We thank Anuja Kibe and Jun. Prof. Redmond Smyth for critically reviewing the manuscript. We thank Tatyana Koch for expert technical assistance. We thank Kristyna Pekarkova for the help with recording experimental videos. The work in our laboratory is supported by the Helmholtz Association and funding from the European Research Council (ERC) Grant Nr. 948636 (to NC).

1. Sample preparation
<ol>
	<li>Clone the sequence of interest into the vector containing the Lambda DNA fragments, which serve&#160;as the handle sequences (Figure 2)43,50.</li>
	<li>First generate a DNA template for subsequent in vitro transcription via PCR (Figure 2B; reaction 1). At this PCR step, the T7 promoter is added in the 5&#39; end of the sense DNA molecule32,33,43,50. Set the PCR reaction according to Table 1. Run the PCR in 50 &#181;L aliquots with appropriate cycles in the thermocycler.</li>
	<li>Prepare the handles by two separate PCR reactions (Table 1, Figure 2B; reaction 2 and 3). First, generate the 5&#39; handle by PCR. Then, generate the 3&#39; handle and simultaneously label it with digoxigenin by using a 5&#39; digoxigenin-labeled primer32,33,43,50.</li>
	<li>After the PCR, purify the DNA using silica spin columns.</li>
	<li>Carry out the in vitro transcription reaction using T7 RNA polymerase (Table 2)32,33,43,50. Incubate the reaction at 37 &#176;C for 2-4 h depending on the length of the RNA. Next, add DNase I to the reaction and incubate at 37 &#176;C for 30 min to digest the DNA template. Purify the RNA using silica spin columns.</li>
	<li>During the labeling reaction of the 5&#39; handle (Table 3), add biotin-16-dUTP at the 3&#39; end of the handle by T4 DNA polymerase38,50. Perform the reaction at room temperature for 1-2 h. Afterwards, purify the DNA using silica spin columns. 
	NOTE: Since the 5&#39; handle must be labeled at its 3&#39; end (Figure 2B), the labeling cannot be performed during the PCR.</li>
	<li>Mix the components mentioned above - 5&#39; handle (3&#39; labeled with biotin), 3&#39; handle (5&#39; labeled with digoxigenin), and RNA - in a 1:1:1 molar ratio in annealing buffer (80% formamide, 400 mM NaCl, 40 mM HEPES, pH 7.5, 0.5 mM EDTA, pH 8), to obtain the desired RNA/DNA hybrid (Table 4). Heat the annealing mixture up to 85 &#176;C for 10 min and then slowly cool down to 4 &#176;C.</li>
	<li>Mix the annealed sample with 1/10 of volume of 3 M sodium acetate (pH 5), 3 volumes of ice-cold ethanol and incubate at -80 &#176;C for at least 1 h or at -20 &#176;C overnight.</li>
	<li>Centrifuge the samples at 15,000 &#215; g for 30 min at 4 &#176;C. Discard the supernatant and dry the pellet (usually not visible) under vacuum.</li>
	<li>Finally, resuspend, the pellet in 50 &#181;L of RNase-free water and make aliquots. Store the aliquots at -80 &#176;C until used. For short term storage, the samples can be also stored at -20 &#176;C.</li>
</ol>
2. Instrument setup
NOTE: The following protocol is optimized for the commercial optical tweezers instrument C-Trap from LUMICKS company. Therefore, adjustments to the presented steps might be necessary while using other optical tweezers instruments. If not used, the microfluidics system of the machine is kept in bleach (sodium hypochlorite solution) and must be washed before use.
<ol>
	<li>Discard the bleach and fill the syringes with 1 mL of RNase-free water.</li>
	<li>Add 50 &#181;L of 0.5 M sodium thiosulfate to at least 1 mL of the RNase-free water and thoroughly wash the system (1 bar, at least 0.5 mL) to eliminate the remaining bleach in the system.</li>
	<li>Discard the sodium thiosulfate solution from the syringes. Replace syringes with fresh ones and wash the system with at least 0.5 mL of RNase-free water. 
	NOTE: Be careful, that the microfluidics system never runs dry to avoid air bubbles in the system.</li>
	<li>Put 2 drops of immersion oil (refractive index of 1.33) or approximately 70 &#181;L of water on top of the objective.</li>
	<li>Place the flow cell inside the holding frame in its position.</li>
	<li>Put 2 drops of immersion oil (refractive index of 1.51) on top of the flow cell.</li>
	<li>Turn on the laser device in the tweezers machine. Once it is running, turn on the trapping laser in the software interface at 100%.</li>
	<li>Using diagnostic cameras (Z-finder), adjust the Z-axis to the middle of the chamber between the second and the third reflections (interfaces) where the refraction rings are the biggest, by turning the micro screw. 
	NOTE: Each time the objective is moved closer to the measuring chamber and the focal plane of the objective crosses the interface between two phases, a reflection can be recognized in the Z-finder mode. There are 4 interfaces possible: (i) water/immersion oil and bottom glass (ii) bottom glass and buffer inside the chamber (iii) buffer inside the chamber and top glass (iv) top glass and immersion oil for condenser.</li>
	<li>Adjust the condenser position (set trapping laser to approximately 50%) so the condenser touches the immersion oil on top of the measuring chamber.</li>
	<li>Adjust the focus by moving slowly down/up with the condenser, so approx. 10 light bands are shown in the moon mode (diagnostic cameras).</li>
</ol>
3. Sample measurement
<ol>
	<li>Incubate anti-digoxigenin-coated beads (AD) with the sample constructs (3 &#181;L of 0.1% (w/v) AD bead suspension + 4 &#181;L of sample) and with 1 &#181;L of RNase inhibitors and 8 &#181;L of the assay buffer (300 mM KCl, 5 mM MgCl2, 20 mM HEPES, pH 7.6, 0.05% Tween 20, 5 mM DTT) at RT for 10-20 min. After the incubation, dilute the sample in 500 &#181;L of assay buffer. 
	NOTE: It is recommended to add oxygen scavengers, particularly during fluorescence measurements to the buffer in order to prevent oxidative damage. Here oxygen scavenger system containing glucose (8.3 mg/mL), glucose oxidase (40 U/mL) and catalase (185 U/mL) was used.</li>
	<li>Mix 0.8 &#181;L of 1% (w/v) streptavidin-coated (SA) beads with 1 mL of assay buffer.</li>
	<li>Discard water from the syringes and fill the syringes with respective suspensions/solutions. Wash for at least 2 min at approximately 1 bar, and then start catching beads. 
