This work was supported by the National Science Foundation grants MCB-1244297 and MCB-1517245 to PEM.

1. Protein Preparation
<ol>
	<li>DNA cloning.
	<ol>
		<li>Synthesize a DNA sequence of interest, for example, the DNA sequence of NI10C10, or isolate via PCR from the host organism using standard molecular biology techniques11. Flank the gene of interest with restriction sites during synthesis or by placing sites in the 5&#8217;- end of the PCR primers to correspond to a module in the plasmid pEMI91 (Addgene #74888)12.</li>
		<li>Separately digest both the plasmid pEMI9112 and the DNA sequence of interest with a pair of restriction sites so that the sequence of interest will be flanked by the tandem I91 repeats (see Figure 1). Follow the standard protocol for the restriction sites.</li>
		<li>Purify the digested product using gel electrophoresis and then ligate the products using T4 DNA ligase, following a standard protocol. After ligation, transform the plasmid into E. coli cells for plasmid purification and purify the plasmid using standard protocols. Sequence using T7 primers or internal pEMI91 primers to verify that the sequence was successfully transformed.</li>
	</ol>
	</li>
	<li>Transformation.
	<ol>
		<li>Remove protein expression cells, such as C41 (DE3) pLysS cells, from a -80 &#176;C freezer and thaw completely on ice.</li>
		<li>Add 1 &#181;L of plasmid DNA to the cells and stir briefly; pipetting up and down is not advisable as it will introduce air bubbles and warm the cells.</li>
		<li>Incubate the culture tube containing the cells and the plasmid on ice for 30 min.</li>
		<li>Heat shock the cells in a water bath at 42 &#176;C for 45 s.</li>
		<li>Return the tube to ice for 2 min.</li>
		<li>Place 950 &#181;L of LB broth in the cells and shake at 250 rpm for 1 h at 37 &#176;C.</li>
		<li>Place about 200 &#181;L of the transformation onto LB plates containing 100 &#181;g/mL Ampicillin. If using a plasmid other than pEMI91, then use the appropriate antibiotic for that plasmid. Place plate overnight in an incubator at 37 &#176;C. The next day, remove plate from incubator, wrap with parafilm, and store refrigerated up to 1 month.</li>
	</ol>
	</li>
	<li>Protein expression.
	<ol>
		<li>Inoculate 15 mL of LB broth medium with 100 &#181;g/mL ampicillin in a 50 mL tube at 37 &#176;C for overnight growth by touching a sterile pipette tip to a single bacterial colony on the plate and pipetting up and down in the LB.</li>
		<li>Transfer the 15 mL culture into 1 L of LB broth medium with 100 &#181;g/mL ampicillin and shake for 4-12 h at 37 &#176;C (until OD600 &#62; 0.8).</li>
		<li>Collect 10 mL of the culture for plasmid preparation.</li>
		<li>Add 0.2-1 mM isopropyl-&#946;-D-thiogalactopyranoside (IPTG) and lower the temperature to room temperature for overnight expression.</li>
		<li>Harvest cells the next day by splitting into four tubes and centrifuging at 4,000 &#215; g for 40 min, and then freeze the pellet at -80 &#176;C for several hours.</li>
		<li>Send the plasmids extracted from the cells for sequencing with the primers corresponding to the inserted cassette of the protein12 and verify the fidelity of the inserted DNA.</li>
	</ol>
	</li>
	<li>Protein purification.
	<ol>
		<li>Thaw one tube of frozen cells for 30 min at room temperature.</li>
		<li>Suspend thawed cells in lysis buffer (38 mL of water, 2 mL of glycerol, 50 mM Tris-HCl pH 7.6, 150 mM NaCl, 1 mM CaCl2, 10 &#181;g/mL DNase, 1 mM PMSF, 1 mM TCEP, 500 &#181;g/mL lysozyme) and shake on ice for 1 h.</li>
		<li>Freeze the lysed cells at -80 &#176;C for several hours and then re-thaw at room temperature.</li>
		<li>Spin the lysate down at 13,100 &#215; g for 30 min at 4 &#176;C.</li>
		<li>Run the supernatant through a gravity flow column for the specific tag (e.g., Strep-tag or His-tag).</li>
		<li>Perform a buffer exchange using a centrifugal filter using the appropriate buffer for Strep-tag or His-tag and store at 4 &#176;C before use.</li>
	</ol>
	</li>
</ol>
2. Slides Preparation for Sample Preparation
<ol>
	<li>Prepare glass slides using Piranha solution.
	<ol>
		<li>Place 10-30 pieces of round glass cover slips (radius of 7.5 mm) into a 40 mL glass beaker. 
		Note: Perform this step and following steps in a hood. Follow standard procedures for dealing with dangerous chemicals during this step.</li>
		<li>Add 30 mL of concentrated (18.4 M) sulfuric acid.</li>
		<li>Add 10 mL of 30% hydrogen peroxide.</li>
		<li>Heat the mixture for 10-30 min to 95 &#176;C.</li>
		<li>Carefully decant the Piranha solution into a separate waste container (to be reused or discarded).</li>
		<li>Rinse the slides with deionized water to remove Piranha solution.</li>
		<li>Suspend the slides in 40 mL of acetone.</li>
		<li>Decant acetone into a waste container and re-suspend slides in 40 mL of ethanol.</li>
		<li>Use clean forceps to carefully extract one slide at a time and dry with argon or purified air.</li>
		<li>Place clean and dried slides under vacuum until the gold evaporation, or under argon for long-term storage.</li>
	</ol>
	</li>
	<li>Prepare gold coated slides (optional). 
	Note: This step requires the use of a vacuum (e-beam) evaporator, which is available in many clean room facilities at major universities.
	<ol>
		<li>Cut five microscope slides (25 mm x 75 mm rectangular shape) in half using a glasscutter.</li>
		<li>Apply double-sided adhesive tape across the middle of each of the 10 half microscope slides.</li>
		<li>Cut the sticky side of a sticky note and apply to the double-sided tape so that the sticky side of the note is face-up. The sticky note is generally less adhesive, but still firmly attaches to slides to prevent future breaking. Then those half microscope slides can now work as glass slides holder.</li>
		<li>Carefully press four clean glass slides (from section 2.1.10) to each corner of the sticky side of the holder.</li>
		<li>Move the glass slides to an e-beam evaporator. Follow the specifications of the evaporator to apply 70 nm of chromium and 300 nm of gold to the surface.</li>
		<li>Store the gold-coated slides under argon.</li>
	</ol>
	</li>
</ol>

3. Sample Preparation
<ol>
	<li>Prepare slide.
	<ol>
		<li>Select a clean iron disk (15 mm diameter) and attach an adhesive tab to it.</li>
		<li>Unattach the cover piece of the adhesive tab.</li>
		<li>Select a piece of clean glass (from section 2.1.10) or gold (from section 2.2.7) and firmly place it onto the sticky side of the iron disk, this will be the sample slide.</li>
	</ol>
	</li>
	<li>Deposit protein onto slide.
	<ol>
		<li>Dialyze the polyprotein into the buffer for the experiment (25 mM Tris-HCl, pH 7.6 with 150 mM NaCl) using a buffer exchange column such as a 0.5-mL centrifugal filter. Spin the protein in the column for 10 min at 13,000 rpm and then reverse the column and elute the protein into a new buffer by spinning at 1,000 rpm for 2 min.</li>
		<li>Determine the approximate protein concentration by measuring the absorbance at 280 nm, using a spectrophotometer.</li>
		<li>Dilute the protein to 10-100 &#181;g/mL in a final volume of 100 &#181;L.</li>
		<li>Apply the 60&#160;&#181;L of protein solution onto the center of the slide. Be careful at this stage not to let any liquid enter beneath into the gap between the glass and the iron slide, as this can cause swelling of the adhesive during the experiment and uncontrolled sample movements. 
		Note: Gold slides are hydrophobic, and a spherical droplet will form, while glass slides are more hydrophilic and the protein solution may spread.</li>
		<li>Let the sample sit at room temperature for 10-60 min. During this time, proceed to the next step to begin setting up the atomic force microscope.</li>
	</ol>
	</li>
</ol>

4. Atomic Force Microscope (AFM) Setup
Note: The following is a general description for setting up the AFM, and some specific details may differ depending on the specific instrumentation used. The instrumentation used is partially home-built and described in detail in Scholl13.
<ol>
	<li>Mount the cantilever onto an AFM cell.
	<ol>
		<li>Choose the cantilever with appropriate properties for the application. Use cantilevers with spring constants 4-10 pN/nm for low unfolding forces (~10-50 pN), while use cantilevers with spring constants 15-100 pN/nm for high unfolding forces.</li>
		<li>Carefully pick up the cantilever from the end and place it into the probe holding cell.</li>
		<li>Make sure the holding cell firmly holds the cantilever in place before continuing.</li>
		<li>Place the holding cell into the AFM head for alignment of the laser.</li>
	</ol>
	</li>
	<li>Align the laser in the AFM head.
	<ol>
		<li>Place the head onto an inverted microscope and connect a battery pack to the AFM head for powering the laser in the head.</li>
		<li>Attach a camera to the microscope detector so that the laser light can be visualized onto a TV or monitor.</li>
		<li>Position the laser so that it is located onto the tip of the cantilever</li>
	</ol>
	</li>
	<li>Mount head with sample.
	<ol>
		<li>Flush 10 &#181;L of buffer into each of the ports of the probe holding cell.</li>
		<li>Take the sample slide which has been incubating and decant 40 &#181;L of fluid from the incubated solution. Add 40 &#181;L of the buffer on the slide. Place the sample slide onto the magnet above the piezo.</li>
		<li>Make sure that the AFM stage is in the elevated position. Then place the AFM head onto the stage so that the AFM cantilever is above the sample droplet.</li>
	</ol>
	</li>
	<li>Center the reflected laser beam onto the photodiode.
	<ol>
		<li>Cut a small piece of paper and place it in front of the AFM photodiode.</li>
		<li>Reposition the AFM laser using the knobs so that the laser spot on the paper becomes focused and bright.</li>
		<li>Adjust the AFM head mirror so that the laser hits the photodiode to maximize the total signal in all quadrants (A+B+C+D) and causes the difference signal between top two and bottom two quadrants to be zero (A+B-C-D).</li>
	</ol>
	</li>
</ol>

5. Atomic Force Microscope Calibration
<ol>
	<li>Measure the power spectrum.
	<ol>
		<li>On the filter connected to the AFM, turn the filter setting for the AFM head signal to full bandwidth.</li>
		<li>On the AFM itself, make sure that the piezo is off as this will add noise to the signal.</li>
		<li>Use the AFM software14 to measure the average of 512 calculations of the power spectrum from 1,024 data points per calculation.</li>
		<li>Use the AFM software15 to integrate the power spectral density across the first peak, which corresponds to the main mode of vibration for the cantilever.</li>
	</ol>
	</li>
	<li>Calculate photodiode sensitivity.
	<ol>
		<li>On the AFM itself, turn the piezo controller on and change the filter setting to 500 Hz (low-pass filter).</li>
		<li>Monitor the difference signal which should be fluctuating around zero in the AFM software. Swiftly move the AFM head down a few hundred micrometers using micropositioners. Repeat moving the head down and monitor the difference signal for consistent jumps. The surface is very nearby when the signal jumps start increasing in height.</li>
		<li>As soon as the deflection signal saturates upon contact with the surface, move the head away from the surface slightly.</li>
		<li>In the AFM software, use the input controls to regulate the piezo voltage to find the surface, by moving up or down 100-5,000 nm. If the surface is still not in reach, continue to reduce the distance between the head and the sample manually.</li>
		<li>In the AFM software, conduct a pulling experiment with a scan size of about 500 nm so that the tip comes in contact for about 100 nm of piezo travel.</li>
		<li>Measure the slope of the linear region of the photodiode signal versus piezo displacement curve where the AFM tip remains in contact with the substrate surface.</li>
		<li>Calculate the spring constant and sensitivity using the slope and the integrated power spectrum (<img alt="Equation 1" src="/files/ftp_upload/55989/55989eq1.jpg" />).</li>
	</ol>
	</li>
</ol>

6. Data Acquisition
<ol>
	<li>Preparation.
	<ol>
		<li>On the filters connected to the AFM, set the filter setting so that the sampling frequency is at least twice the bandwidth (Nyquist criterion), which will give an upper bound for the low-pass cutoff.</li>
	</ol>
	</li>
	<li>Measurements.
	<ol>
		<li>Set the scan size to the total theoretical size of the unfolding polyprotein (number of amino acids &#215; 0.365) plus about 40% to allow for pressing against the substrate. For example, pulling a 9x I91 construct will have a theoretical unfolded length of about 300 nm, so the scan size should be about 420 nm.</li>
		<li>In the AFM software, position the cantilever so that it is 80% of the scan-size away from the surface (in this case, about 340 nm away from the surface).</li>
		<li>In the AFM software, set the scan speed to 300 nm/s initially. 
		Note: This speed can be modified to be slower, or faster, depending on the application.</li>
		<li>Perform a pulling experiment and measure the resulting position of the piezo and the photodiode signal. Use the AFM software to initiate the scan.</li>
		<li>In the AFM software, continue to perform measurements until there are about 10,000 recordings. 
		Note: Generally, the pickup rate of a positive-controlled molecule is around 0.5%16, so 10,000 are necessary to be able to collect enough data to analyze.</li>
	</ol>
	</li>
</ol>

7. Data Analysis
<ol>
	<li>Normalize data.
	<ol>
		<li>For each recording, compute the extension using extension = displacement - F/kc (kc is from 5.2.7).</li>
		<li>Determine the baseline force level by taking the average of either the beginning or end of the force-extension trace, where no force peaks are present and where the cantilever is not pressing against the surface. The entire force-extension curve can then be shifted by this mean value.</li>
		<li>Translate the data so that the extension is aligned along the zero y-axis. This is because the molecule should be set to zero extension at the beginning of a trace.</li>
	</ol>
	</li>
	<li>Identifying protein of interest.
	<ol>
		<li>Identify recordings with at least four I91 events that provide a single-molecule fingerprint. These recordings capture true &#8220;single-molecule measurements&#8221;.</li>
		<li>Save events that are denoted single-molecule events for further analysis.</li>
	</ol>
	</li>
	<li>Determine contour-length increment.
	<ol>
		<li>For each recording, fit a worm-like chain model to unfolding event, <img alt="Equation 2" src="/files/ftp_upload/55989/55989eq2.jpg" /> where <img alt="Equation 3" src="/files/ftp_upload/55989/55989eq3.jpg" /> at room temperature, p is the persistence length (typically 0.4 to 1 nm), x is the extension in nanometers, and L is the contour length in nanometers. 
		Note: Form of the worm-like chain is an interpolated solution to the exact, a more exact numerical solution is found in Bouchiat et al.17. Alternate fitting models can be found in Su et al.18.</li>
		<li>Calculate the difference between the values for L determined for two consecutive unfolding events, and this difference is coined the &#8220;contour-length increment&#8221;.</li>
	</ol>
	</li>
	<li>Determine rupture force.
	<ol>
		<li>Calculate the rupture force for a given unfolding event by taking the highest point before the biggest drop in the unfolding curve.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Rief, M., Gautel, M., Oesterhelt, F., Fernandez, J. M., Gaub, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reversible+Unfolding+of+Individual+Titin+Immunoglobulin+Domains+by+AFM.">Reversible Unfolding of Individual Titin Immunoglobulin Domains by AFM.</a> Science. 276 (5315), 1109-1112 (1997).</li><li>Fisher, T. E., Oberhauser, A. F., Carrion-Vazquez, M., Marszalek, P. E., Fernandez, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+study+of+protein+mechanics+with+the+atomic+force+microscope.">The study of protein mechanics with the atomic force microscope.</a> Trends in Biochemical Sciences. 24 (10), 379-384 (1999).</li><li>Ng, S., Rounsevell, R., Steward, A., Randles, L., Clarke, J. <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+studies+of+protein+folding+by+atomic+force+microscopy(AFM).">Single molecule studies of protein folding by atomic force microscopy(AFM).</a> Abstracts of Papers of the American Chemical Society. 227, U545-U545 (2004).</li><li>Rico, F., Chu, C., Moy, V. T., Braga, P. C., Ricci, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods in Molecular Biology. 736, 331-353 (2011).</li><li>Muller, D. J., Dufrene, Y. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic+force+microscopy+as+a+multifunctional+molecular+toolbox+in+nanobiotechnology.">Atomic force microscopy as a multifunctional molecular toolbox in nanobiotechnology.</a> Nature Nanotechnology. 3 (5), 261-269 (2008).</li><li>Lv, 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=Designed+biomaterials+to+mimic+the+mechanical+properties+of+muscles.">Designed biomaterials to mimic the mechanical properties of muscles.</a> Nature. 465 (7294), 69-73 (2010).</li><li>Kim, 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=Nanomechanics+of+Streptavidin+Hubs+for+Molecular+Materials.">Nanomechanics of Streptavidin Hubs for Molecular Materials.</a> Advanced Materials. 23 (47), 5684-5688 (2011).</li><li>Gonzalez, M. A., 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=Self-Adhesive+Hydrogels+from+Intrinsically+Unstructured+Proteins.">Self-Adhesive Hydrogels from Intrinsically Unstructured Proteins.</a> Advanced Materials. , (2017).</li><li>Oberhauser, A. F., Hansma, P. K., Carrion-Vazquez, M., Fernandez, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stepwise+unfolding+of+titin+under+force-clamp+atomic+force+microscopy.">Stepwise unfolding of titin under force-clamp atomic force microscopy.</a> Proceedings of the National Academy of Sciences. 98 (2), 468-472 (2001).</li><li>Li, Q., Scholl, Z. N., Marszalek, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capturing+the+Mechanical+Unfolding+Pathway+of+a+Large+Protein+with+Coiled-Coil+Probes.">Capturing the Mechanical Unfolding Pathway of a Large Protein with Coiled-Coil Probes.</a> Angewandte Chemie International Edition. 53 (49), 13429-13433 (2014).</li><li>Davis, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Basic methods in molecular biology. , (2012).</li><li>Scholl, Z. N., Josephs, E. A., Marszalek, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Modular,+Non-Degenerate+Polyprotein+Scaffold+for+Atomic+Force+Spectroscopy.">A Modular, Non-Degenerate Polyprotein Scaffold for Atomic Force Spectroscopy.</a> Biomacromolecules. , (2016).</li><li>Scholl, Z. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The (Un) Folding of Multidomain Proteins Through the Lens of Single-molecule Force-spectroscopy and Computer Simulation. , (2016).