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>

<ol>
	<li>Ramakrishnan, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+we+have+learned+from+ribosome+structures.">What we have learned from ribosome structures.</a> Biochemical Society Transactions. 36, 567-574 (2008).</li><li>Frank, J., Agrawal, R. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+ratchet-like+inter-subunit+reorganization+of+the+ribosome+during+translocation.">A ratchet-like inter-subunit reorganization of the ribosome during translocation.</a> Nature. 406 (6793), 318-322 (2000).</li><li>Moazed, D., Noller, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intermediate+states+in+the+movement+of+transfer+RNA+in+the+ribosome.">Intermediate states in the movement of transfer RNA in the ribosome.</a> Nature. 342 (6246), 142-148 (1989).</li><li>Schmeing, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+crystal+structure+of+the+ribosome+bound+to+EF-Tu+and+aminoacyl-tRNA.">The crystal structure of the ribosome bound to EF-Tu and aminoacyl-tRNA.</a> Science. 326 (5953), 688-694 (2009).</li><li>Valle, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Locking+and+unlocking+of+ribosomal+motions.">Locking and unlocking of ribosomal motions.</a> Cell. 114 (1), 123-134 (2003).</li><li>Gromadski, K. B., Rodnina, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetic+determinants+of+high-fidelity+tRNA+discrimination+on+the+ribosome.">Kinetic determinants of high-fidelity tRNA discrimination on the ribosome.</a> Molecular Cell. 13 (2), 191-200 (2004).</li><li>Blanchard, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=tRNA+dynamics+on+the+ribosome+during+translation.">tRNA dynamics on the ribosome during translation.</a> Proceedings of the National Academy of Sciences of the United States of America. 101 (35), 12893-12898 (2004).</li><li>Blanchard, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=tRNA+selection+and+kinetic+proofreading+in+translation.">tRNA selection and kinetic proofreading in translation.</a> Nature Structural and Molecular Biology. 11 (10), 1008-1014 (2004).</li><li>Wang, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+structural+dynamics+of+ef-g-ribosome+interaction+during+translocation.">Single-molecule structural dynamics of ef-g-ribosome interaction during translocation.</a> Biochemistry. 46 (38), 10767-10775 (2007).</li><li>Cornish, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Following+the+movement+of+the+L1+stalk+between+three+functional+states+in+single+ribosomes.">Following the movement of the L1 stalk between three functional states in single ribosomes.</a> Proceedings of the National Academy of Sciences of the United States of America. 106 (8), 2571-2576 (2009).</li><li>Cornish, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spontaneous+intersubunit+rotation+in+single+ribosomes.">Spontaneous intersubunit rotation in single ribosomes.</a> Molecular Cell. 30 (5), 578-588 (2008).</li><li>Chen, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordinated+conformational+and+compositional+dynamics+drive+ribosome+translocation.">Coordinated conformational and compositional dynamics drive ribosome translocation.</a> Nature Structural and Molecular Biology. 20 (6), 718-727 (2013).</li><li>Feng, X. A., Poyton, M. F., Ha, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multicolor+single-molecule+FRET+for+DNA+and+RNA+processes.">Multicolor single-molecule FRET for DNA and RNA processes.</a> Current Opinion in Structural Biology. 70, 26-33 (2021).</li><li>Ly, C. T., Altuntop, M. E., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+study+of+viomycin's+inhibition+mechanism+on+ribosome+translocation.">Single-molecule study of viomycin's inhibition mechanism on ribosome translocation.</a> Biochemistry. 49 (45), 9732-9738 (2010).</li><li>Altuntop, M. E., Ly, C. T., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+study+of+ribosome+hierarchic+dynamics+at+the+peptidyl+transferase+center.">Single-molecule study of ribosome hierarchic dynamics at the peptidyl transferase center.</a> Biophysical Journal. 