Name: single molecule fluorescence energy transfer study of ribosome protein synthesis
Uploaded: 2021-07-06T15:00:00.000+00:00
Duration: 8 min 7 s
Description: Watch this Scientific Journal Video about Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis at JoVE.com

This work is supported by the US National Institutes of Health (R01GM111452) and the Welch Foundation (E-1721).

1. Preparation of labeled ribosome and tRNA for FRET detection
<ol>
	<li>Isolate ribosome without L27 from E. coli strain IW312 according to standard protocols20,21. Extract the regular ribosome from E. coli strain MRE600.</li>
	<li>Clone the L27&#39;s rpmA gene with C-terminal His-tag into pET-21b (+) plasmid, which is transformed and expressed in BL21(DE3)pLysS cells15. Purify the protein via a prepacked sepharose column.</li>
	<li>Labeling of L27
	<ol>
		<li>Incubate 20-100 &#956;M of purified L27 in 100 &#956;L with 2-10-fold excess of TCEP (Tris-2-carboxyethyl-phosphine) at room temperature (RT) for 30 min. Buffer exchange the solution into PBS buffer (10 mM Na2HPO4, 1.8 mM KH2PO4, 2.7 mM KCl; 0.137 M NaCl), using a nap-5 column. Add 10-fold excess Cy5-maleimide mono-reactive dye in DMSO into the solution and incubate at RT in the dark for 2 h.</li>
		<li>Load the labeled protein solution (500 &#956;L) mentioned above on a Nap-5 column to remove the excess dye. Ensure that the protein elutes in the first colored band, followed by the free dye elute in the second band, respectively. Collect the eluted protein in 1 mL of TAM10 buffer (20 mM tris-HCl (pH 7.5), 10 mM Mg (OAc)2, 30 mM NH4Cl, 70 mM KCl, 0.5 mM EDTA, and 7 mM BME (2-mercaptoethanol)).</li>
	</ol>
	</li>
	<li>Reconstitute the ribosome with L27. Mix the labeled L27 protein with an equal amount of IW312 ribosome in TAM10 buffer and incubate at 37 &#176;C for 1 h. Remove the free protein by sucrose cushion ultracentrifugation (100,000 x g, overnight) to allow the ribosome to form a blue-colored pellet at the bottom of the tube.</li>
	<li>Label the tRNAPhe according to the previously published report22. First, incubate the tRNA (10 A260 in 1 mL of water) with 100 &#956;L of NaBH4 (100 mM stock, adding dropwise) at 0 &#176;C for 1 h. Free the tRNAs from unreacted NaBH4 via three-time ethanol precipitation by adding 1/10 volume of 3 M KAc (pH 5.0) and 2.5x volume of ethanol. Incubate the reduced tRNA with 1 &#956;L of Cy3-NHS dye solution (1 mg in 10 &#956;L of DMSO) in a minimum volume (~20 &#956;L) of 0.1 M sodium formate (pH 3.7). After 2 h of incubation at 37 &#176;C, separate the tRNA from the unreacted dye via a small G25 Sephadex column, and then remove the free dye via two-time ethanol precipitation.</li>
</ol>
2. Preparation of the ribosome complexes
<ol>
	<li>Express and purify the His-tagged proteins of IF1, IF2, IF3, EF-G, EF-Tu, EF-Ts, and tRNA synthetases using standard methods15.</li>
	<li>Prepare the initiation mix by adding the following ingredients in TAM10 buffer at 37 &#176;C for 15 min: 1 &#181;M of the labeled ribosome; 1.5 &#181;M each of IF1, IF2, and IF3; 2 &#181;M biotinylated mRNA (coding MF for the first two amino acids); 4 &#181;M of charged fMet-tRNAfMet; and 1 mM GTP.</li>
	<li>Prepare the EF-Tu_G mix by mixing the following ingredients in TAM10 buffer at 37 &#176;C for 15 min: 4 &#181;M EF-Tu, 0.4 &#181;M EF-Ts, 2 &#181;M EF-G, 4 mM GTP, 4 mM PEP, and 0.02 mg/mL of Pyruvate Kinase.</li>
