Hybrid Ensemble and Single-molecule Assay to Image the Motion of Fully Reconstituted CMG

We study the motion dynamics of different components of a eukaryotic replisome at the single molecule level, in order to obtain a deep quantitative understanding of how cells can duplicate their entire genomes every cell cycle. In a breakthrough in 2015, eukaryotic DNA replication was fully reconstituted in vitro from purified protein components, and this gave us a lot of control over the system. And since then, it's been used extensively to answer questions about different stages of DNA replication with increasing temporal and spatial resolution.

And this was done using complementary approaches such as cryo-EM and single molecule biophysics. In the single molecule field, one of the biggest challenges with this system is the number of proteins involved and also the concentrations at which these components are required in order to maximize the efficiency of the overall reaction, because this can complicate the single molecule imaging. The protocol described here introduces a hybrid approach in which a protein complex is first assembled in an efficient manner using ensemble biochemistry before then being introduced and studied in the single molecule setting.

This avoids the introduction of too high protein concentrations at the single molecule level. We illustrate this approach for studying the behavior of the replicative helicase CMG on DNA at the single molecule level following its assembly via the origin-based pathway. But this approach can also be used to study the dynamics of many other types of DNA binding protein complexes.

Proceed to perform the reaction after functionalizing both ends of the linear DNA substrate. First, take for S400 spin columns and vortex them for at least 30 seconds to resuspend the resin. Then centrifuge them for one minute at 735 G to remove the storage buffer.

Transfer the columns to clean 1.5 milliliter tubes. Immediately add 100 microliters of the doubly functionalized linear DNA solution to each column. Centrifuge the columns for two minutes at 735 G to flow the DNA through the column while retaining the unincorporated nucleotides.

After pooling together the flow through containing the DNA from the four columns, measure the volume with a pipette. Next, vortex M280 streptavidin-coated magnetic beads for 30 seconds to resuspend them. Transfer four milligrams of the resuspended magnetic beads to a clean 1.5 milliliter tube.

Place the tube in a magnetic rack for one minute to collect the beads before removing the storage buffer. Resuspend the beads in one milliliter of buffer A by vortexing for five seconds before incubating the beads at room temperature for five minutes. Collect the beads by placing the tube in a magnetic holder for one minute.

After removing buffer A, resuspend the beads in 400 microliters of 2x buffer A by pipetting. Then, add 400 microliters of the functionalized DNA solution to the beads and mix gently by pipetting. Incubate the bead-DNA mixture overnight at four degrees celsius with end over end rotation.

The next day, use the magnetic rack to remove the supernatant before calculating the total amount of DNA bound to the magnetic beads. Then, wash the beads sequentially with buffer B and buffer C.Resuspend the beads in 300 microliters of buffer C and make four 75 microliter aliquots. To begin, take one 75 microliter aliquot containing one milligram of DNA bound streptavidin-coated magnetic beads and remove the supernatant with the magnetic rack.

Wash the beads with 200 microliters of loading buffer. Then, resuspend the beads in 75 microliters of loading buffer and mix gently by pipetting. Next, add the origin recognition complex at a final concentration of 37.5 nanomolar to the bead bound DNA and incubate the reaction for five minutes at 30 degrees celsius with 800 revolutions per minute agitation.

Add Cdc6 at a final concentration of 50 nanomolar and incubate the reaction as demonstrated before. Then, add Mcm2-7/Cdt1 at a final concentration of 100 nanomolar and incubate the reaction for 20 minutes as demonstrated previously. After that, add Ddk at a final concentration of 100 nanomolar and incubate similarly for 30 minutes.

After removing the supernatant, wash the bead bound DNA containing phosphorylated Mcm2-7 hexamers with 200 microliters of high salt wash buffer. Mixed by pipetting, ensuring the beads are fully resuspended. Next, remove the buffer, and wash the beads once with 200 microliters of CMG buffer.

To assemble and activate the fluorescently labeled CMG onto bead bound DNA, remove the CMG buffer and resuspend the DNA bound beads in 50 microliters of CMG buffer supplemented with five millimolar ATP. Next, mix all the components mentioned on the screen in one tube and place it on ice. Immediately add the resuspended bead bound DNA to the protein mix.

Incubate the reaction for 15 minutes at 30 degrees celsius with 800 RPM agitation. Wash the beads sequentially with HSW buffer and CMG buffer. After removing the buffer, resuspend the CMG containing DNA bound magnetic beads in 200 microliters of elution buffer.

Then, incubate at room temperature for one hour with 800 RPM agitation. Place the tube in a magnetic rack and allow the beads to be collected for five minutes. Carefully collect the supernatant containing the eluted DNA-CMG complexes without disrupting the settled beads and transfer it to a new tube.

To ensure that no beads remain in the solution, place the collected supernatant again in a magnetic rack and keep for another five minutes. Carefully collect the supernatant and transfer it to a new tube. Add 1, 400 microliters of CMG buffer to the 200 microliters of supernatant.

The sample is now ready for single molecule imaging. Perform the single molecule imaging in a commercial setup combining dual beam optical tweezers with confocal microscopy. Use inject channel one for bead trapping, channel two for DNA protein complex binding, channel three for checking the presence of CMG, and channels four and five as protein reservoirs and buffer exchange locations.

Flow all solutions into the flow cell at a constant pressure of 0.5 bar in the injection port. Then, turn off the flow in channels four and five. After initially flowing all solutions, reduce the pressure to 0.2 bar.

Move trapping lasers to channel one until one bead is caught in each optical trap. Move trapped beads to channel two by moving the joystick while pressing the trigger. Using the joystick without pressing the trigger, fish a DNA-CMG complex by moving the right bead towards and away from the left bead until a DNA tether is trapped.

Move the beads to channel three and immediately stop the flow in all channels. Adjust the 561 nanometer laser power to four microwatts in the excitation laser panel and then take a one frame test scan of the DNA in channel three. If present, CMG will appear as two dimensional diffraction limited spots.

In this case, move the DNA tether to either channel four or five for imaging. Otherwise, turn the flow back on, discard the beads and repeat the steps. In channels four or five, input two piconewtons in the force spectroscopy panel, and click on the enable button to start a force clamp.

Once the initial tension reaches two piconewtons, disable the force clamp before imaging by clicking again on the enable button in the force spectroscopy panel. Click the start scan button in the image scan panel. Image fluorescent CMG every five seconds with a 561 nanometer laser at a power of four microwatts as measured at the objective.

In an aggregation free reaction, CMG appeared as discreet, symmetrical diffraction limited spots sparsely crowding the DNA. On the contrary, aggregates are less discreet, sometimes asymmetrical blobs crowding a larger length of the DNA. If the assay was successfully executed and high purity of the purified proteins was achieved, long range motion of CMG in the presence of ATP could be observed.