99 (9), 3002-3009 (2010).</li><li>Wang, Y., Xiao, M., Li, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterogeneity+of+single+molecule+FRET+signals+reveals+multiple+active+ribosome+subpopulations.">Heterogeneity of single molecule FRET signals reveals multiple active ribosome subpopulations.</a> Proteins. 82 (1), 1-9 (2014).</li><li>Xiao, M., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=L27-tRNA+interaction+revealed+by+mutagenesis+and+pH+titration.">L27-tRNA interaction revealed by mutagenesis and pH titration.</a> Biophysical Chemistry. 167, 8-15 (2012).</li><li>Wang, Y., Xiao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+the+ribosomal+protein+L27+revealed+by+single-molecule+FRET+study.">Role of the ribosomal protein L27 revealed by single-molecule FRET study.</a> Protein Science. 21 (11), 1696-1704 (2012).</li><li>Lin, R., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+smFRET+microscope+for+the+quantification+of+label-free+DNA+oligos.">Automated smFRET microscope for the quantification of label-free DNA oligos.</a> Biomedical Optics Express. 10 (2), 682-693 (2019).</li><li>Moazed, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interconversion+of+active+and+inactive+30+S+ribosomal+subunits+is+accompanied+by+a+conformational+change+in+the+decoding+region+of+16+S+rRNA.">Interconversion of active and inactive 30 S ribosomal subunits is accompanied by a conformational change in the decoding region of 16 S rRNA.</a> Journal of Molecular Biology. 191 (3), 483-493 (1986).</li><li>Wower, I. K., Wower, J., Zimmermann, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ribosomal+protein+L27+participates+in+both+50+S+subunit+assembly+and+the+peptidyl+transferase+reaction.">Ribosomal protein L27 participates in both 50 S subunit assembly and the peptidyl transferase reaction.</a> Journal of Biological Chemistry. 273 (31), 19847-19852 (1998).</li><li>Pan, D., Qin, H., Cooperman, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synthesis+and+functional+activity+of+tRNAs+labeled+with+fluorescent+hydrazides+in+the+D-loop.">Synthesis and functional activity of tRNAs labeled with fluorescent hydrazides in the D-loop.</a> RNA. 15 (2), 346-354 (2009).</li><li>Yang, H., Xiao, M., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+data-driven+algorithm+to+reveal+and+track+the+ribosome+heterogeneity+in+single+molecule+studies.">A novel data-driven algorithm to reveal and track the ribosome heterogeneity in single molecule studies.</a> Biophysical Chemistry. 199, 39-45 (2015).</li><li>Metskas, L. A., Rhoades, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+FRET+of+intrinsically+disordered+proteins.">Single-molecule FRET of intrinsically disordered proteins.</a> Annual Review of Physical Chemistry. 71, 391-414 (2020).</li><li>Hellenkamp, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precision+and+accuracy+of+single-molecule+FRET+measurements-a+multi-laboratory+benchmark+study.">Precision and accuracy of single-molecule FRET measurements-a multi-laboratory benchmark study.</a> Nature Methods. 15 (9), 669-676 (2018).</li><li>Tsai, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-Efficiency+"-1"+and+"-2"+Ribosomal+Frameshiftings+Revealed+by+Force+Spectroscopy.">High-Efficiency "-1" and "-2" Ribosomal Frameshiftings Revealed by Force Spectroscopy.</a> ACS Chemical Biology. 12 (6), 1629-1635 (2017).</li><li>Yin, H., Xu, S., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+DNA+rulers+to+study+the+mechanism+of+ribosome+translocation+with+single-nucleotide+resolution.">Dual DNA rulers to study the mechanism of ribosome translocation with single-nucleotide resolution.</a> Journal of Visualized Experiments: JoVE. (149), e59918 (2019).</li><li>Desai, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Co-temporal+force+and+fluorescence+measurements+reveal+a+ribosomal+gear+shift+mechanism+of+translation+regulation+by+structured+mRNAs.">Co-temporal force and fluorescence measurements reveal a ribosomal gear shift mechanism of translation regulation by structured mRNAs.</a> Molecular Cell. 75 (5), 1007-1019 (2019).</li></ol>

