The overall goal of the following method article is to demonstrate a protocol useful to study RNA interactions with proteins or other factors using optical tweezers coupled with confocal microscopy. Over the past decade, several sophisticated techniques have been developed to investigate the properties of single macromolecules. One of these techniques is single molecule optical tweezers.
With the new fluorescence assisted optical tweezers, not only forces required for unfolding of RNA molecules, as well as binding events during translation can be monitored in detail. Here we focus on measurements in which RNA molecules are stretched with and without transacting factors that regulate translation. We also illustrate how the technique can be applied to monitor translation using labeled ribosomes.
The protocol will be demonstrated by two graduate students in my group, Lukas Pekarek and Stefan Park. In this video, we are going to guide you through the setup and execution of an optical tweezers experiment. We also outline the simple data analysis workflow.
The advantage of this technique is that it is straightforward and robust. A molecule is tethered between two beads, strapped in the focus of laser beams, and change and force upon displacement can be precisely measured in so-called force-ramp experiments. Secondary structures inside a tether unfold at a certain force depending on the internal energy of the molecule.
This dynamic transition between multiple structures can be further investigated with constant force measurements. Furthermore, the parallel use of fluorescent imaging can be used to visualize binding events in real time. A distinct force-distance profile of a tether might vary after binding of potential transacting factors that influence its stability.
First, the RNA of interest has to be cloned into a DNA vector. Upstream and downstream of this sequence, additional parts, the handles are needed to tether the final DNA-RNA construct between the beads. After successful cloning and amplification of the plasmid, three different PCR reactions are performed.
For each PCR reaction, mix the reagents according to table one in the protocol. Run the PCR in 50 microliter aliquots in an appropriate thermal cycler. The first one results in a double stranded DNA of the whole construct with the T7 promoter for subsequent in-vitro transcription.
The second PCR produces the five prime handles. Whereas the last reaction results in the three prime handles. Both 5 prime and 3 prime handles need to be labeled according to the used beads.
The three prime handles are labeled during the PCR by using a digoxigenin labeled reverse primer, and the five prime handles have to be labeled in an extra step through biotinylation. The three prime biotinylation of the five prime handles is performed using T 40 and a polymerase. The reaction is performed for one to two hours at room temperature.
Next, carry out the in-vitro transcription using T7 polymerase. Incubate the reaction at 37 degree for two to four hours, depending on the length of the RNA. In one last step, the handles are annealed with the RNA from the in-vitro transcription to obtain the final DNA-RNA construct with the single stranded region of interest between the double stranded handles.
To anneal the desired RNA-DNA hybrid, mix the handles and the RNA in annealing buffer. Heat, the annealing mixture up to 85 degree for 10 minutes, and then slowly cool down the sample to four degree. Mix the annealed sample with 1/10th of volume of three molar sodium acetate, and three volumes of ice cold ethanol and incubate minus 80 degree for at least an hour.
Centrifuge the samples at 15, 000 xg for 30 minute at four degree. Discard the supernatant and dry the pellet under vacuum. Small aliquots of the resuspended pellet are frozen in liquid nitrogen and kept at minus 80 degree until they are used.
First remove the bleach out of the syringes and fill in one mL of RNase free water. Add 50 microliter of 0.5 molar sodium thiosulfate and wash the system with one bar. Be careful that the microfluidic system never runs dry to avoid air bubbles in the system.
Next, discard the remaining sodium thiosulfate solution and replace the syringes with new ones. Then wash again with at least 0.5 mL of RNase free water. To set up the optical system of the machine, put two drops of immersion oil onto the objective.
Put the flow cell in place and rise the objective until the liquid touches the flow cell. Then put two drops of immersion oil on top of the flow cell. Now turn on the trap steering unit and the trapping laser.
The objective is adjusted using the Z finder tool of the diagnostic cameras of the machine. By turning the micro screw, the Z axis is adjusted to the middle of the chamber between the second and the third reflection. Where the refraction rings are the biggest.
Switch the diagnostic cameras to moon mode and reduce the trapping laser to around 50%Then lower the condenser and adjust its position so you see approximately 10 light bands. The measuring chamber has five channels, and before the experiment, different setup possibilities should be considered. For a simple force-ramp experiment, beads-buffer-beads is a convenient channel arrangement.