	<li>Prepare the EF-Tu_NoG mix similar to step 2.3 without EF-G.</li>
	<li>Prepare the Phe mix by mixing the following ingredients in nuclease-free water at 37 &#176;C for 15 min: 100 mM Tris (pH 7.5), 20 mM MgAc2, 1 mM EDTA, 4 mM ATP, 7 mM BME, 2 mM Phenylalanine-specific synthetase, 2 A260/mL of labeled tRNAPhe, and 50 &#956;M phenylalanine.</li>
	<li>Prepare the pre-translocation complex (PRE) by mixing the initiation, EF-Tu_NoG, and Phe mixes in the ratio of 1:2:2, at 37 &#176;C for 2 min. After incubation, purify the PRE by 1.1 M sucrose cushion ultracentrifugation overnight at 100,000 x g.</li>
	<li>Prepare the post-translocation complex (POST) by mixing the initiation, EF-Tu_WG, and Phe mixes in the ratio of 1:2:2, at 37 &#176;C for 10 min. After incubation, purify the POST by 1.1 M sucrose cushion ultracentrifugation overnight at 100,000 x g.</li>
</ol>
3. Preparation of sample slides
<ol>
	<li>Clean the microscope glass slides (75 mm x 25 mm) containing six pairs of holes (1 mm in diameter). Each pair forms an inlet-outlet of the sample chamber. Ensure that the distance between each inlet-outlet hole is 13 mm and the center-to-center distance between two channels is 6 mm. Place six slides in a glass crucible.</li>
	<li>Fill the crucible with acetone and sonicate them for 5 min. Decant the acetone and rinse the slides three times with ultrapure water. Fill the crucible again with water and 1 mL of 10 M KOH. Sonicate for 20 min.</li>
	<li>Rinse the slides three times with ultrapure water. Fill the crucible again with ethanol and sonicate for 5 min. Completely decant the solvent. Let the slides dry in the fume hood.</li>
	<li>Clean the microscope glass coverslips (#1.5 thickness, 24 x 40 mm). Perform the cleaning steps as described in steps 3.1-3.3.</li>
	<li>Bake the cleaned slides and coverslips at 300 &#176;C for 3 h. Keep them in the furnace overnight to cool.</li>
	<li>Coat the coverslip with aminosilane. In the crucible containing coverslips, pour in methanol mix (100 mL of methanol, 5 mL of water, 0.5 mL of HAc, and 1 mL of aminosilane). Warm the crucible in a water bath for 10 min, sonicate it for 10 min, and then warm the crucible in the water bath for another 10 min. Decant the methanol mixture, rinse the coverslip well in three crucibles of clean water. Purge the surfaces with a dry nitrogen stream through a needle.</li>
	<li>Coat the coverslips with biotin-PEG in a laminar clean hood.
	<ol>
		<li>To make the PEG solution, use 98 mg of PEG and 2-3 mg of biotin-PEG in 200 &#956;L of 100 mM NaHCO3 solution.</li>
		<li>Lay one coverslip flat on a surface. Carefully drop 60 &#956;L of PEG solution on the top edge. Then, lay another coverslip on top of it, letting the capillary effect spread the solution between the two coverslips. Ensure no bubbles are formed.</li>
		<li>Repeat step 3.7.2 for two more pairs of coverslips. Coat six pieces of coverslips with biotin. If more coverslips are needed, adjust the amount of PEG and Biotin PEG accordingly to make more PEG solution.</li>
		<li>Store these coverslips in an empty tip-box filled with water and incubate for 3 h in the dark.</li>
		<li>After 3 h, separate the coverslips, rinse with water in three consecutive crucibles and purge with a dry nitrogen stream. Place the coated coverslips on a ceramic rack. Mark the surface with coating.</li>
	</ol>
	</li>
	<li>Assemble the sample chambers19.
	<ol>
		<li>Pull sharp pipette tips through the holes on the glass slides until they fit tight. Scrape the sharp ends off until it is completely flat to the glass surface. Cut the other end of the tip to be approximately 5 mm for sample loading.</li>
		<li>Cut a double-faced tape with the channel pattern on it (25 x 40 mm).</li>
		<li>Stick the double-faced tape onto the glass slides on the flat side.</li>
		<li>Stick the coated coverslip onto the double-faced tape. Press on it to make a tight seal of the sample chamber. Make sure the coated side is facing inward.</li>
	</ol>
	</li>
</ol>
4. Single molecule FRET imaging
<ol>
	<li>Turn on the computer.</li>
	<li>Turn on the microscope switch.</li>
	<li>Start the laser control interface and turn on the laser. Push the enable button on the laser control box to warm up the laser.</li>
	<li>Turn on the cameras.</li>
	<li>Start the microscope-associated software program. Click the upper shutter Off and click the lamp On.</li>
	<li>Prepare the TAM10_WT buffer by adding 10 &#956;L of 5% Tween-20 into 1 mL of TAM10 buffer.</li>
	<li>Prepare the deoxy solution by weighing 3 mg of glucose oxidase in a small microcentrifuge tube. Add 45 &#956;L of catalase. Vortex gently to dissolve the solid. Spin at 20,000 x g in a microcentrifuge for 1 min. Take the supernatant.</li>
	<li>Prepare 30% glucose solution in water and 200 mM Trolox solution in DMSO.</li>
	<li>Prepare 0.5 mg/mL of streptavidin solution in nuclease-free water.</li>
	<li>Add one drop of imaging oil to the TIRF objective.</li>
	<li>Put the sample chambers on the objective.</li>
	<li>Fill the chamber with 10 &#956;L of streptavidin solution. Wait for 1 min.</li>
	<li>Wash the chamber with 30 mL of TAM10_WT buffer and collect the run-through solution with a folded filter paper. Wait for 1 min.</li>
	<li>Dilute the ribosome complexes (PRE or POST) to 10-50 nM concentration with TAM10_WT buffer.</li>
	<li>Load 20 &#956;L of the ribosome sample into the channel. Wait for 2 min.</li>
	<li>Make the imaging buffer (50 &#956;L of TAM10 buffer plus 0.5 &#956;L each of deoxy, glucose, and Trolox solutions). Mix the buffer well.</li>
	<li>Flush the chamber with 30 &#956;L of the imaging buffer.</li>
	<li>Set the camera to acquire 100 ms/frame, 16 bits, 4 x 4 binning.</li>
	<li>Click the upper shutter On and the lamp Off.</li>
	<li>Start camera acquisition. Spread the images into three windows representing two individual cameras and the overlay.</li>
	<li>Click on Auto-focus. Fine-tune to find the best focus.</li>
	<li>Click on a method. Set the field points, file storage, and the loop number.</li>
	<li>Click on Run. 
	NOTE: A new window appears with real-time imaging. The progress of the time series and field of view series are shown at the top of the window.</li>
	<li>Once the image acquisition is complete, close the popup window.</li>
	<li>Open the ND file (acquired file).</li>
	<li>Click on ROI/ simple ROI editor. Choose the option Circle.</li>
	<li>Select the ROI on the image. Click on Finish when it is complete.</li>
	<li>Click on Measurement/ Time Measurement to show the data either as a plot or as a spreadsheet.</li>
	<li>Open the Export tab. Set the export parameters.</li>
	<li>Click on Export to save the intensities.</li>
</ol>

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Y. Wang declares no conflicts of interest.

SmFRET is sensitive to background signals. First, it is necessary to coat the sample chamber with 0.05% tween and then be added concurrently with the ribosome solution to block non-specific binding of the ribosome to the surface. To see fluorescence from the acceptor Cy5 emission, the oxygen scavenger cocktail (deoxy, glucose, and Trolox solutions) is essential. Without this solution, the bleaching is too fast in the acceptor channel to obtain useful information. Another critical step for ribosome experiments, specifical...

The ribosome is a &#248; 20 nm large ribonucleoprotein complex of a large (50S) and a small (30S) subunit. It assembles long peptides along the mRNA template processively and cooperatively. The ribosome 30S binds to the fMet-tRNAfMet and mRNA to start protein synthesis, and the 50S then joins to form the 70S initiation complex. The tRNAs bring amino acids to the ribosome at the A-site (aminoacyl- tRNA binding site), while the elongated peptidyl chain is held at the P-site (peptidyl- tRNA binding site). In the pre-translocation complex, the peptidyl chain is transferred to the tRNA at the A-site with one amino acid added. Meanwhile, the P-site tRNA is deacylated. Then, the A-, P- tRNAs move to the P-, E- sites to form the post-translocation complex, in which the E-site represents the tRNA exit site. In this state, the peptidyl-tRNA moves back to the P-site. The elongation cycle continues between the pre- and post-conformations while the ribosome translocates on the mRNA, one codon at a time1. The ribosome is highly coordinative of different functional sites to make this process efficient and accurate, such as inter-subunit ratcheting2, tRNA hybridization fluctuations3, GTPase activations4, L1 stalk opening-closing5, etc. Consequently, ribosomes quickly de-phase because every molecule moves at its own pace. The conventional methods can only deduce apparent average parameters, but low-populated or short-lived species will be masked in the average effect6. Single molecule method can break this limitation by detecting each ribosome individually, then identify different species via statistical reconstruction7. Different labeling sites have been implemented to probe ribosome dynamics, such as the interactions between tRNA-tRNA8, EF-G-L119, L1-tRNA10, etc. In addition, by labeling the large and small subunits, respectively, inter-subunit ratcheting kinetics and coordination with factors are observed11,12. Meanwhile, the smFRET method has broad applications in other central biological processes, and multi-color FRET methods are emerging13.
Previously a novel ribosome FRET pair was developed14,15. The recombinant ribosomal protein L27 has been expressed, purified, and labeled, and incorporated back into the ribosome. This protein interacted with the tRNAs at a close distance and helped stabilize the P-site tRNA in the post-translocation complex. When tRNA moved from the A- to the P-site, the distance between this protein and the tRNA is shortened, which can be distinguished by the smFRET signal. Multiple ribosome subpopulations have been identified using statistical methods and mutagenesis, and spontaneous exchange of these populations in the pre- but not post- translocation complex suggests the ribosome is more flexible before moving on the mRNA, and more rigid during decoding16,17,18. These variations are essential to the ribosome function. Here, the protocol describes the details of ribosome/tRNA-labeling, their incorporation in the ribosome, smFRET sample preparation, and data acquisition/analysis19.

<table><tbody><tr><td>Aminosilane</td><td>Laysanbio</td><td>MPEG-SIL-5000</td><td></td></tr><tr><td>Biotin-PEG</td><td>Laysanbio</td><td>Biotin-PEG-SVA-5000</td><td></td></tr><tr><td>BL21(DE3)pLysS cells</td><td>Novagen</td><td>71403</td><td></td></tr><tr><td>Catalase</td><td>millipore sigma</td><td>C3515</td><td></td></tr><tr><td>CS150FNX Micro Ultracentrifuge</td><td>nuaire</td><td></td><td></td></tr><tr><td>Cy3/C5-maleimide</td><td>ApexBio</td><td>A8138/A8140</td><td></td></tr><tr><td>ECLIPSE Ti2 inverted microscope</td><td>Nikon</td><td></td><td></td></tr><tr><td>EdgeGARD Laminar Flow Hood</td><td>Baker</td><td></td><td></td></tr><tr><td>Glucose oxidase</td><td>millipore sigma</td><td>G2133</td><td></td></tr><tr><td>Histrap HP column (Prepacked sepharose column)</td><td>Cytiva</td><td>17524701</td><td></td></tr><tr><td>Microscope cover slip</td><td>VWR</td><td>48393-230</td><td></td></tr><tr><td>Microscope glass slides</td><td>VWR</td><td>470235-792</td><td></td></tr><tr><td>ORCA-Flash4.0 V3 camera</td><td>Hamamatsu</td><td></td><td></td></tr><tr><td>PEG (5,000)</td><td>Laysanbio</td><td>MPEG-SVA-5000</td><td></td></tr><tr><td>pET-21b (+) plasmid</td><td>Novagen</td><td>69741</td><td></td></tr><tr><td>Sonicator</td><td>VWR</td><td>CPX-952-518R</td><td></td></tr><tr><td>TCEP</td><td>Apexbio</td><td>B6055</td><td></td></tr><tr><td>Trolox</td><td>millipore sigma</td><td>238813</td><td></td></tr></tbody></table>