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 ...

single-molecule-fluorescence-energy-transfer-study-ribosome-protein

Research

Education

JoVE Journal

JoVE Core

Cell Biology

Biochemistry

Unit 1

Transmembrane Transport in Endoplasmic Reticulum and Peroxisomes

SCIENTISTS IN ACTION

2.5K 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 ...

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 ...

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 ...

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 ...

An Optimized Single-Molecule Pull-Down Assay for Quantification of Protein Phosphorylation

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to measure protein phosphorylation cannot preserve the heterogeneity in phosphorylation across individual proteins. The single-molecule pull-down (SiMPull) assay was developed to investigate the composition of macromolecular complexes via immunoprecipitation of proteins on a glass coverslip followed by single-molecule imaging. The current technique is an adaptation of SiMPull that provides robust quantification of the phosphorylation state of full-length membrane receptors at the single-molecule level. Imaging thousands of individual receptors in this way allows for quantifying protein phosphorylation patterns. The present protocol details the optimized SiMPull procedure, from sample preparation to imaging. Optimization of glass preparation and antibody fixation protocols further enhances data quality. The current protocol provides code for the single-molecule data analysis that calculates the fraction of receptors phosphorylated within a sample. While this work focuses on phosphorylation of the epidermal growth factor receptor (EGFR), the protocol can be generalized to other membrane receptors and cytosolic signaling molecules.

The present protocol describes sample preparation and data analysis to quantify protein phosphorylation using an improved single-molecule pull-down (SiMPull) assay.

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to ...

Studying TRF1/2 Interactions with Telomeric DNA Using Single-Molecule Assay

Analyzing Telomeric Protein-DNA Interactions Using Single-Molecule Magnetic Tweezers

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper function depends on tightly regulated processes of replication, elongation, and damage response. The shelterin complex, especially Telomere Repeat-binding Factor 1 (TRF1) and TRF2, plays a pivotal role in telomere protection and has emerged as a potential anti-cancer target for drug discovery. These proteins bind to the repetitive telomeric DNA motif TTAGGG, facilitating the formation of protective structures and recruitment of other telomeric proteins. Structural methods and advanced imaging techniques have provided insights into telomeric protein-DNA interactions, but probing the dynamic processes requires single-molecule approaches. Tools like magnetic tweezers, optical tweezers, and atomic force microscopy (AFM) have been employed to study telomeric protein-DNA interactions, revealing important details such as TRF2-dependent DNA distortion and telomerase catalysis. However, the preparation of single-molecule constructs with telomeric repetitive motifs continues to be a challenging task, potentially limiting the breadth of studies utilizing single-molecule mechanical methods. To address this, we developed a method to study interactions using full-length human telomeric DNA with magnetic tweezers. This protocol describes how to express and purify TRF2, prepare telomeric DNA, set up single-molecule mechanical assays, and analyze data. This detailed guide will benefit researchers in telomere biology and telomere-targeted drug discovery.

Author Spotlight: Advanced Single-Molecule Techniques for Investigating Telomeric Protein-DNA Interactions

This protocol demonstrates using single-molecule magnetic tweezers to study interactions between telomeric DNA-binding proteins (Telomere Repeat-binding Factor 1 [TRF1] and TRF2) and long telomeres extracted from human cells. It describes the preparatory steps for telomeres and telomeric repeat-binding factors, the execution of single-molecule experiments, and the data collection and analysis methods.

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper ...

In Situ Nucleosome Reconstitution for Single-Molecule Studies

In Situ Nucleosome Assembly for Single-Molecule Correlative Force and Fluorescence Microscopy

Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding their biophysical properties and interactions with chromatin-binding proteins. Nucleosome reconstitution on DNA for these studies typically involves a salt dialysis procedure that provides precise control over the placement and number of nucleosomes formed along a DNA tether. However, this protocol is time-consuming and requires a substantial amount of DNA and histone octamers as inputs. To offer an alternative strategy, an in situ nucleosome reconstitution method for single-molecule force and fluorescence microscopy that utilizes the histone chaperone Nap1 is described. This method enables users to assemble nucleosomes on any DNA template without the need for strong nucleosome positioning sequences, adjust nucleosome density on demand, and use fewer reagents. In situ nucleosome formation occurs within seconds, offering a simpler experimental workflow and a convenient transition into single-molecule measurements. Examples of two downstream assays for probing nucleosome mechanics and visualizing the behavior of individual proteins on chromatin are further described.

Author Spotlight: Efficient Nucleosome Reconstitution for Single-Molecule Techniques

This article presents a detailed experimental procedure for reconstituting nucleosome-containing DNA tethers for single-molecule correlative force and fluorescence microscopy. It further describes several downstream experiments that can be conducted to visualize the binding behavior of chromatin-interacting proteins and analyze changes in the physical properties of nucleosomes.

Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding ...