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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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...

Watch this Scientific Journal Video about Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis at JoVE.com

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

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

Protein-protein interactions play an essential role in the function of a living organism. Once an interaction has been identified and validated it is necessary to characterize it at the structural and mechanistic level. Several biochemical and biophysical methods exist for such purpose. Among them, fluorescence anisotropy is a powerful technique particularly used when the fluorescence intensity of a fluorophore-labeled protein remains constant upon protein-protein interaction. In this technique, a fluorophore-labeled protein is excited with vertically polarized light of an appropriate wavelength that selectively excites a subset of the fluorophores according to their relative orientation with the incoming beam. The resulting emission also has a directionality whose relationship in the vertical and horizontal planes defines anisotropy (r) as follows: r=(IVV-IVH)/(IVV+2IVH), where IVV and IVH are the fluorescence intensities of the vertical and horizontal components, respectively. Fluorescence anisotropy is sensitive to the rotational diffusion of a fluorophore, namely the apparent molecular size of a fluorophore attached to a protein, which is altered upon protein-protein interaction. In the present text, the use of fluorescence anisotropy as a tool to study protein-protein interactions was exemplified to address the binding between the protein mutated in the Shwachman-Diamond Syndrome (SBDS) and the Elongation factor like-1 GTPase (EFL1). Conventionally, labeling of a protein with a fluorophore is carried out on the thiol groups (cysteine) or in the amino groups (the N-terminal amine or lysine) of the protein. However, SBDS possesses several cysteines and lysines that did not allow site directed labeling of it. As an alternative technique, the dye 4&#39;,5&#39;-bis(1,3,2 dithioarsolan-2-yl) fluorescein was used to specifically label a tetracysteine motif, Cys-Cys-Pro-Gly-Cys-Cys, genetically engineered in the C-terminus of the recombinant SBDS protein. Fitting of the experimental data provided quantitative and mechanistic information on the binding mode between these proteins.

Protein interactions are at the heart of a cell&#39;s function. Calorimetric and spectroscopic techniques are commonly used to characterize them. Here we describe fluorescence anisotropy as a tool to study the interaction between the protein mutated in the Shwachman-Diamond Syndrome (SBDS) and the Elongation factor-like 1 GTPase (EFL1).

Protein-protein interactions play an essential role in the function of a living organism. Once an interaction has been identified and validated it is ...

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 ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing approaches suffer from significant limitations, we have developed a new method for determining TF-DNA binding affinities with high sensitivity on a large scale. The assay relies on the established fluorescence anisotropy (FA) principle but introduces important technical improvements. First, we measure a full FA competitive titration curve in a single well by incorporating TF and a fluorescently labeled reference DNA in a porous agarose gel matrix. Unlabeled DNA oligomer is loaded on the top as a competitor and, through diffusion, forms a spatio-temporal gradient. The resulting FA gradient is then read out using a customized epifluorescence microscope setup. This improved setup greatly increases the sensitivity of FA signal detection, allowing both weak and strong binding to be reliably quantified, even for molecules of similar molecular weights. In this fashion, we can measure one titration curve per well of a multi-well plate, and through a fitting procedure, we can extract both the absolute dissociation constant (KD) and active protein concentration. By testing all single-point mutation variants of a given consensus binding sequence, we can survey the entire binding specificity landscape of a TF, typically on a single plate. The resulting position weight matrices (PWMs) outperform those derived from other methods in predicting in vivo TF occupancy. Here, we present a detailed guide for implementing HiP-FA on a conventional automated fluorescent microscope and the data analysis pipeline.

Here we present a novel method for determining binding affinities at equilibrium and in solution with high sensitivity on a large scale. This improves the quantitative analysis of transcription factor-DNA binding. The method is based on automated fluorescence anisotropy measurements in a controlled delivery system.

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing ...

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. ...

OaAEP1-Mediated Enzymatic Synthesis and Immobilization of Polymerized Protein for Single-Molecule Force Spectroscopy

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled protein polymerization process is always a challenge. Here, we have developed an enzymatic methodology for constructing polymerized protein step by step in a rationally-controlled sequence. In this method, the C-terminus of a protein monomer is NGL for protein conjugation using OaAEP1 (Oldenlandia affinis asparaginyl endopeptidases) 1) while the N-terminus was a cleavable TEV (tobacco etch virus) cleavage site plus an L (ENLYFQ/GL) for temporary N-terminal protecting. Consequently, OaAEP1 was able to add only one protein monomer at a time, and then the TEV protease cleaved the N-terminus between Q and G to expose the NH2-Gly-Leu. Then the unit is ready for next OaAEP1 ligation. The engineered polyprotein is examined by unfolding individual protein domain using atomic force microscopy-based single-molecule force spectroscopy (AFM-SMFS). Therefore, this study provides a useful strategy for polyprotein engineering and immobilization.

Here, we present a protocol to conjugate protein monomer by enzymes forming protein polymer with a controlled sequence and immobilize it on the surface for single-molecule force spectroscopy studies.

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled ...

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...

F&#246;rster Resonance Energy Transfer Measurements in Living Plant Cells

Sensitized emission-based F&#246;rster resonance energy transfer (FRET) experiments are easily done but depend on the microscopic setup. Confocal laser scanning microscopes have become a workhorse for biologists. Commercial systems offer high flexibility in laser power adjustment and detector sensitivity and often combine different detectors to obtain the perfect image. However, the comparison of intensity-based data from different experiments and setups is often impossible due to this flexibility. Biologist-friendly procedures are of advantage and allow for simple and reliable adjustment of laser and detector settings. 
Furthermore, as FRET experiments in living cells are affected by the variability in protein expression and donor-acceptor ratios, protein expression levels must be considered for data evaluation. Described here is a simple protocol for reliable and reproducible FRET measurements, including routines for the estimation of protein expression and adjustment of laser intensity and detector settings. Data evaluation will be performed by calibration with a fluorophore fusion of known FRET efficiency. To improve simplicity, correction factors have been compared that have been obtained in cells and by measuring recombinant fluorescent proteins.

A protocol is provided for setting up a standard confocal laser-scanning microscope for in vivo F&#246;rster resonance energy transfer measurements, followed by data evaluation.

Sensitized emission-based F&#246;rster resonance energy transfer (FRET) experiments are easily done but depend on the microscopic setup. Confocal laser ...