The authors acknowledge funding from a European Research Council advanced grant to Hermann Gaub. Michael A. Nash gratefully acknowledges funding from Society in Science - The Branco Weiss Fellowship program. The authors thank Edward A. Bayer, Yoav Barak, and Daniel B. Fried at the Weizmann Institute of Science for generously providing the proteins used in this study. The authors thank Hermann E. Gaub, Elias M. Puchner, and Stefan W. Stahl for helpful discussions.

A schematic of the pulling geometry used in this work to probe the cohesin-dockerin interaction is shown in Figure 1A. The protein immobilization protocol reported here for cantilever and cover glass functionalization is a modified version of the procedure published previously27. The proteins were expressed from plasmid vectors in E. coli using conventional methods. The proteins were designed with a solvent-accessible thiol group, which was used in combination with maleimide chemistry to tether the protein via a stable thioether linkage to the cover glass surface and cantilever. The engineered cysteine residues in both the CBM-cohesin and xylanase-dockerin fusion proteins were located towards the N-terminal side of the proteins, away from the cohesin-dockerin binding interface11. A detailed overview of the chemical bonding employed in protein immobilization is shown in Figure 1B.
1. Sample Preparation
<ol>
	<li>Buffers
	<ol>
		<li>Prepare Tris buffered saline supplemented with calcium (TBS): 25 mM TRIS, 75 mM NaCl, 1 mM CaCl2, pH 7.2</li>
		<li>Prepare sodium borate buffer: 50 mM Na2B4O7, pH 8.5 
		The process flow diagram showing sample preparation steps is shown in Figure 2. 
		When handling cantilevers and cover glasses, self-locking tweezers are recommended.</li>
	</ol>
	</li>
	<li>Aminosilanization of cover glass (approximately&#160;1.5 hr)
	<ol>
		<li>Place cover glass (24 mm diameter, 0.5 mm thickness) in a PTFE holder.</li>
		<li>Sonicate cover glass in 1:1 ethanol : ultrapure water (v/v) for 15 min.</li>
		<li>Rinse cover glass with ultrapure water.</li>
		<li>Place cover glass in piranha solution (1:1 H2SO4 (concentrated) : H2O2 (30%) (v/v)) for 30 min, then thoroughly rinse with ultrapure water. Dry cover glass under a gentle stream of N2. Caution: piranha solution is extremely corrosive. Eye and skin protection are required.</li>
		<li>Submerge cover glass in 45:5:1 ethanol : ultrapure water : 3-aminopropyl dimethyl ethoxysilane (v/v). Place on a shaker at RT for 60 min (approximately&#160;50 rpm).</li>
		<li>Submerge cover glass sequentially in ethanol and ultrapure water (2x each). Dry cover glass under a gentle stream of N2.</li>
		<li>Bake cover glass in an oven (80 &#176;C for 30 min).</li>
		<li>Silanized cover glasses may be stored under argon for up to 6 weeks.</li>
	</ol>
	</li>
	<li>Aminosilanization of cantilevers (approximately&#160;1.5 hr) 
	NOTE: The presented protocol for tip functionalization is appropriate for silicon cantilever tips.
	<ol>
		<li>Place cantilevers on a clean glass slide. Treat with UV-ozone for 15 min.</li>
		<li>Submerge cantilevers for 3 min&#160;in 1:1 ethanol : 3-aminopropyl dimethyl ethoxysilane (v/v) with a catalytic amount (0.25%, (v/v)) of ultrapure water.</li>
		<li>Rinse cantilevers with gentle stirring sequentially for 60 sec&#160;in beakers of toluene, ethanol, and ultrapure water. Carefully dry cantilevers on filter paper between rinses.</li>
		<li>Place levers on a clean glass slide and bake (80 &#176;C for 30 min).</li>
	</ol>
	</li>
	<li>Protein disulfide reduction (approximately&#160;3 hr) 
	All solutions should be prepared to obtain approximately 30 &#181;l&#160;of diluted protein per cantilever and 20 &#181;l&#160;of diluted protein per cover glass. Protein solutions should be mixed with Tris(2-carboxyethyl)phosphine (TCEP) disulfide reducing gel in a ratio of 1:2 (v/v).
	<ol>
		<li>Prepare aliquots of TCEP disulfide reducing beads in micro-tubes. It is recommended to cut micropipette tips with scissors to widen the hole diameter when pipetting the TCEP bead slurry.</li>
		<li>Rinse TCEP bead slurry with 1 ml&#160;TBS buffer, and centrifuge at 850 rcf for 3 min.</li>
		<li>Carefully remove and discard the supernatant with a micropipette.</li>
		<li>Repeat steps 1.4.2-1.4.3&#160;2x.</li>
		<li>Apply concentrated protein solution (1-10 mg/ml) to the TCEP beads (1:2 protein : TCEP bead slurry (v/v)) and gently mix by stirring with micropipette tip. Avoid introducing air bubbles.</li>
		<li>Place protein/TCEP bead slurry mixture on a rotator for 2.5 hr.</li>