</li><li>Pawlak, K., Strzelecki, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanopuller-open+data+acquisition+platform+for+AFM+force+spectroscopy+experiments.">Nanopuller-open data acquisition platform for AFM force spectroscopy experiments.</a> Ultramicroscopy. 164, 17-23 (2016).</li><li>. Nanopuller Available from: <a target="_blank" href="https://sourceforge.net/projects/nanopuller/">https://sourceforge.net/projects/nanopuller/</a> (2018)</li><li>Scholl, Z. N., Marszalek, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improving+single+molecule+force+spectroscopy+through+automated+real-time+data+collection+and+quantification+of+experimental+conditions.">Improving single molecule force spectroscopy through automated real-time data collection and quantification of experimental conditions.</a> Ultramicroscopy. 136, 7-14 (2014).</li><li>Bouchiat, C., 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=Estimating+the+persistence+length+of+a+worm-like+chain+molecule+from+force-extension+measurements.">Estimating the persistence length of a worm-like chain molecule from force-extension measurements.</a> Biophysical journal. 76 (1), 409-413 (1999).</li><li>Su, T., Purohit, P. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanics+of+forced+unfolding+of+proteins.">Mechanics of forced unfolding of proteins.</a> Acta. 5 (6), 1855-1863 (2009).</li><li>Steward, A., Toca-Herrera, J. L., Clarke, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Versatile+cloning+system+for+construction+of+multimeric+proteins+for+use+in+atomic+force+microscopy.">Versatile cloning system for construction of multimeric proteins for use in atomic force microscopy.</a> Protein science. 11 (9), 2179-2183 (2002).</li><li>Scholl, Z. N., Josephs, E. A., Marszalek, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modular,+Nondegenerate+Polyprotein+Scaffolds+for+Atomic+Force+Spectroscopy.">Modular, Nondegenerate Polyprotein Scaffolds for Atomic Force Spectroscopy.</a> Biomacromolecules. 17 (7), 2502-2505 (2016).</li><li>Hoffmann, 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=Rapid+and+Robust+Polyprotein+Production+Facilitates+Single-Molecule+Mechanical+Characterization+of+&#946;-Barrel+Assembly+Machinery+Polypeptide+Transport+Associated+Domains.">Rapid and Robust Polyprotein Production Facilitates Single-Molecule Mechanical Characterization of &#946;-Barrel Assembly Machinery Polypeptide Transport Associated Domains.</a> ACS. 9 (9), 8811-8821 (2015).</li><li>Dudko, O. K., Hummer, G., Szabo, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Theory,+analysis,+and+interpretation+of+single-molecule+force+spectroscopy+experiments.">Theory, analysis, and interpretation of single-molecule force spectroscopy experiments.</a> Proceedings of the National Academy of Sciences of the United States of America. 105 (41), 15755-15760 (2008).</li><li>Popa, I., Berkovich, R., Alegre-Cebollada, J., Rivas-Pardo, J. A., Fernandez, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Halotag+Tethers+to+Study+Titin+Folding+at+the+Single+Molecule+Level.">Halotag Tethers to Study Titin Folding at the Single Molecule Level.</a> Biophysical journal. 106 (2), 391a (2014).</li><li>Yu, H., Siewny, M. G., Edwards, D. T., Sanders, A. W., Perkins, T. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hidden+dynamics+in+the+unfolding+of+individual+bacteriorhodopsin+proteins.">Hidden dynamics in the unfolding of individual bacteriorhodopsin proteins.</a> Science. 355 (6328), 945-950 (2017).</li><li>Rico, F., Gonzalez, L., Casuso, I., Puig-Vidal, M., Scheuring, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-speed+force+spectroscopy+unfolds+titin+at+the+velocity+of+molecular+dynamics+simulations.">High-speed force spectroscopy unfolds titin at the velocity of molecular dynamics simulations.</a> Science. 342 (6159), 741-743 (2013).</li><li>He, Y., Lu, M., Cao, J., Lu, H. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manipulating+protein+conformations+by+single-molecule+AFM-FRET+nanoscopy.">Manipulating protein conformations by single-molecule AFM-FRET nanoscopy.</a> ACS nano. 6 (2), 1221-1229 (2012).</li><li>Fotiadis, D., Scheuring, S., M&#252;ller, S. A., Engel, A., M&#252;ller, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+and+manipulation+of+biological+structures+with+the+AFM.">Imaging and manipulation of biological structures with the AFM.</a> Micron. 33 (4), 385-397 (2002).</li><li>Edwards, D. T., Faulk, J. K., LeBlanc, M. A., Perkins, T. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Force+Spectroscopy+with+9-&#956;s+Resolution+and+Sub-pN+Stability+by+Tailoring+AFM+Cantilever+Geometry.">Force Spectroscopy with 9-&#956;s Resolution and Sub-pN Stability by Tailoring AFM Cantilever Geometry.</a> Biophysical journal. 113 (12), 2595-2600 (2017).</li><li>Dudko, O. K., Mathe, J., Szabo, A., Meller, A., Hummer, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracting+kinetics+from+single-molecule+force+spectroscopy:+Nanopore+unzipping+of+DNA+hairpins.">Extracting kinetics from single-molecule force spectroscopy: Nanopore unzipping of DNA hairpins.</a> Biophysical. 92 (12), 4188-4195 (2007).</li><li>Scholl, Z. N., Li, Q., Yang, W., Marszalek, P. 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+Force+Spectroscopy+Reveals+the+Calcium+Dependence+of+the+Alternative+Conformations+in+the+Native+State+of+a+&#946;&#947;-Crystallin+Protein.">Single-molecule Force Spectroscopy Reveals the Calcium Dependence of the Alternative Conformations in the Native State of a &#946;&#947;-Crystallin Protein.</a> Journal of Biological Chemistry. 291 (35), 18263-18275 (2016).</li></ol>