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.

The smFRET had the ribosome labeled at the middle position of tRNA traffic, to distinguish the tRNA translocation from the A- to the P-site (Figure 1)15. The distance from the L27 labeling residue to the A- or P-site tRNA is 52 or 61 &#197;, respectively, corresponding to FRET efficiency of 0.47 and 0.65. After the image collection, fluorescence intensities from the donor and acceptor channels were retrieved and plotted as time lapses (Figure 1</s...

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Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

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 ...

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2.6K Views.  University of Houston. 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 intermed...

Video: Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

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 ...

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 ...

A Fluorescence-based Method to Study Bacterial Gene Regulation in Infected Tissues

Bacterial virulence genes are often regulated at the transcriptional level by multiple factors that respond to different environmental signals. Some factors act directly on virulence genes; others control pathogenesis by adjusting the expression of downstream regulators or the accumulation of signals that affect regulator activity. While regulation has been studied extensively during in vitro growth, relatively little is known about how gene expression is adjusted during infection. Such information is important when a particular gene product is a candidate for therapeutic intervention. Transcriptional approaches like quantitative, real-time RT-PCR and RNA-Seq are powerful ways to examine gene expression on a global level but suffer from many technical challenges including low abundance of bacterial RNA compared to host RNA, and sample degradation by RNases. Evaluating regulation using fluorescent reporters is relatively easy and can be multiplexed with fluorescent proteins with unique spectral properties. The method allows for single-cell, spatiotemporal analysis of gene expression in tissues that exhibit complex three-dimensional architecture and physiochemical gradients that affect bacterial regulatory networks. Such information is lost when data are averaged over the bulk population. Herein, we describe a method for quantifying gene expression in bacterial pathogens in situ. The method is based on simple tissue processing and direct observation of fluorescence from reporter proteins. We demonstrate the utility of this system by examining the expression of Staphylococcus aureus thermonuclease (nuc), whose gene product is required for immune evasion and full virulence ex vivo and in vivo. We show that nuc-gfp is strongly expressed in renal abscesses and reveal heterogeneous gene expression due in part to apparent spatial regulation of nuc promoter activity in abscesses fully engaged with the immune response. The method can be applied to any bacterium with a manipulatable genetic system and any infection model, providing valuable information for preclinical studies and drug development.

Described here is a method for analyzing bacterial gene expression in animal tissues at a cellular level. This method provides a resource for studying the phenotypic diversity occurring within a bacterial population in response to the tissue environment during an infection.

Bacterial virulence genes are often regulated at the transcriptional level by multiple factors that respond to different environmental signals. Some ...

Genetics

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

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. ...

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed extensive studies of cap-dependent translation and its regulation, high resolution kinetic characterization of this translation pathway is still lacking. Recently, we developed an in vitro assay to measure cap-dependent translation kinetics with single-molecule resolution. The assay is based on fluorescently labeled antibody binding to nascent epitope-tagged polypeptide. By imaging the binding and dissociation of antibodies to and from nascent peptide&#8211;ribosome&#8211;mRNA complexes, the translation progression on individual mRNAs can be tracked. Here, we present a protocol for establishing this assay, including mRNA and PEGylated slide preparations, real-time imaging of translation, and analysis of single molecule trajectories. This assay enables tracking of individual cap-dependent translation events and resolves key translation kinetics, such as initiation and elongation rates. The assay can be widely applied to distinct translation systems and should broadly benefit in vitro studies of cap-dependent translation kinetics and translational control mechanisms.