	</ol>
	</li>
	<li>PEGylation of cover glasses and cantilevers (approximately&#160;1.5 hr)
	<ol>
		<li>Prior to modification with NHS-PEG-maleimide linkers, soak aminosilanized cantilevers and cover glasses in sodium borate buffer (pH 8.5) for 45 min to deprotonate primary amine groups on the surface.</li>
		<li>Ensure that the NHS-PEG-maleimide powder is warmed up to RT&#160;before opening the cap and weighing the appropriate amount for a 25 mM solution. Unused NHS-PEG-maleimide should be stored under argon at -20 &#176;C. Approximately 30 &#181;l&#160;of polymer solution per cantilever, and 90 &#181;l&#160;per 2 cover glasses (sandwiched together) is required.</li>
		<li>After weighing the 5 kDa NHS-PEG-maleimide, add sodium borate buffer and vortex to obtain a 25 mM solution. 
		Note: The solution should be used as quickly as possible due to the extremely short half-life of NHS at pH 8.5. Vortexing and transfer of the liquid onto the cantilevers/cover glasses should be completed within 1-2 min.</li>
		<li>Incubate cantilevers in 30 &#181;l&#160;droplets of NHS-PEG-maleimide solution in a Petri dish. For cover glasses, place 90 &#181;l&#160;of NHS-PEG-maleimide solution onto a single cover glass, and add a second cover glass on top creating a cover glass sandwich with NHS-PEG-maleimide solution in the middle.</li>
		<li>Incubate the cantilevers/cover glasses with the NHS-PEG-maleimide solution in a water-saturated atmosphere at RT&#160;for 1 hr.</li>
	</ol>
	</li>
	<li>Protein conjugation (approximately&#160;2 hr) 
	Critical: Minimize the exposure of PEGylated cantilevers and cover glasses to air.
	<ol>
		<li>Centrifuge TCEP-bead/ reduced protein solutions at 100 rcf for 1 min and collect the supernatant.</li>
		<li>Dilute protein solution with TBS. Aim for a protein concentration during surface conjugation in a range of 0.5-2 mg/ml. Set reduced and diluted protein solutions aside for a few minutes while rinsing cantilevers and cover glasses.</li>
		<li>Rinse cantilevers and cover glasses in three sequential beakers of ultrapure water.</li>
		<li>Carefully remove residual liquid from cover glasses by carefully touching the edges to a filter paper under a gentle stream of N2. Carefully remove residual liquid from cantilevers by touching to a filter paper. Apply diluted protein solution immediately.</li>
		<li>Mount cover glasses in an appropriate sample holder that is compatible with the AFM instrument.</li>
		<li>Incubate PEGylated cover glasses and cantilevers with respective diluted protein solutions at RT for 1-2 hr.</li>
		<li>Rinse cantilevers in three sequential beakers with TBS to remove unbound proteins. Pipette rinse cover glasses at least 10x.</li>
		<li>Store cantilevers and cover glasses under TBS prior to measurement.</li>
	</ol>
	</li>
</ol>
2. Data Acquisition
In this work, a custom-built AFM28 controlled by an MFP-3D AFM controller from Asylum Research with custom written Igor Pro software was used. Cantilever deflection is measured via the optical beam deflection method29. The sample preparation and data analysis protocols provided here are applicable regardless of the exact AFM model used. However, the AFM model should be suitable for measuring in liquids and support an accessible speed range on the z-piezo of approximately 200-5,000 nm/sec.
<ol>
	<li>Mount the functionalized cantilever and glass surface on the AFM. During the whole procedure, the surface should stay covered with buffer. When mounting the cantilever, minimize exposure to air. Upon correct adjustment of the laser beam, let the system equilibrate for at least 30 min to reduce any drift effects and readjust if necessary.</li>
	<li>Record a thermal noise spectrum with the cantilever far away from the surface, i.e. in the absence of damping effects.</li>
	<li>Use a minimally invasive method like the acoustic approach to find the surface without damaging the cantilever tip prior to measurement. If possible, manually approach the surface with the cantilever and use headphones to listen to the thermal noise on the raw deflection output from the AFM controller. As soon as the cantilever draws near the surface, a distinct change in sound is audible. 
	Note: The cantilever tip should now be within 2-5 &#181;m of the surface. The nature of the sound change is dependent on the cantilever used. The resonance frequency of the cantilever used in this work is approximately 25 kHz in water, above the human audible range. Due to damping effects near the surface, the resonance is shifted towards lower frequencies bringing the cantilever resonance into the audible range. Hence, an apparent increase in frequency and sound intensity is perceived. 