	NOTE: Depending on the experimental set-up, different channel arrangements may be used (Figure 3). Typically, one flow channel is filled with anti-digoxigenin beads carrying the RNA molecule. A second channel is filled with the streptavidin-coated beads. Buffer channel is used to form the tethers. A fourth channel can be employed to load the RNA binding protein (Figure 3C), or alternatively RBP can be added directly in the buffer channel (Figure 3B).</li>
	<li>To capture the beads, move the optical traps apart from each other. First move to the AD channel and catch an AD-bead in trap 1. Next, move the stage to the SA-channel and catch a single SA bead by trap 2. 
	NOTE: Try to stay at the interface of the buffer and bead channels to avoid losing the already caught bead, or to prevent catching multiple beads by the same trap.</li>
	<li>Once the beads of the right size are captured, move to the buffer channel and stop the laminar flow. Next, perform force calibration to check trap stiffness. The respective stiffness values should not differ in the x/y axis by more than 10-15%. 
	NOTE: Adjust the laser power or the laser split between the traps according to bead size. Force calibration does not have to be done for every bead pair as long as the bead templates match (similarity score &#62; 0.9). However, it should be performed regularly, or at least every time assay conditions are changed.</li>
	<li>Start fishing for a tether by moving the beads close to each other, waiting for a few seconds, and then moving them back apart, repeat until a tether is formed. A tether formation results in an increase of measured force upon pulling the two beads away from each other. 
	NOTE: To avoid formation of multiple tethers, the beads should not be moved too close. Upon catching a tether between the two beads, tether quality can be checked by finding the overstretching plateau. The plateau should be between 50 to 60 pN for a single tether.</li>
	<li>Upon obtaining a tether, start the measurement. Depending on the phenomenon studied different measurement setups should be chosen (Figure 1B-D). 
	NOTE: Usually at the beginning of the experiment, a force-ramp experiment is conducted to check the tether quality and probe the behavior. Afterward, one may also start the constant-force or constant-position experiments to study the state transitions further. Once sufficient number of measurements have been performed on an RNA sample to determine its behavior, labeled factors can be added to the system to perform confocal measurements.</li>
	<li>To perform fluorescence measurements, turn on the confocal lasers and photon counter unit in the optical tweezers instrument.</li>
	<li>Turn on the excitation laser of desired wavelength in the software interface and set the power of the laser to 5% or higher, depending on the fluorophore. 
	NOTE: While not measuring lower the power setting of the excitation laser to 0% to avoid excessive photodamage to the sample.</li>
	<li>Start imaging the sample by using image functions of the software. 
	NOTE: In order to get well-focused images, the focal plane of the confocal microscope and optical traps have to be aligned. For this purpose, autofluorescence of the polystyrene beads in the blue laser channel can be employed. The focal plane of optical traps is moved up or down in the z-axis until the image of beads reaches its highest diameter. At this position, the fluorescence signal from the molecule tethered between the beads can be measured.</li>
	<li>To use the kymograph function, specify the x-y position of the kymograph axis so that it allows detection of the tether between the beads.</li>
	<li>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 microfluidics system.</li>
</ol>
4. Data analysis
<ol>
	<li>Raw data pre-processing
	<ol>
		<li>By using a simple script, downsample the raw data enough to (i) allow faster subsequent data processing but (ii) still contain all the critical information (Figure 4A). Usually, 100-5000 Hz is suitable for this purpose. 
		NOTE: The data gathering frequency in optical tweezers experiments is often higher than it is necessary for the analysis - in the presented experiments, the data gathering frequency is set to 78 125 Hz by default. Since storage space is limited, it is convenient and timesaving to reduce the sampling rate of the data. Here, the raw data were downsampled by a factor of 30.</li>
		<li>Next, employ a signal filter to reduce the high frequency measurement noise from the signal (Figure 4A). Adjust the filter degree and cut-off frequency parameters accordingly to optimize data output of different experiments (Figure 5). 
		NOTE: Amongst signal filters, Butterworth filter51 is one of the most widely used. A custom-written python script allowing the pre-processing of raw data is provided in the supplementary data. Downsampling and signal filtering parameters (cut-off frequency, filter degree) need to be optimized for different experiments.</li>
	</ol>
	</li>
	<li>For force-ramp data analysis, use the following steps.
	<ol>
		<li>Mark the steps either manually by finding corresponding points on the force trajectory plot or by using custom-written scripts. Unfolding steps are characterized by a sudden drop in force combined with an increase in distance in the force-distance (FD) curve.</li>
		<li>Once unfolding events are marked, fit different regions of the FD curve using appropriate models (Figure 4D). 
		NOTE: For the region before the first unfolding step, the tether can be considered &#34;double-stranded&#34; and is commonly fit using an extensible Worm-like-chain model (WLC)47,52,53. The parts after the first unfolding event are considered a combination of double-stranded nucleotides (handles) and single-stranded nucleotides (unfolded RNA molecule). Therefore, data fitting&#160;is more complex - usually a combination of 2 WLC models or WLC and Freely-jointed chain (FJC) models are employed36,39,52. The extensible WLC model has two main fit parameters the contour length (LC) and the persistence length (LP). Contour length corresponds to the length of the fully stretched molecule and persistence length defines the bending properties of the molecule of interest. The model can be described with the following equation (1). WLC can be used to model the behavior of both folded as well as unfolded regions, although for each of these a separate model with different parameters has to be employed. 
		(1) <img alt="Equation 1" src="/files/ftp_upload/62589/62589eq01.jpg" /> 
		where x is extension, LC is contour length, F is force, LP is persistence length, kB is Boltzmann constant, T is Thermodynamic temperature, and S is stretch modulus. 
		The second model called Freely-jointed chain (FJC) is commonly used to describe behavior of unfolded single stranded regions. It uses similar parameters of the polymers but treats each unit of the &#34;chain&#34; as a rigid rod, here corresponding to the nucleotides of the unfolded single stranded region. The following equation (2) describes this model: 
		(2) <img alt="Equation 2" src="/files/ftp_upload/62589/62589eq02.jpg" /> 
		NOTE: Our lab has recently developed an algorithm that allows batch processing of the raw force-ramp data called &#34;Practical Optical Tweezers Analysis TOol (POTATO)54.&#34; The algorithm downsamples and filters the data, then it identifies possible unfolding steps and finally performs data fitting. The POTATO is built in a user-friendly graphical user interface (GUI) (https://github.com/REMI-HIRI/POTATO).</li>
	</ol>
	</li>
	<li>Process constant-force data as follows: 
	NOTE: The following instructions can be analogically applied on constant-position data.
	<ol>
		<li>For the constant-force data, plot the distance over time (Figure 5). A histogram showing the frequency (counts) of different conformations over the relative change in position is a useful way to characterize various dominant and minor states (Figure 7).</li>
		<li>Fit the histogram using (multiple) Gaussian functions to estimate the overall percentage of individual conformers at a given force (Figure 7C). The Gaussian fits, mean position, and the standard deviation outlines the force-related relationship among different populations. 
		NOTE: A custom-written python script allowing pre-processing and basic bimodal Gaussian fitting of constant-force data is provided in the supplementary data. Parameters (cut-off frequency, filter degree, expected means, standard deviation values and amplitudes) need to be optimized for different experiments.</li>
		<li>Next, employ the Hidden Markov model to further analyze the states, which may uncover additional folding intermediates (conformers)55. For further information on the constant-force and Hidden Markov model, one may refer to55,56,57,58.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Here, we demonstrate the use of fluorescence-coupled optical tweezers to study interactions and dynamic behavior of RNA molecules with various ligands. Below, critical steps and limitations of the present technique are discussed.
Critical steps in the protocol 
As for many other methods, the quality of the sample is pivotal to obtain reliable data. Therefore, to obtain the highest possible quality samples, it is worth it to spend time to optimize the procedure for sample preparation. The optimization steps include proper primer design, annealing temperatures, RNA and protein purification steps.
Throughout the experiment use of filtered tips and solutions is crucial in order to maintain RNase-free conditions. In addition, the microfluidics system is kept in bleach when not in use. Before starting measurements, it is important to wash the system properly with sodium thiosulfate and RNase-free water to remove the bleach from the system.
In case the same-sized beads are used throughout the experiment, it is not required to perform force calibration each time. Nevertheless, force calibration checks should be done regularly for the reproducibility of experiments.
Modifications and troubleshooting of the method 
Fluorophore stability and photobleaching 
A complication during fluorescence measurements is photobleaching. Since the time frame to monitor translation can be extended from seconds to minutes depending on the system, photobleaching during the measurements should be also considered and minimized as much as possible73. One option is to employ more stable fluorophores, which are less prone to photobleaching, such as recently introduced quantum dots49,74,75. Further stability is also achieved by removing oxygen molecules using an &#34;oxygen scavenger&#34; system, such as glucose oxidase coupled with catalase. Glucose oxidase removes oxygen from the environment by turning it into hydrogen peroxide, which is then decomposed by catalase. Alternative oxygen scavenging systems can also be employed76,77.
Microfluidics 
Maintaining a continuous laminar flow is essential for proper measurements. Most importantly, the system should never run dry. Unfortunately, RBPs or other trans-acting factors of interest are often available only in small volumes for the experiments, therefore maintaining continuous flow can be challenging and cost intensive. If air bubbles are introduced into the system during the sample application, manual pressure or ethanol wash is usually sufficient for their removal.
Limitations of the method 
Combination of OT with confocal microscopy also brings some limitations. First, the focal plane of the confocal unit must be aligned properly with trap centers to allow proper recording of fluorescence signal. Furthermore, for confocal measurements, handles of at least 2 kb at each site are usually needed17. Although in principle using longer handles is possible, one should consider the energy contribution of the handles and the change in the persistence length for the accuracy of data analysis78. Another crucial point is the oxygen scavengers, which are used to increase the half-life of the fluorophores, also lead to relatively quick changes in pH of the solutions76. These changes can be partially compensated by increasing the concentration of the buffering compound; however, during the measurements, samples should be replenished regularly (every 30-60 min) to ensure consistent conditions through the experiment.

Transfer of genetic information from DNA to proteins through mRNAs is a complex biochemical process, which is precisely regulated on all levels through macromolecular interactions inside cells. For translational regulation, RNA-protein interactions confer a critical role to rapidly react to various stimuli and signals1,2. Some RNA-protein interactions affect mRNA stability and thereby alter the time an RNA is translationally active. Other RNA-protein interactions are associated with recoding mechanisms such as stop-codon readthrough, bypassing, or programmed ribosomal frameshifting (PRF)3,4,5,6,7. Recently, a number of RNA-binding proteins (RBPs) have been demonstrated to interact with stimulatory mRNA elements and the translation machinery to dictate when and how much recoding will occur in the cell7,8,9,10,11. Thus, to dissect the role of RNA-binding proteins in translation and how they modulate RNA structure and stability, it is pivotal to study the interaction principles and mechanical properties of these RNA-protein complexes in detail.
Decades of work have laid the foundation to study the multi-step and multi-component process of translation, which relies on intricate communication between the RNA and protein components of the translation machinery to achieve speed and accuracy12,13,14. A crucial next step in understanding complex regulatory events is determining the forces, timescales, and structural determinants during translation at high precision12,15,16,17. The study of RNA conformational dynamics and especially how trans-acting auxiliary factors act on the RNA structure during translation have been further illuminated by the emergence of single-molecule tools, including optical tweezers or zero-mode waveguides16,17,18,19,20,21,22,23,24,25,26.
Optical tweezers (OT) represent a highly precise single-molecule technique, which has been applied to study many sorts of RNA-dependent dynamic processes including transcription, and translation26,27,28,29,30,31,32. The use of optical tweezers has allowed probing of molecular interactions, nucleic acid structures, and thermodynamic properties, kinetics, and energetics of these processes in detail16,17,22,33,34,35,36,37,38,39. Optical tweezers assay is based on the entrapment of microscopic objects with a focused laser beam. In a typical OT experiment, the molecule of interest is tethered between two transparent (usually polystyrene) beads (Figure 1A)27. These beads are then caught by optical traps, which behave like springs. Thus, the force applied on the molecule can be calculated based on the bead&#39;s displacement from the center of the focused laser beam (trap center). Recently, optical tweezers have been combined with confocal microscopy (Figure 1B), enabling fluorescence or F&#246;rster resonance energy transfer (FRET) measurements40,41,42. This opens a whole new field of possible experiments allowing simultaneous measurement and, therefore, precise correlation of force spectroscopy and fluorescence data.
Here, we demonstrate experiments using the optical tweezers combined with confocal microscopy to study protein-RNA interactions regulating translational frameshifting. Between the objective and the condenser, a flow cell with five channels enables continuous sample application with laminar flow. Through the microfluidic channels, various components can be injected directly, which decreases the hands-on time as well as allowing very little sample consumption throughout the experiment.
First, a basic guideline to assist the design of OT experiments is proposed and advantages as well as pitfalls of various setups are discussed. Next, the preparation of samples and experimental workflows are described, and a protocol for the data analysis is provided. To represent an example, we outline the results obtained from RNA stretching experiments to study the SARS-CoV-2 frameshifting RNA element (Figure 2A) with the trans-acting factor the short isoform of zinc-finger antiviral protein (ZAP), which alters the translation of the viral RNA from an alternative reading frame43. Additionally, it is demonstrated that fluorescence-labeled ribosomes can be employed in this OT confocal assay, which would be useful to monitor the processivity and speed of the translation machinery. The method presented here can be used to rapidly test the effect of different buffers, ligands, or other cellular components to study various aspects of translation. Finally, common experimental pitfalls and how to troubleshoot them are discussed. Below, some crucial points in experimental design are outlined.
Construct design 
In principle, there are two common approaches to create an OT-compatible RNA construct. The first approach employs a long RNA molecule that is hybridized with complementary DNA handles, thus yielding a construct consisting of two RNA/DNA hybrid regions flanking a single-stranded RNA sequence in the middle (Figure 2B). This approach is employed in most OT RNA experiments33,44,45.
The second approach takes advantage of dsDNA handles with short (around 20 nt) overhangs15,17. These overhangs are then hybridized with the RNA molecule. Although more complicated in design, the use of dsDNA handles overcomes some of limitations of the DNA/RNA-hybrid system. In principle, even very long handles (&#62;10kb) can be implemented, which is more convenient for confocal measurements. In addition, the RNA molecule can be ligated to DNA handles to increase tether stability.
End-labeling strategy 
The construct must be tethered to beads via a strong molecular interaction. While there are approaches available for covalent bonding of handles to beads46, strong but non-covalent interactions such as streptavidin-biotin and digoxigenin-antibody are commonly used in OT experiments15,33,35,45. In the described protocol, the construct is labeled with biotin or digoxigenin, and the beads are coated with streptavidin or antibodies against digoxigenin, respectively (Figure 1A). This approach would be suitable for applying forces up to approximately 60 pN (per tether)47. Furthermore, the use of different 5&#39; and 3&#39; labeling strategies allow determining the orientation of the tether formed between the beads17.
Protein labeling for fluorescence measurements 
For the confocal imaging, there are several commonly used approaches for fluorescence labeling. For instance, fluorophores can be covalently attached to amino acid residues that are found natively in proteins or introduced by site-directed mutagenesis through a reactive organic group. Thiol or amine-reactive dyes can be used for labeling of cysteine and lysine residues, respectively. There are several reversible protection methods to increase the specificity of labeling48,49, however native proteins would typically be labeled at multiple residues. Although the small size of the fluorophore may confer an advantage, non-specific labeling might interfere with the protein activity and thus signal intensity may vary49. Also, depending on the labeling efficiency signal intensity may differ between different experiments. Therefore, an activity check should be performed prior to the experiment.
In case the protein of interest contains an N- or C-terminal tag, such as a His-tag or strep-tag, specific labeling of these tags represents another popular approach. Moreover, tag-targeted labeling reduces the chance of the fluorophore interfering with protein activity and can enhance solubility49. However, tag-specific labeling usually yields mono-fluorophore labeled proteins, which might be challenging to detect. Another way of specific labeling can be accomplished by employing antibodies.
Microfluidics setup 
The combination of OT with a microfluidics system allows a rapid transition between different experimental conditions. Moreover, current systems take advantage of maintaining the laminar flow inside the flow cell, which precludes the mixing of liquids from other channels in the perpendicular direction relative to the flow direction. Therefore, laminar flow is particularly advantageous for the experimental design. Currently, flow cells with up to 5 channels are commonly employed (Figure 3).