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Tracking individual translation events allows for high-resolution kinetic studies of cap-dependent translation mechanisms. Here we demonstrate an in vitro single-molecule assay based on imaging interactions between fluorescently labeled antibodies and epitope-tagged nascent peptides. This method enables single-molecule characterization of initiation and peptide elongation kinetics during active in vitro cap-dependent translation.

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed ...

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.

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 ...

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One &#945;-Synuclein Monomer at a Time

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural dynamics and its contributions to biology. While the FRET-based ruler reports on inter-dye distances in the 3-10 nm range, other spectroscopic techniques, such as protein-induced fluorescence enhancement (PIFE), report on the proximity between a dye and a protein surface in the shorter 0-3 nm range. Regardless of the method of choice, its use in measuring freely-diffusing biomolecules one at a time retrieves histograms of the experimental parameter yielding separate centrally-distributed sub-populations of biomolecules, where each sub-population represents either a single conformation that stayed unchanged within milliseconds, or multiple conformations that interconvert much faster than milliseconds, and hence an averaged-out sub-population. In single-molecule FRET, where the reported parameter in histograms is the inter-dye FRET efficiency, an intrinsically disordered protein, such as the &#945;-Synuclein monomer in buffer, was previously reported as exhibiting a single averaged-out sub-population of multiple conformations interconverting rapidly. While these past findings depend on the 3-10 nm range of the FRET-based ruler, we sought to put this protein to the test using single-molecule PIFE, where we track the fluorescence lifetime of site-specific sCy3-labeled &#945;-Synuclein proteins one at a time. Interestingly, using this shorter range spectroscopic proximity sensor, sCy3-labeled &#945;-Synuclein exhibits several lifetime sub-populations with distinctly different mean lifetimes that interconvert in 10-100 ms. These results show that while &#945;-Synuclein might be disordered globally, it nonetheless attains stable local structures. In summary, in this work we highlight the advantage of using different spectroscopic proximity sensors that track local or global structural changes one biomolecule at a time.

Time-resolved single-molecule protein-induced fluorescence enhancement is a useful fluorescence spectroscopic proximity sensor sensitive to local structural changes in proteins. Here we show it can be used to uncover stable local conformations in &#945;-Synuclein, which is otherwise known as globularly unstructured and unstable when measured using the longer range FRET ruler.

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural ...

F&#246;rster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between biomolecules in vitro as well as within cells. In FRET, the efficiency of energy transfer, measured by changes in fluorescence intensity or lifetime, relates to the distance between two fluorescent molecules or labels. Determination of dynamics and conformational changes from the distances are just some examples of applications of this method to biological systems. Under certain conditions, this methodology can add to and enhance existing X-ray crystal structures by providing information regarding dynamics, flexibility, and adaptation to binding surfaces. We describe the use of FRET and associated distance determinations to elucidate structural properties, through the identification of a binding site or the orientations of dimer subunits. Through judicious choice of labeling sites, and often employment of multiple labeling strategies, we have successfully applied these mapping methods to determine global structural properties in a protein-DNA complex and the SecA-SecYEG protein translocation system. In the SecA-SecYEG system, we have used FRET mapping methods to identify the preprotein-binding site and determine the local conformation of the bound signal sequence region. This study outlines the steps for performing FRET mapping studies, including identification of appropriate labeling sites, discussion of possible labels including non-native amino acid residues, labeling procedures, how to perform measurements, and interpreting the data.

The study details the methodology of FRET mapping including the selection of labeling sites, choice of dyes, acquisition, and data analysis. This methodology is effective at determining binding sites, conformational changes, and dynamic motions in protein systems and is most useful if performed in conjunction with existing 3-D structural information.

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between ...

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. ...

Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

Summary

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Chapters in this video

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

Quantitative Immunofluorescence to Measure Global Localized Translation

A Fluorescence-based Method to Study Bacterial Gene Regulation in Infected Tissues

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

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

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One α-Synuclein Monomer at a Time

Förster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

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