For measurements with the fourth compound, like fluorescent ribosomes or transacting factors, beads-beads-buffer factor is a better arrangement, but it makes it harder to catch beads. An aliquot of the annealed construct is incubated with three microliter AD bead suspension, one microliter RNase inhibitors, and eight microliter of the assay buffer at room temperature for 10 to 20 minutes. Then the sample is diluted in 500 microliter assay buffer and put into the respective channel.
For the second kind of beads, 0.8 microliter of assay beads are mixed with one mL of assay buffer and administered into the corresponding channel. The channels are opened and washed with low pressure. Now turn on the trapping laser.
To catch beads, the lasers are moved apart from each other and an AD bead is caught in trap one. Next the stage is moved to the SA bead channel and a bead is caught in trap two. To avoid losing the beads, or to catch a second one in the same trap, the stage is moved to the buffer channel and all channels are closed.
In the buffer, a trap calibration is performed. The caught beads should be set as visual templates for subsequent measurements. And the similarity score should be kept above 90%between measurements to guarantee comparability.
Finally, the beads are moved closer together and one can start to catch a single tether. This is done moving the beads close for a few seconds and then slowly depart the beads again. A tether formation results in an increase of measured force upon pulling the two beads away from each other.
The number and quality of tethers can be checked by finding the overstretching plateau and compare the trajectory with the freely jointed chain or worm-like chain model. The overstretching plateau should be between 50 to 60 piconewton for a single tether. To perform the fluorescence measurements, turn on the confocal lasers and photon count on unit and optical tweezers machine.
Afterwards, turn on the excitation laser of desired wavelength and the software interface and set the power of the laser to 5%or higher, depending on the fluorophore. Now imaging can be started by using the image or kymograph functions of the software. In order to get well focused images, make sure that focal plane of the confocal microscope and the optical traps are aligned.
For this purpose, autofluorescence of the polystyrene beads in the blue laser channel can be employed. Move up and down with the focal plane of the optical traps until the autofluorescence of the beads reaches its highest diameter. At this position, the fluorescence signals taking place at the tethered molecule can be detected.
For both functions it is important to select the appropriate regions of interest. Throughout the measurement buffer composition can be easily changed by either moving the beads to different channels, or by changing the buffer supplied in the microfluidic system. To avoid photobleaching, always turn off the excitation laser when not measuring.
The raw data output from the device often contains a lot of noise. Therefore, in order to further analyze scattered data, one has first to pre-process them. The first step is to down sample and filter the data with the low pass Butterworth filter, good results were obtained.
For force-ramp experiments, the unfolding events have to be marked at the corresponding folded and unfolded regions of the first distance curve can be fitted using worm-like chain or freely jointed chain models. For the constant force data, the distance over time can be plotted and it is useful to generate a histogram to quantify the predominant conformation at a given force. Here it is shown that the filter parameters are crucial to distinguish different folding states.
For better understanding what can be achieved with this technique, some representative results are shown. This is how a standard force-distance curve from a force-ramp experiment looks like. In this case, we studied the SARS-coronavirus-2 frame shifting element.
We observed unfolding in multiple steps around 15 piconewton. When the direction is reversed and the beads are getting closer again, the tether refolds back at slightly lower force. When we measured the same RNA in presence of the zinc finger antiviral protein ZAP, a similar pattern of unfolding was observed.
But the secondary structure of the RNA did not refold. This is a good example to show what big influence protein binding can have on RNA kinetics. Looking at constant force data the unfolding kinetics can be further investigated.
The distance over time is plotted for different forces and the histogram of the different observed states is added on the right. At 10 piconewton, the RNA is completely in the folded state. At 11.5 piconewton it switches between states, and at 13 piconewton the RNA is permanently in an unfolded state.
However, when the SARS-coronavirus-2 frame shifting element was measured together with ZAP, the RNA was completely in an unfolded state at 11 piconewton, and even showed very little transition towards a folded state at 10 piconewton. This indicates that ZAP binds to the single stranded frame shifting element and prevents the formation of a secondary structure. Finally, we will look at some optical tweezers data coupled with confocal microscopy.
In this experiment, a fluorescent dye was used that labels double stranded nucleic acids, non specifically with increasing force. The kymograph shows that as the beads are moved apart from each other, the fluorescence intensity increases and it completely disappears once the tether is stretched too much and breaks. In the last figure fluorescently labeled ribosomes were added through channel four.
In this experiment specific ribosome binding to the RNA tether was observed. We hope this video gave you an insight into optical tweezers and what can be performed this incredible technique.