	In cases where an audio output of the deflection signal is not available, the surface can be approached with the z-piezo while an active feedback on the deflection signal is enabled. As soon as the deflection signal increases by a defined amount due to indentation of the surface, the approach is stopped.</li>
	<li>Determine the inverse optical lever sensitivity, (InvOLS) which represents the tip displacement distance (in nm) per volt deflection signal. Do this by indenting the surface with the cantilever tip. A deflection set point voltage corresponding to a cantilever tip displacement of approximately 3 nm is recommended.</li>
	<li>Determine the spring constant of the cantilever by fitting a simple harmonic oscillator response function to the thermal noise spectrum, according to the equipartition theorem30,31.</li>
	<li>Initialize an experimental routine. For this work, the following set of measurement parameters was used: approach speed: 3,000 nm/sec; indentation force: 180 pN; surface dwell time: 10 msec; retract velocities: 0.2, 0.7, 2.0, 5.0 &#181;m/sec with sampling rates of 2,000, 5,000, 15,000, 20,000 Hz respectively; retract distance: 500 nm. 
	Note: The sampling rate should not be set higher than 10 points/nm to avoid oversampling and to keep data sizes reasonable.</li>
	<li>After each force-distance trace, actuate the x- and y-piezo stages to expose a new surface location to the cantilever in each force-distance curve. This technique samples a larger area of the cover glass surface during long-term measurements.</li>
	<li>Use periodic rezeroing of the deflection stage (i.e.&#160;photodiode position) and height of the z-piezo chassis during long-term measurements in case the deflection signal drifts out of range, or contact is lost with the surface.</li>
	<li>Upon completion of the measurement run, perform another InvOLS measurement with a significantly higher indentation force than used prior to measurements to obtain a more precise InvOLS value.</li>
	<li>Record another thermal noise spectrum far away from the surface. Determine the spring constant at the end of the experimental run.</li>
</ol>
3. Data Analysis
The flow diagram in Figure 3 illustrates the process of data analysis. Perform all data manipulations using an appropriate software package such as Igor Pro or MATLAB. First convert the raw signal from the detector into units of force, and correct for offset and drift. Subsequently, use models of biopolymer elasticity to locate energy barriers in the unfolding pathways, and identify protein subdomains. Finally, kinetic and energetic parameters of the receptor-ligand interaction are obtained.
<ol>
	<li>Unit conversion and data corrections
	<ol>
		<li>Multiply the raw deflection signal (volts) by the InvOLS (nm/volt) and spring constant (pN/nm) to convert the detector voltage into units of force.</li>
		<li>Offset the data such that the unloaded cantilever has a force value of zero pN by first averaging the force values from the last 10% of the force-distance trace (acquired farthest away from the surface), and then subtracting the average from all force values in the data trace.</li>
		<li>Offset each trace in the x-direction such that the first intercept with the distance axis occurs at a distance of 0 nm.</li>
		<li>The InvOLS is dependent on the laser spot position on the cantilever. Even small amounts of drift in the optical readout system may cause noticeable changes in the InvOLS when the footprint of the cantilever is comparable to the laser spot size. Correct for this by analysis of the noise on the deflection signal at zero force. Assuming constant ambient conditions, noise on the deflection signal is directly proportional to the InvOLS.
		<ol>
			<li>Measure the route mean square (RMS) deflection value (noise level under zero force) of the last 10% of each force-distance trace.</li>
			<li>Plot the noise vs. the curve number and apply a suitable fit. Typically an exponential fit in the form of <img alt="" fo:content-width="1in" height="22" src="/files/ftp_upload/50950/50950_clip_image002.jpg" width="109" />&#160;will work best, where N is the noise, ni is the curve number, and N0 and k are fit parameters. A linear fit may also be appropriate for certain data sets.</li>
			<li>Determine a scaling factor (SF) for each curve: 
			 