<table><tbody><tr><td>&lt;strong&gt;Bacterial Strains&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>E. coli HB101</td><td>lab collection</td><td>N/A</td><td>cloning of the vectors</td></tr><tr><td>&lt;strong&gt;Chemicals and enzymes&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich</td><td>31424</td><td>Buffers</td></tr><tr><td>Biotin-16-dUTP</td><td>Roche</td><td>11093070910</td><td>Biotinylation</td></tr><tr><td>BSA</td><td>Sigma-Aldrich</td><td>A4737</td><td>Buffers</td></tr><tr><td>Catalase</td><td>Lumicks</td><td>N/A</td><td>Oxygen scavanger system</td></tr><tr><td>Dithiothreitol (DTT)</td><td>Melford Labs</td><td>D11000</td><td>Buffers</td></tr><tr><td>DNAse I from bovine pancreas</td><td>Sigma-Aldrich</td><td>D4527</td><td>in vitro transcription</td></tr><tr><td>dNTPs</td><td>Th.Geyer</td><td>11786181</td><td>PCR</td></tr><tr><td>EDTA</td><td>Sigma-Aldrich</td><td>E9884</td><td>Buffers</td></tr><tr><td>Formamide</td><td>Sigma-Aldrich</td><td>11814320001</td><td>Buffers</td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G8270-1KG</td><td>Oxygen scavanger system</td></tr><tr><td>Glucose-oxidase</td><td>Lumicks</td><td>N/A</td><td>Oxygen scavanger system</td></tr><tr><td>HEPES</td><td>Carl Roth</td><td>HN78.3</td><td>Buffers</td></tr><tr><td>Magnesium chloride</td><td>Carl Roth</td><td>2189.1</td><td>Buffers</td></tr><tr><td>Phusion DNA polymerase</td><td>NEB</td><td>M0530L</td><td>Gibson assembly, cloning</td></tr><tr><td>Potassium chloride</td><td>Merck</td><td>529552-1KG</td><td>Buffers</td></tr><tr><td>PrimeSTAR GXL DNA Polymerase</td><td>Takara Bio Clontech</td><td>R050A</td><td>PCR</td></tr><tr><td>Pyrophosphotase, thermostabile, inorganic</td><td>NEB</td><td>M0296L</td><td>in vitro transcription</td></tr><tr><td>RNase Inhibitor</td><td>Molox</td><td>1000379515</td><td>Buffers</td></tr><tr><td>rNTPS</td><td>life technologies</td><td>R0481</td><td>in vitro transcription</td></tr><tr><td>Sodium thiosulophate</td><td>Sigma-Aldrich</td><td>S6672-500G</td><td>Bleach deactivation</td></tr><tr><td>Sytox Green</td><td>Lumicks</td><td>N/A</td><td>confocal measurements</td></tr><tr><td>T4 DNA Polymerase</td><td>NEB</td><td>M0203S</td><td>Biotinylation</td></tr><tr><td>T5 exonuclease</td><td>NEB</td><td>M0363S</td><td>Gibson assembly, cloning</td></tr><tr><td>T7 RNA polymerase</td><td>Produced in-house</td><td>N/A</td><td>in vitro transcription</td></tr><tr><td>Taq DNA polymerase</td><td>NEB</td><td>M0267S</td><td>PCR</td></tr><tr><td>Taq ligase</td><td>Biozym</td><td>L6060L</td><td>Gibson assembly, cloning</td></tr><tr><td>TWEEN 20 BioXtra</td><td>Sigma-Aldrich</td><td>P7949</td><td>Buffers</td></tr><tr><td>&lt;strong&gt;Kits&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Monolith Protein Labeling Kit RED-NHS 2nd Generation (Amine Reactive)</td><td>Nanotemper</td><td>MO-L011</td><td>Used for ribosome labeling</td></tr><tr><td>Purefrex 2.0</td><td>GeneFrontier</td><td>PF201-0.25-EX</td><td>Ribosomes used for the labeling</td></tr><tr><td>&lt;strong&gt;Oligonucleotides&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>5&#039; handle T7 forward</td><td>Microsynth</td><td>custom order</td><td>5&amp;rsquo; - CTTAATACGACTCACTATAGGTC&lt;br/&gt; CTTTCTGTGGACGCC - 3&amp;rsquo;, used to generate OT in vitro transcription template in PCR 1</td></tr><tr><td>3&amp;rsquo; handle reverse</td><td>Microsynth</td><td>custom order</td><td>5&#039; -&amp;nbsp; GTCAAAGTGCGCCCCGTTATCC - 3&#039;, used to generate OT in vitro transcription template in PCR 1</td></tr><tr><td>5&#039; handle forward</td><td>Microsynth</td><td>custom order</td><td>5&#039; - TCCTTTCTGTGGACGCCGC - 3&#039; , used to generate 5&#039; handle in PCR 2</td></tr><tr><td>5&amp;rsquo; handle reverse</td><td>Microsynth</td><td>custom order</td><td>5&amp;rsquo; - CATAAATACCTCTTTACTAATATA&lt;br/&gt; TATACCTTCGTAAGCTAGCGT - 3&amp;rsquo;, used to generate 5&#039; handle in PCR 2</td></tr><tr><td>3&amp;rsquo; handle forward</td><td>Microsynth</td><td>custom order</td><td>5&#039; - ATCCTGCAACCTGCTCTTCGCC&lt;br/&gt; AG - 3&#039;, used to generate 3&#039; handle in PCR 3</td></tr><tr><td>3&amp;rsquo; handle reverse 5&amp;rsquo;labeled with digoxigenin</td><td>Microsynth</td><td>custom order</td><td>5&#039; -[Dig]-GTCAAAGTGCGCCCCGTTATCC - 3&#039;, used to generate 3&#039; handle in PCR 3</td></tr><tr><td>&lt;strong&gt;DNA vectors&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>pMZ_OT</td><td>produced in-house</td><td>N/A</td><td>further description in &quot;Structural studies of Cardiovirus 2A protein reveal the molecular basis for RNA recognition and translational control&quot;&lt;br/&gt; Chris H. Hill, Sawsan Napthine, Lukas Pekarek, Anuja Kibe, Andrew E. Firth, Stephen C. Graham, Neva Caliskan, Ian Brierley&lt;br/&gt; bioRxiv 2020.08.11.245035; doi: https://doi-org.remotexs.ntu.edu.sg/10.1101/2020.08.11.245035</td></tr><tr><td>&lt;strong&gt;Software and Algorithms&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Atom</td><td>https://atom.io/packages/ide-python</td><td>N/A</td><td></td></tr><tr><td>Bluelake</td><td>Lumicks</td><td>N/A</td><td></td></tr><tr><td>Graphpad</td><td>https://www.graphpad.com/</td><td>N/A</td><td></td></tr><tr><td>InkScape 0.92.3</td><td>https://inkscape.org/</td><td>N/A</td><td></td></tr><tr><td>Matlab</td><td>https://www.mathworks.com/products/matlab.html</td><td>N/A</td><td></td></tr><tr><td>POTATO</td><td>https://github.com/lpekarek/POTATO.git</td><td>N/A</td><td></td></tr><tr><td>RNAstructure</td><td>https://rna.urmc.rochester.edu/RNAstructure.html</td><td>N/A</td><td></td></tr><tr><td>Spyder</td><td>https://www.spyder-ide.org/</td><td>N/A</td><td></td></tr><tr><td>&lt;strong&gt;Other&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Streptavidin Coated Polystyrene Particles, 1.5-1.9 &amp;micro;m, 5 ml, 1.0% w/v</td><td>Spherotech</td><td>SVP-15-5</td><td></td></tr><tr><td>Anti-digoxigenin Coated Polystyrene Particles, 2.0-2.4 &amp;micro;m, 2 ml, 0.1% w/v</td><td>Spherotech</td><td>DIGP-20-2</td><td></td></tr><tr><td>Syringes</td><td>VWR</td><td>TERUMO SS+03L1</td><td></td></tr><tr><td>&lt;strong&gt;Devices&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>C-trap</td><td>Lumicks</td><td>N/A</td><td>&amp;nbsp;optical tweezers coupled with confocal microscopy</td></tr></tbody></table>