The authors have nothing to disclose.

A critical step in the protocol is the use of a polyprotein, described in step 1.1.2, which serves as a positive control to &#34;fingerprint&#34; single-molecule events. Generally, there must be unfolding events of the polyprotein proteins (for I91, this means an unfolding force of about 200 pN and contour length increment of about 28 nm) to unambiguously conclude that the protein of interest has been unfolded. For example, when the protein of interest is flanked by three I91 domains from either side, then there must be ...

Advances in single molecule force spectroscopy (SMFS) by AFM have enabled mechanical manipulation and precise characterization of single protein molecules. This characterization has produced novel insights about protein mechanics1,2, protein folding3, protein-ligand interactions4, protein-protein interactions5, and protein-based engineered materials6,7,8. SMFS is especially useful for studying protein unfolding, as stretching by AFM allows the chemical and physical bonds within the protein molecule to gradually extend according to their stiffness, which gives rise to a continually increasing contour length. This overstretching of a protein molecule can produce an abrupt transition in the force-extension curve resulting in a rupture event (or force peak). The force peak gives direct information on the unfolding force and structural change of the protein during the mechanical unfolding process. One of the first studies using AFM measured titin1 and found novel aspects of protein unfolding and refolding under physiological conditions without the use of unnatural denaturants like concentrated chemicals or extreme temperatures.
SMFS experiments are conducted on a variety of instruments, though here we consider only the AFM. The AFM is composed of four main elements: the probe, the detector, the sample holder, and the piezoelectric scanner. The probe is a sharp tip on the free-swinging end of a cantilever. After calibration, bending of the cantilever during stretching of an attached molecule is measured using a laser beam that is reflected off the back of the cantilever to precisely determine forces using Hooke&#39;s law. The reflected laser beam projects into a quadrant photodiode detector which produces a voltage in proportion to the displacement of the laser beam from the diode center. The substrate with the protein sample in fluid is mounted onto a 3D piezoelectric stage that can be controlled with sub-nanometer precision. A computer reads the voltage from the photodiode detectors and controls the 3D stage through a computer-controlled voltage supply. These piezo actuator stages are usually equipped with capacitive or strain-gauge position sensors to precisely measure piezo displacement and to correct hysteresis through feed-back control system. The sensor signal output from the piezo controller is converted into distance using the voltage constant of the piezo that is factory-calibrated. An example force-extension curve from a pulling experiment is shown in Figure 2.
There are two types of AFM-SMFS experiments: constant velocity and constant force pulling measurements. Constant force SMFS measurements are described in Oberhauser et al.9, while here we focus on constant velocity measurements. A typical AFM constant-velocity pulling experiment is done by providing voltage to a piezo to gently move a substrate relative to a cantilever tip. A typical experiment has the tip initially pressing against the surface. The pulling measurement is begun by moving the substrate away from the tip to bring out of contact. If a protein comes into contact with the tip initially, it will be pulled and the unfolding trace of force against displacement will be measured. The substrate is then brought back into contact with the tip and a relaxing trace is measured where protein folding can be determined from the force displacement.

<table>
	<tbody>
		<tr>
			<td>AFM Specimen Discs, 15mm diameter</td>
			<td>Ted Pella, Inc.</td>
			<td>16218</td>
			<td>Serve as base for glass substrate</td>
		</tr>
		<tr>
			<td>Round Glass Coverslips, 15mm diamiter No.1 Thick</td>
			<td>Ted Pella, Inc.</td>
			<td>26024</td>
			<td>serve as glass substrate and base for gold coating</td>
		</tr>
		<tr>
			<td>Adhesive Tabs</td>
			<td>Ted Pella, Inc.</td>
			<td>16079</td>
			<td>Paste on AFM Specimen Discs to provide a sticky face for attaching glass coverslips</td>
		</tr>
		<tr>
			<td>STD Multimode head assembly</td>
			<td>Bruker Nano Inc.</td>
			<td>1B75C</td>
			<td>AFM head</td>
		</tr>
		<tr>
			<td>Glass probe holder</td>
			<td>Bruker Nano Inc.</td>
			<td>MTFML-V2</td>
			<td>Glass probe holder for scanning in fluid with the MultiMode AFM.&#160;&#160;</td>
		</tr>
		<tr>
			<td>Microlever AFM probes</td>
			<td>Bruker Nano Inc.</td>
			<td>MLCT</td>
			<td>Silicon Nitride cantilevers with Silicon Nitride tips, ideal for contact imaging modes</td>
		</tr>
		<tr>
			<td>AFM probes with Au coated tips</td>
			<td>Bruker Nano Inc.</td>
			<td>OBL-10</td>
			<td>Cantilevers for pulling on proteins with low unfolding force</td>
		</tr>
		<tr>
			<td>Multifunction Data Acquisition (DAQ) Card,16-Bit, 1 MS/s (Multichannel), 1.25 MS/s (1-Channel), 32 Analog Inputs</td>
			<td>National Instruments</td>
			<td>PCI-6259</td>
			<td>Data Acquisition for signals from AFM head and Piezo Actuators</td>
		</tr>
		<tr>
			<td>LISA Linear Piezo Stage Actuators</td>
			<td>Physik Instrumente LP</td>
			<td>P-753.11C</td>
			<td>Piezo Actuator to control the position of substrate and perform pulling measurements</td>
		</tr>
		<tr>
			<td>XY Piezo Stage</td>
			<td>Physik Instrumente LP</td>
			<td>P-541.2CD</td>
			<td>Piezo Actuator to control the position of substrate and scan on substrate surface</td>
		</tr>
	</tbody>
</table>

force spectroscopy of single protein molecules using an atomic force microscope

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. Atomic force microscopy (AFM) can address this problem by enabling stretching and relaxation of single protein molecules, which gives direct evidence of specific unfolding and refolding characteristics. AFM-based single-molecule force-spectroscopy (AFM-SMFS) provides a means to consistently measure high-energy conformations in proteins that are not possible in traditional bulk (biochemical) measurements. Although numerous papers were published to show principles of AFM-SMFS, it is not easy to conduct SMFS experiments due to a lack of an exhaustively complete protocol. In this study, we briefly illustrate the principles of AFM and extensively detail the protocols, procedures, and data analysis as a guideline to achieve good results from SMFS experiments. We demonstrate representative SMFS results of single protein mechanical unfolding measurements and we provide troubleshooting strategies for some commonly encountered problems.