			Equation 1: 
			<img alt="" fo:content-width="2in" height="50" src="/files/ftp_upload/50950/50950eq1.jpg" /> 
			where, ni is the curve number, nf is the final curve number, and C is an offset.</li>
			<li>Next divide all the force values in each individual curve by the scaling factor. This procedure scales each curve by the ratio of the RMS noise value of the current curve to the RMS noise value of the final trace that was acquired immediately prior to the InvOLS measurement.</li>
			<li>Perform a deflection correction to transform the distance axis (z) to molecular extension (z*). This accounts for bending of the cantilever under force which shortens the distance between cantilever tip and sample from the value reported by the z-piezo sensor position. 
			 
			Equation 2: 
			<img alt="" fo:content-width="1in" height="50" src="/files/ftp_upload/50950/50950eq2.jpg" /> 
			Where z is the measured z-sensor position, F the force acting on the cantilever and k the spring constant.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Contour length analysis 
	The contour length of a protein is the maximum stretched length of the polypeptide chain. The folding state of a protein refers to its geometry and end to end distance determined by secondary and tertiary structure. The contour length of a protein is directly related to its folding state9,25,32. The position of specific ruptures in force-extension traces varies widely due to polydispersity of PEG linkers, as well as external parameters such as temperature, buffer properties and loading rates. This complicates direct data analysis but can be overcome by transforming force-extension data into contour length space. This technique enables averaging over huge datasets, and allows automatic pattern recognition to be used to identify characteristic unfolding events. It is therefore possible to sort individual force traces depending on the type of interaction exhibited. The following previously described procedure25 is used to transform force-extension data into contour length space.
	<ol>
		<li>Solving the WLC model (Equation 3)23 for the contour length L at a fixed persistence length p results in Equation 4, which provides&#160; the contour length L(x,u). Here, x is the distance and u=F*p/kBT, where kB is Boltzmann&#39;s constant and T is the temperature. Only real solutions can be considered. Additional constraints are x&#60;L, F&#62;0, L&#62;0, x&#62;0; 
		 
		Equation 3: 
		<img alt="" fo:content-width="2.5in" height="50" src="/files/ftp_upload/50950/50950eq3.jpg" /> 
		 
		Equation 4: 
		<img alt="" fo:content-width="3in" height="50" src="/files/ftp_upload/50950/50950eq4a.jpg" /> 
		where, 
		<img alt="" fo:content-width="3.5in" height="60" src="/files/ftp_upload/50950/50950eq4b.jpg" /></li>
		<li>Plot the transformed data points in a force vs. contour length plot. Apply a force threshold of approximately 10 pN to exclude noise. Unspecific interactions can be excluded by applying a long-pass length filter. Assemble a histogram of contour lengths.</li>
		<li>Cross-correlate33 the obtained histograms with a template histogram, and offset along the x-axis to correct for PEG polydispersity. Use the resulting correlation values to measure the similarity of individual data traces. Thereby, data traces can be sorted into predefined classes to simplify further analysis.</li>
		<li>Use a similar technique to find repeating features in a single trace by autocorrelation, e.g. for multiple Ig-domain unfolding.</li>
		<li>Sort traces manually to investigate other unfolding events.</li>
	</ol>
	</li>
	<li>Loading Rate Analysis 
	Extract kinetic and energetic information about receptor-ligand dissociation by applying suitable models to the force spectrum, i.e. the rupture-force vs. ln(loading-rate) plot.
	<ol>
		<li>For a given pulling speed, determine the rupture force and loading rate for rupture events of interest:
		<ol>
			<li>Perform a line fit to a force-time trace in the vicinity of the rupture event of interest. Determine the loading rate from the slope of the line fit to the peak. Repeat this procedure for every trace showing the rupture event of interest.</li>
			<li>Determine the most probable rupture force by applying a Gaussian fit to a histogram of the rupture forces. Alternative fit functions are possible.</li>
			<li>Determine the most probable loading rate.</li>
		</ol>
		</li>
		<li>Repeat steps 3.3.3.1 - 3.3.3.3 for all pulling speeds.</li>
		<li>Plot the most probable rupture forces against the natural logarithm of the most probable loading rates to obtain the force spectrum.</li>
		<li>Apply a suitable theoretical model to the force spectrum to extract kinetic and energetic parameters (Figure 4C). In many cases, the linear Bell-Evans model20,34 can be used and will yield good estimates for koff, the dissociation rate in the absence of force, and Dx, the distance to the transition state along the reaction coordinate, as shown in Equation 5. 
		 