optical tweezers to study rna-protein interactions in translation regulation

RNA adopts diverse structural folds, which are essential for its functions and thereby can impact diverse processes in the cell. In addition, the structure and function of an RNA can be modulated by various trans-acting factors, such as proteins, metabolites or other RNAs. Frameshifting RNA molecules, for instance, are regulatory RNAs located in coding regions, which direct translating ribosomes into an alternative open reading frame, and thereby act as gene switches. They may also adopt different folds after binding to proteins or other trans-factors. To dissect the role of RNA-binding proteins in translation and how they modulate RNA structure and stability, it is crucial to study the interplay and mechanical features of these RNA-protein complexes simultaneously.&#160;This work illustrates how to employ single-molecule-fluorescence-coupled optical tweezers to explore the conformational and thermodynamic landscape of RNA-protein complexes at a high resolution. As an example, the interaction of the SARS-CoV-2 programmed ribosomal frameshifting element with the trans-acting factor short isoform of zinc-finger antiviral protein is elaborated. In addition, fluorescence-labeled ribosomes were monitored using the confocal unit, which would ultimately enable the study of translation elongation. The fluorescence coupled OT assay can be widely applied to explore diverse RNA-protein complexes or trans-acting factors regulating translation and could facilitate studies of RNA-based gene regulation.

Watch this Scientific Journal Video about Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation at JoVE.com

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

This protocol presents a complete experimental workflow for studying RNA-protein interactions using optical tweezers. Several possible experimental setups are outlined including the combination of optical tweezers with confocal microscopy.

RNA adopts diverse structural folds, which are essential for its functions and thereby can impact diverse processes in the cell. In addition, the ...

optical-tweezers-to-study-rna-protein-interactions-translation

Nanyang Technological University

Research

JoVE Journal

Biochemistry

4.9K Views.  Helmholtz Zentrum für Infektionsforschung (Helmholtz Centre for Infection Research). Het algemene doel van het volgende methodeartikel is om een protocol aan te tonen dat nuttig is om RNA-interacties met eiwitten of andere factoren te bestuderen met behulp van optische pincetten in combinatie met confocale microscopie. In het afgelopen decennium zijn verschillende geavanceerde technieken ontwikkeld om de eigenschappen van afzonderlijke macromoleculen te onderzoeken. Een van deze technieken is een enkel molecuul optisch pincet.Met het nieuwe fluorescentieondersteunde op...

Video: Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

Quantitative Immunofluorescence to Measure Global Localized Translation

The mechanisms regulating mRNA translation are involved in various biological processes, such as germ line development, cell differentiation, and organogenesis, as well as in multiple diseases. Numerous publications have convincingly shown that specific mechanisms tightly regulate mRNA translation. Increased interest in the translation-induced regulation of protein expression has led to the development of novel methods to study and follow de novo protein synthesis in cellulo. However, most of these methods are complex, making them costly and often limiting the number of mRNA targets that can be studied. This manuscript proposes a method that requires only basic reagents and a confocal fluorescence imaging system to measure and visualize the changes in mRNA translation that occur in any cell line under various conditions. This method was recently used to show localized translation in the subcellular structures of adherent cells over a short period of time, thus offering the possibility of visualizing de novo translation for a short period during a variety of biological processes or of validating changes in translational activity in response to specific stimuli.