Representative results from this protocol are shown in Figure 2. Both panels show representative force-extension curves from proteins. The top shows results from a I91 polyprotein, while the bottom shows the I91 protein flanking a protein-of-interest, the NI10C molecule. These recordings show the characteristic force of I91 (200 pN) and contour length increment (28 nm) which indicates that the alignment and calibration of the AFM was successful. Th...

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Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. ...

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8.6K Views.  Duke University. We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

Video: Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the propagation of neurodegenerative disorders. Such assemblies consist in multiple protein subunits organized in a non-covalent way to form large macromolecular objects that can execute a variety of cellular functions or cause detrimental consequences. Atomic insights into the assembly mechanisms and the functioning of those macromolecular assemblies remain often scarce since their inherent insolubility and non-crystallinity often drastically reduces the quality of the data obtained from most techniques used in structural biology, such as X-ray crystallography and solution Nuclear Magnetic Resonance (NMR). We here present magic-angle spinning solid-state NMR spectroscopy (SSNMR) as a powerful method to investigate structures of macromolecular assemblies at atomic resolution. SSNMR can reveal atomic details on the assembled complex without size and solubility limitations. The protocol presented here describes the essential steps from the production of 13C/15N isotope-labeled macromolecular protein assemblies to the acquisition of standard SSNMR spectra and their analysis and interpretation. As an example, we show the pipeline of a SSNMR structural analysis of a filamentous protein assembly.

Structures of supramolecular protein assemblies at atomic resolution are of high relevance because of their crucial roles in a variety of biological phenomena. Herein, we present a protocol to perform high-resolution structural studies on insoluble and non-crystalline macromolecular protein assemblies by magic-angle spinning solid-state nuclear magnetic resonance spectroscopy (MAS SSNMR).

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the ...

Capturing the Interaction Kinetics of an Ion Channel Protein with Small Molecules by the Bio-layer Interferometry Assay

The bio-layer interferometry (BLI) assay is a valuable tool for measuring protein-protein and protein-small molecule interactions. Here, we first describe the application of this novel label-free technique to study the interaction of human EAG1 (hEAG1) channel proteins with the small molecule PIP2. hEAG1 channel has been recognized as potential therapeutic target because of its aberrant overexpression in cancers and a few gain-of-function mutations involved in some types of neurological diseases. We purified&#160;hEAG1 channel proteins from a mammalian stable expression system and measured the interaction with PIP2 by BLI. The successful measurement of the kinetics of binding between hEAG1 protein and PIP2 demonstrates that the BLI assay is a potential high-throughput approach used for novel small-molecule ligand screening in ion channel pharmacology.

The protocol here describes the interactions of purified hEAG1 ion channel protein with the small molecule lipid ligand phosphatidylinositol 4, 5-bisphosphate (PIP2). The measurement demonstrates that BLI could be a potential method for novel small-molecule ion channel ligand screening.

The bio-layer interferometry (BLI) assay is a valuable tool for measuring protein-protein and protein-small molecule interactions. Here, we first describe ...

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 ...

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Imaging of Extracellular Vesicles by Atomic Force Microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

Atomic Absorbance Spectroscopy to Measure Intracellular Zinc Pools in Mammalian Cells

Transition metals are essential micronutrients for organisms but can be toxic to cells at high concentrations by competing with physiological metals in proteins and generating redox stress. Pathological conditions that lead to metal depletion or accumulation are causal agents of different human diseases. Some examples include anemia, acrodermatitis enteropathica, and Wilson&#8217;s and Menkes&#8217; diseases. It is therefore important to be able to measure the levels and transport of transition metals in biological samples with high sensitivity and accuracy in order to facilitate research exploring how these elements contribute to normal physiological functions and toxicity. Zinc (Zn), for example, is a cofactor in many mammalian proteins, participates in signaling events, and is a secondary messenger in cells. In excess, Zn is toxic and can inhibit absorption of other metals, while in deficit, it can lead to a variety of potentially lethal conditions.

Graphite furnace atomic absorption spectroscopy (GF-AAS) provides a highly sensitive and effective method for determining Zn and other transition metal concentrations in diverse biological samples. Electrothermal atomization via GF-AAS quantifies metals by atomizing small volumes of samples for subsequent selective absorption analysis using wavelength of excitation of the element of interest. Within the limits of linearity of the Beer-Lambert Law, the absorbance of light by the metal is directly proportional to concentration of the analyte. Compared to other methods of determining Zn content, GF-AAS detects both free and complexed Zn in proteins and possibly in small intracellular molecules with high sensitivity in small sample volumes. Moreover, GF-AAS is also more readily accessible than inductively coupled plasma mass spectrometry (ICP-MS) or synchrotron-based X-ray fluorescence. In this method, the systematic sample preparation of different cultured cell lines for analyses in a GF-AAS is described. Variations in this trace element were compared in both whole cell lysates and subcellular fractions of proliferating and differentiated cells as proof of principle.