		Equation 5: 
		<img alt="" fo:content-width="2in" height="50" src="/files/ftp_upload/50950/50950eq5.jpg" /></li>
	</ol>
	</li>
</ol>

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J. Mol. Biol. 381 (5), 1253-1266 (2008).</li><li>Ortiz, C., Hadziioannou, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Entropic+Elasticity+of+Single+Polymer+Chains+of+Poly(methacrylic+acid)+Measured+by+Atomic+Force+Microscopy.">Entropic Elasticity of Single Polymer Chains of Poly(methacrylic acid) Measured by Atomic Force Microscopy.</a> Macromolecules. 32 (3), 780-787 (1999).</li><li>Bustamante, C., Marko, J. F., Siggia, E. D., Smith, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Entropic+Elasticity+of+l-phage+DNA.">Entropic Elasticity of l-phage DNA.</a> Science. 265 (5178), 1599-1600 (1994).</li><li>Marko, J. F., Siggia, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stretching+DNA.">Stretching DNA.</a> Macromolecules. 28 (26), 8759-8770 (1995).</li><li>Puchner, E. M., Franzen, G., Gautel, 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=Comparing+Proteins+by+Their+Unfolding+Pattern.">Comparing Proteins by Their Unfolding Pattern.</a> Biophys. J. 95 (1), 426-434 (2008).</li><li>Livadaru, L., Netz, R. R., Kreuzer, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stretching+Response+of+Discrete+Semiflexible+Polymers.">Stretching Response of Discrete Semiflexible Polymers.</a> Macromolecules. 36 (10), 3732-3744 (2003).</li><li>Zimmermann, J. L., Nicolaus, T., Neuert, G., Blank, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thiol-based,+site-specific+and+covalent+immobilization+of+biomolecules+for+single-molecule+experiments.">Thiol-based, site-specific and covalent immobilization of biomolecules for single-molecule experiments.</a> Nat. Protoc. 5 (6), 975-985 (2010).</li><li>Gumpp, H., Stahl, S. W., Strackharn, M., Puchner, E. 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=Ultrastable+combined+atomic+force+and+total+internal+fluorescence+microscope.">Ultrastable combined atomic force and total internal fluorescence microscope.</a> Rev. Sci. Instrum. 80 (6), 063704 (2009).</li><li>Gustafsson, M. G. 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=Scanning+force+microscope+springs+optimized+for+optical-beam+deflection+and+with+tips+made+by+controlled+fracture.">Scanning force microscope springs optimized for optical-beam deflection and with tips made by controlled fracture.</a> J. Appl. Phys. 76 (1), 172 (1994).</li><li>Cook, S. M., Lang, K. M., Chynoweth, K. M., Wigton, M., Simmonds, R. W., Sch&#228;ffer, T. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Practical+implementation+of+dynamic+methods+for+measuring+atomic+force+microscope+cantilever+spring+constants.">Practical implementation of dynamic methods for measuring atomic force microscope cantilever spring constants.</a> Nanotechnology. 17 (9), 2135-2145 (2006).</li><li>Proksch, R., Sch&#228;ffer, T. E., Cleveland, J. P., Callahan, R. C., Viani, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Finite+optical+spot+size+and+position+corrections+in+thermal+spring+constant+calibration.">Finite optical spot size and position corrections in thermal spring constant calibration.</a> Nanotechnology. 15 (9), 1344-1350 (2004).</li><li>Ainavarapu, S. R. K., Brujic, 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=Contour+Length+and+Refolding+Rate+of+a+Small+Protein+Controlled+by+Engineered+Disulfide+Bonds.">Contour Length and Refolding Rate of a Small Protein Controlled by Engineered Disulfide Bonds.</a> Biophys. J. 92 (1), 225-233 (2007).</li><li>Dietz, H., Rief, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detecting+Molecular+Fingerprints+in+Single+Molecule+Force+Spectroscopy+Using+Pattern+Recognition.">Detecting Molecular Fingerprints in Single Molecule Force Spectroscopy Using Pattern Recognition.</a> Jap. J. Appl. Phys. 46, 5540-5542 (2007).</li><li>Evans, E., Ritchie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+strength+of+molecular+adhesion+bonds.">Dynamic strength of molecular adhesion bonds.</a> Biophys. J. 72 (4), 1541-1555 (1997).</li><li>Dietz, H., Rief, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+structure+by+mechanical+triangulation.">Protein structure by mechanical triangulation.</a> Proc. Natl. Acad. Sci. U.S.A. 103 (5), 1244-1247 (2006).</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=Intrinsic+Rates+and+Activation+Free+Energies+from+Single-Molecule+Pulling+Experiments.">Intrinsic Rates and Activation Free Energies from Single-Molecule Pulling Experiments.</a> Phys. Rev. Lett. 96 (10), 108101 (2006).</li><li>Friddle, R. W., Noy, A., De Yoreo, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interpreting+the+widespread+nonlinear+force+spectra+of+intermolecular+bonds.">Interpreting the widespread nonlinear force spectra of intermolecular bonds.</a> Proc. Natl. Acad. Sci. U.S.A. 109 (34), 13573-13578 (2012).</li></ol>