This manuscript describes a method to visualize and quantify localized translation events in subcellular compartments. The approach proposed in this manuscript requires a basic confocal imaging system and reagents and is rapid and cost-effective.

The mechanisms regulating mRNA translation are involved in various biological processes, such as germ line development, cell differentiation, and ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Toeprinting Analysis of Translation Initiation Complex Formation on Mammalian mRNAs

Translation initiation is the rate-limiting step of protein synthesis and represents a key point at which cells regulate their protein output. Regulation of protein synthesis is the key to cellular stress-response, and dysregulation is central to many disease states, such as cancer. For instance, although cellular stress leads to the inhibition of global translation by attenuating cap-dependent initiation, certain stress-response proteins are selectively translated in a cap-independent manner. Discreet RNA regulatory elements, such as cellular internal ribosome entry sites (IRESes), allow for the translation of these specific mRNAs. Identification of such mRNAs, and the characterization of their regulatory mechanisms, have been a key area in molecular biology. Toeprinting is a method for the study of RNA structure and function as it pertains to translation initiation. The goal of toeprinting is to assess the ability of in vitro transcribed RNA to form stable complexes with ribosomes under a variety of conditions, in order to determine which sequences, structural elements, or accessory factors are involved in ribosome binding&#8212;a pre-cursor for efficient translation initiation. Alongside other techniques, such as western analysis and polysome profiling, toeprinting allows for a robust characterization of mechanisms for the regulation of translation initiation.

Toeprinting aims to measure the ability of in vitro transcribed RNA to form translation initiation complexes with ribosomes under a variety of conditions. This protocol describes a method for toeprinting mammalian RNA and can be used to study both cap-dependent and IRES-driven translation.

Translation initiation is the rate-limiting step of protein synthesis and represents a key point at which cells regulate their protein output. Regulation ...

An Oligonucleotide-based Tandem RNA Isolation Procedure to Recover Eukaryotic mRNA-Protein Complexes

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein complexes is essential to understanding mRNA regulation inferred by interacting proteins or non-coding RNAs. Herein, we describe a tandem RNA isolation procedure (TRIP) that enables the purification of endogenously formed mRNA-protein complexes from cellular extracts. The two-step protocol involves the isolation of polyadenylated mRNAs with antisense oligo(dT) beads and subsequent capture of an mRNA of interest with 3&#39;-biotinylated 2&#39;-O-methylated antisense RNA oligonucleotides, which can then be isolated with streptavidin beads. TRIP was used to recover in vivo crosslinked mRNA-ribonucleoprotein (mRNP) complexes from yeast, nematodes and human cells for further RNA and protein analysis. Thus, TRIP is a versatile approach that can be adapted to all types of polyadenylated RNAs across organisms to study the dynamic re-arrangement of mRNPs imposed by intracellular or environmental cues.

A tandem RNA isolation procedure (TRIP) for recovery of endogenously formed mRNA-protein complexes is described. Specifically, RNA-protein complexes are crosslinked in vivo, polyadenylated RNAs are isolated from extracts with oligo(dT) beads, and particular mRNAs are captured with modified RNA antisense oligonucleotides. Proteins bound to mRNAs are detected by immunoblot analysis.

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein ...

An Assay for Quantifying Protein-RNA Binding in Bacteria

In the initiation step of protein translation, the ribosome binds to the initiation region of the mRNA. Translation initiation can be blocked by binding of an RNA binding protein (RBP) to the initiation region of the mRNA, which interferes with ribosome binding. In the presented method, we utilize this blocking phenomenon to quantify the binding affinity of RBPs to their cognate and non-cognate binding sites. To do this, we insert a test binding site in the initiation region of a reporter mRNA and induce the expression of the test RBP. In the case of RBP-RNA binding, we observed a sigmoidal repression of the reporter expression as a function of RBP concentration. In the case of no-affinity or very low affinity between binding site and RBP, no significant repression was observed. The method is carried out in live bacterial cells, and does not require expensive or sophisticated machinery. It is useful for quantifying and comparing between the binding affinities of different RBPs that are functional in bacteria to a set of designed binding sites. This method may be inappropriate for binding sites with high structural complexity. This is due to the possibility of repression of ribosomal initiation by complex mRNA structure in the absence of RBP, which would result in lower basal reporter gene expression, and thus less-observable reporter repression upon RBP binding.

In this method, we quantify the binding affinity of RNA binding proteins (RBPs) to cognate and non-cognate binding sites using a simple, live, reporter assay in bacterial cells. The assay is based on repression of a reporter gene.

In the initiation step of protein translation, the ribosome binds to the initiation region of the mRNA. Translation initiation can be blocked by binding ...

Genetics

Nanomanipulation of Single RNA Molecules by Optical Tweezers

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be increasingly important. As RNA function often depends on its ability to adopt alternative structures, it is difficult to predict RNA three-dimensional structures directly from sequence. Single-molecule approaches show potentials to solve the problem of RNA structural polymorphism by monitoring molecular structures one molecule at a time. This work presents a method to precisely manipulate the folding and structure of single RNA molecules using optical tweezers. First, methods to synthesize molecules suitable for single-molecule mechanical work are described. Next, various calibration procedures to ensure the proper operations of the optical tweezers are discussed. Next, various experiments are explained. To demonstrate the utility of the technique, results of mechanically unfolding RNA hairpins and a single RNA kissing complex are used as evidence. In these examples, the nanomanipulation technique was used to study folding of each structural domain, including secondary and tertiary, independently. Lastly, the limitations and future applications of the method are discussed.