Cultured primary or established cell lines are commonly used to address fundamental biological and mechanistic questions as an initial approach before using animal models. This protocol describes how to prepare whole cell extracts and subcellular fractions for studies of zinc (Zn) and other trace elements with atomic absorbance spectroscopy.

Transition metals are essential micronutrients for organisms but can be toxic to cells at high concentrations by competing with physiological metals in ...

Evaluation of Protein&#8211;Protein Interactions using an On-Membrane Digestion Technique

Numerous intracellular proteins physically interact in accordance with their intracellular and extracellular circumstances. Indeed, cellular functions largely depend on intracellular protein&#8211;protein interactions. Therefore, research regarding these interactions is indispensable to facilitating the understanding of physiologic processes. Co-precipitation of associated proteins, followed by mass spectrometry (MS) analysis, enables the identification of novel protein interactions. In this study, we have provided details of the novel technique of immunoprecipitation-liquid chromatography (LC)-MS/MS analysis combined with on-membrane digestion for the analysis of protein&#8211;protein interactions. This technique is suitable for crude immunoprecipitants and can improve the throughput of proteomic analyses. Tagged recombinant proteins were precipitated using specific antibodies; next, immunoprecipitants blotted onto polyvinylidene difluoride membrane pieces were subjected to reductive alkylation. Following trypsinization, the digested protein residues were analyzed using LC-MS/MS. Using this technique, we were able to identify several candidate associated proteins. Thus, this method is convenient and useful for the characterization of novel protein&#8211;protein interactions.

Here, we present a protocol for an on-membrane digestion technique for the preparation of samples for mass spectrometry. This technique facilitates the convenient analysis of protein&#8211;protein interactions.

Numerous intracellular proteins physically interact in accordance with their intracellular and extracellular circumstances. Indeed, cellular functions ...

Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy

The phenomenon of protein misfolding and aggregation results in the formation of highly heterogeneous protein aggregates, which are associated with neurodegenerative conditions such as Alzheimer&#8217;s and Parkinson&#8217;s diseases. In particular low molecular weight aggregates, amyloid oligomers, have been shown to possess generic cytotoxic properties and are implicated as neurotoxins in many forms of dementia. We illustrate the use of methods based on atomic force microscopy (AFM) to address the challenging task of characterizing the morphological, structural and chemical properties of these aggregates, which are difficult to study using conventional structural methods or bulk biophysical methods because of their heterogeneity and transient nature. Scanning probe microscopy approaches are now capable of investigating the morphology of amyloid aggregates with sub-nanometer resolution. We show here that infrared (IR) nanospectroscopy (AFM-IR), which simultaneously exploits the high resolution of AFM and the chemical recognition power of IR spectroscopy, can go further and enable the characterization of the structural properties of individual protein aggregates, and thus offer insights into the aggregation mechanisms. Since the approach that we describe can be applied also to the investigations of the interactions of protein assemblies with small molecules and antibodies, it can deliver fundamental information to develop new therapeutic compounds to diagnose or treat neurodegenerative disorders.

We describe the application of infrared nanospectroscopy and high-resolution atomic force microscopy to visualize the process of protein self-assembly into oligomeric aggregates and amyloid fibrils, which is closely associated with the onset and development of a wide range of human neurodegenerative disorders.

The phenomenon of protein misfolding and aggregation results in the formation of highly heterogeneous protein aggregates, which are associated with ...

Creating Highly Specific Chemically Induced Protein Dimerization Systems by Stepwise Phage Selection of a Combinatorial Single-Domain Antibody Library

Protein dimerization events that occur only in the presence of a small-molecule ligand enable the development of small-molecule biosensors for the dissection and manipulation of biological pathways. Currently, only a limited number of chemically induced dimerization (CID) systems exist and engineering new ones with desired sensitivity and selectivity for specific small-molecule ligands remains a challenge in the field of protein engineering. We here describe a high throughput screening method, combinatorial binders-enabled selection of CID (COMBINES-CID), for the de novo engineering of CID systems applicable to a large variety of ligands. This method uses the two-step selection of a phage-displayed combinatorial nanobody library to obtain 1) &#34;anchor binders&#34; that first bind to a ligand of interest and then 2) &#34;dimerization binders&#34; that only bind to anchor binder-ligand complexes. To select anchor binders, a combinatorial library of over 109 complementarity-determining region (CDR)-randomized nanobodies is screened with a biotinylated ligand and hits are validated with the unlabeled ligand by bio-layer interferometry (BLI). To obtain dimerization binders, the nanobody library is screened with anchor binder-ligand complexes as targets for positive screening and the unbound anchor binders for negative screening. COMBINES-CID is broadly applicable to select CID binders with other immunoglobulin, non-immunoglobulin, or computationally designed scaffolds to create biosensors for in vitro and in vivo detection of drugs, metabolites, signaling molecules, etc.

Creating chemically induced protein dimerization systems with desired affinity and specificity for any given small molecule ligand would have many biological sensing and actuation applications. Here, we describe an efficient, generalizable method for de novo engineering of chemically induced dimerization systems via the stepwise selection of a phage-displayed combinatorial single-domain antibody library.

Protein dimerization events that occur only in the presence of a small-molecule ligand enable the development of small-molecule biosensors for the ...

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 ...