip over the strips and firmly press on the cover slip to form liquid tight channels.

Then use five minute epoxy to seal the edges of the channels to immobilize the molecules for microscopy. First, inject 15 microliters of neu tradin solution into one channel. After two minutes, rinse the channel with 100 microliters of T 50 buffer and then inject 50 microliters of DNA sample into the channel.

After five minutes, rinse away the unbound DNA with 100 microliters of T 50 buffer, and then fill the channel with an imaging buffer that contains the oxygen scavenging system. Now place immersion oil onto the microscope objective, and then use specimen clips to affix the flow cell to the microscope stage. Turn on the 532 nanometer laser.

Use the live view of the fluorescent images on the monitor to fine. Adjust the focus. Begin the data acquisition with the 532 nanometer laser on.

Stop the data acquisition when most of the molecules have photo bleached to process and analyze the images, use a MATLAB script to look through all the single molecule time traces that show multiple transitions between the high and low fret signals. Identify the looped and unloop states, and then find the threshold that separates the two distributions by determining the intersection between the two fitted Gaussian curves. Next, calculate the F front efficiency by dividing the sci-fi intensity by the sum of the SCI three and sci-fi intensities.

Assigning the loop states with high fret values and the unloop states with low fret values. Then using a MATLAB script, analyze the cumulative number of molecules or NFT that looped that is that reached the high fret state at different time lapses. Since the start of data acquisition, the looping rate K sub loop can then be extracted by fitting the NFT with an exponential function.

If NFT increases bi, basically it can be fitted with a double exponential function as illustrated in the equation. In this case, K sub loop is obtained from this equation to determine the J factor flow. 20 microliters of 30 to 50 picaMolar biotin sci five oligo or primer three into one of the nut travain coated channels as just demonstrated.

Next, rinse the channel with 100 microliters of T 50 to wash away unbound oligos, and then flow freshly prepared imaging buffer supplemented with PS three oligo into the chamber. Keep the 532 nanometer laser on and then briefly turn the 640 nanometer laser on to identify the locations of any surface bound sci-fi oligos. Then turn off the 640 nanometer laser and start monitoring the fret signal.

Use a MATLAB script to analyze the number of molecules that start in the unbound state. That is with a low sci five intensity, but later turn into the IL state. That is with a high scifi intensity as a function of time from the scifi intensity traces plot this number of ane molecules versus time fitting the curve with a single exponential function to obtain the ealing rate.

K sub anil, repeat the experiment at different SI three oligo concentrations to confirm the linearity between the an kneeling rate and the reactant concentration. Extract the second order an kneeling rate constant K prime sub ail from the slope. Finally, calculate the J factor where K sub loop is the looping rate measured under the same buffer conditions to test the effect of the intrinsic curvature of double stranded DNA on looping.

In this experiment, one straight and one curved 186 base pair. Double stranded DNAs were each constructed compared to straight DNA. Curved DNA was determined to produce more high fret events during the same time period.

The curved DNA also looped four times more frequently than the straight DNA during the same acquisition time, demonstrating that the intrinsic curvature of the DNA can significantly affect the looping dynamics. In this final analysis step, the J factor was calculated by dividing the looping rate by the first order, a kneeling rate constant, and was determined to be 61 plus or minus three ano molar for the straight DNA and 265 plus or minus 48 nanomolar for the curve DNA in agreement with previous findings. Following this procedure, the effects of accident factor on the looping mobility such as temperature, unstrained and viscosity can be studied.

This protocol will be useful not only for studying the polymer physics of DNA, but also for demonstrating the power of single molecule fluorescence to beginning research students or lay audience.