Optical tweezers have been used to study RNA folding by stretching individual molecules from their 5&#8217; and 3&#8217; ends. Here common procedures are described to synthesize RNA molecules for tweezing, calibration of the instrument, and methods to manipulate single molecules.

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be ...

Bioengineering

Dual DNA Rulers to Study the Mechanism of Ribosome Translocation with Single-Nucleotide Resolution

The ribosome translocation refers to the ribosomal movement on the mRNA by exactly three nucleotides (nt), which is the central step in protein synthesis. To investigate its mechanism, there are two essential technical requirements. First is single-nt resolution that can resolve normal translocation from frameshifting, during which the ribosome moves by other than 3 nt. The second is the capability to probe both the entrance and exit sides of mRNA in order to elucidate the whole picture of translocation. We report the dual DNA ruler assay that is based on the critical dissociation forces of DNA-mRNA duplexes, obtained by force-induced remnant magnetization spectroscopy (FIRMS).&#160; With 2-4 pN force resolution, the dual ruler assay is sufficient to distinguish different translocation steps. By implementing a long linker on the probing DNAs, they can reach the mRNA on the opposite side of the ribosome, so that the mRNA position can be determined for both sides. Therefore, the dual ruler assay is uniquely suited to investigate the ribosome translocation, and nucleic acid motion in general. We show representative results which indicated a looped mRNA conformation and resolved normal translocation from frameshifting.

Dual DNA ruler assay is developed to determine the mRNA position during ribosome translocation, which relies on the dissociation forces of the formed DNA-mRNA duplexes. With single-nucleotide resolution and capability of reaching both ends of mRNA, it can provide mechanistic insights for ribosome translocation and probe other nucleic acid displacements.

The ribosome translocation refers to the ribosomal movement on the mRNA by exactly three nucleotides (nt), which is the central step in protein synthesis. ...

Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately 20 nm to accommodate large tRNA substrates at the A-, P- and E-sites. Consequently, the ribosome dynamics are naturally de-phased quickly. Single molecule method can detect each ribosome separately and distinguish inhomogeneous populations, which is essential to reveal the complicated mechanisms of multi-component systems. We report the details of a smFRET method based on the Nikon Ti2 inverted microscope to probe the ribosome dynamics between the ribosomal protein L27 and tRNAs. The L27 is labeled at its unique Cys 53 position and reconstituted into a ribosome that is engineered to lack L27. The tRNA is labeled at its elbow region. As the tRNA moves to different locations inside the ribosome during the elongation cycle, such as pre- and post- translocation, the FRET efficiencies and dynamics exhibit differences, which have suggested multiple subpopulations. These subpopulations are not detectable by ensemble methods. The TIRF-based smFRET microscope is built on a manual or motorized inverted microscope, with home-built laser illumination. The ribosome samples are purified by ultracentrifugation, loaded into a home-built multi-channel sample cell and then illuminated via an evanescent laser field. The reflection laser spot can be used to achieve feedback control of perfect focus. The fluorescence signals are separated by a motorized filter-turret and collected by two digital CMOS cameras. The intensities are retrieved via the NIS-Elements software.

Single molecule fluorescence energy transfer is a method that tracks the tRNA dynamics during ribosomal protein synthesis. By tracking individual ribosomes, inhomogeneous populations are identified, which shed light on mechanisms. This method can be used to track biological conformational changes in general to reveal dynamic-function relationships in many other complexed biosystems. Single molecule methods can observe non-rate limiting steps and low-populated key intermediates, which are not accessible by conventional ensemble methods due to the average effect.

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately ...

An Optimized Quantitative Pull-Down Analysis of RNA-Binding Proteins Using Short Biotinylated RNA

Protein-RNA interactions regulate gene expression and cellular functions at transcriptional and post-transcriptional levels. For this reason, identifying the binding partners of an RNA of interest remains of high importance to unveil the mechanisms behind many cellular processes. However, RNA molecules might interact transiently and dynamically with some RNA-binding proteins (RBPs), especially with non-canonical ones. Hence, improved methods to isolate and identify such RBPs are greatly needed.
To identify the protein partners of a known RNA sequence efficiently and quantitatively, we developed a method based on the pull-down and characterization of all interacting proteins, starting from cellular total protein extract. We optimized the protein pull-down using biotinylated RNA pre-loaded on streptavidin-coated beads. As a proof of concept, we employed a short RNA sequence known to bind the neurodegeneration-associated protein TDP-43 and a negative control of a different nucleotide composition but the same length. After blocking the beads with yeast tRNA, we loaded the biotinylated RNA sequences on the streptavidin beads and incubated them with the total protein extract from HEK 293T cells. After incubation and several washing steps to remove nonspecific binders, we eluted the interacting proteins with a high-salt solution, compatible with the most commonly used protein quantification reagents and with sample preparation for mass spectrometry. We quantified the enrichment of TDP-43 in the pull-down performed with the known RNA binder compared to the negative control by mass spectrometry. We used the same technique to verify the selective interactions of other proteins computationally predicted to be unique binders of our RNA of interest or of the control. Finally, we validated the protocol by western blot via the detection of TDP-43 with an appropriate antibody. 
This protocol will allow the study of the protein partners of an RNA of interest in near-to-physiological conditions, helping uncover unique and unpredicted protein-RNA interactions.

Here, we present an optimized in vitro method to uncover, quantify, and validate protein interactors of specific RNA sequences, using total protein extract from human cells, streptavidin beads coated with biotinylated RNA, and mass spectrometry analysis.

Protein-RNA interactions regulate gene expression and cellular functions at transcriptional and post-transcriptional levels. For this reason, identifying ...

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 ...

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

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