The authors declare that they have no competing financial interests.

To obtain meaningful data from single molecule force spectroscopy experiments, it is crucial to achieve well-defined and reproducible pulling geometries. The protocol used here results in site-specific immobilization of protein complexes in a defined pulling geometry.
The cantilevers used in this study were chosen due to their force sensitivity and high resonance frequency in water. Moreover, the small tip curvature of approximately 10 nm is advantageous for single molecule experiments due to ...

Cellulosomes are large multienzyme complexes displayed on the surface of anaerobic cellulolytic bacteria (e.g.&#160;C. thermocellum) that have evolved to efficiently depolymerize plant cell wall lignocellulose into soluble oligosaccharides1. A central attribute of cellulosomes is the high-affinity cohesin-dockerin interaction. In the most prominent paradigm, a highly conserved 60-75 amino acid type I dockerin module is displayed at the C-terminal end of the various bacterial enzymes. The dockerin module directs assembly of synergistic combinations of enzymes onto the noncatalytic scaffold protein (&#39;scaffoldin&#39;), which comprises a polyprotein of cohesin domains that are specific for the type I dockerin module. At higher levels, cellulosome architecture can become very complex, incorporating alternative cohesin and dockerin pairs (e.g.&#160;type II, type III) that anchor the structures to the cell surface and allow for the assembly of branched structures containing multiple scaffoldins2. The various cohesin-dockerin types, despite having related structures, exhibit differential binding specificities suppressing cross reactivity with unintended scaffoldins or components from other cellulosome-producing bacterial species. While bioinformatic approaches have successfully identified thousands of unique cellulosomal components at the genetic level, comparatively few protein structures are known, and the mechanisms at work in cohesin-dockerin specificity determination remains an active area of structural biology research.
Since the invention of the atomic force microscope (AFM) by Binnig et al.3, various AFM operational modes have been developed and continuously improved, including noncontact imaging, oscillation mode imaging4, and single molecule force spectroscopy (SMFS)5,6. SMFS has evolved into a widely used technique to directly probe individual proteins7-11, nucleic acids12-15, and synthetic polymers16-19. In a typical SMFS experiment to investigate receptor-ligand binding20,21, an AFM cantilever tip is modified with one of the binding partners, while a flat glass surface is modified with the complementary binding partner. The modified cantilever is brought into contact with the surface allowing the partners to bind. The base of the cantilever is then withdrawn at constant speed and the force is measured using the optical lever deflection method. The resultant force-distance data traces exhibit sawtooth-like peaks if binding was established. In cases where the binding partners are fused to multiple protein domains, each peak in the force-distance trace can be correlated to the unfolding of a single protein domain or folded subdomain, while the last peak corresponds to rupture of the protein binding interface. The specific positions of the force-resistant elements can be used as a fingerprint to identify the various protein domains of interest. This method can be used to interrogate important amino acids involved in protein folding and stabilization. Many models have been reported in the literature to treat the characteristic force extension behavior observed in SMFS experiments. The most commonly used models include the freely jointed chain (FJC) model22, the worm-like chain (WLC) model18,23-25, and the freely rotating chain (FRC) model25,26.
In our prior work11, we used single-molecule force spectroscopy to investigate the interaction of cohesin and dockerin modules. Here, we present an experimental protocol for glass surface and cantilever functionalization with enzyme-dockerin and CBM-cohesin protein constructs. We also present an AFM-based SMFS protocol including data acquisition and analysis procedures. The described protocol can easily be generalized to other molecular systems, and should prove particularly useful to researchers interested in high-affinity receptor ligand pairs.

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			<td height="42" style="height:42px;width:216px;">3-Aminopropyl dimethyl ethoxysilane</td>
			<td style="width:139px;">ABCR GmbH</td>
			<td style="width:119px;">AB110423</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">5 kDa NHS-PEG-maleimide</td>
			<td style="width:139px;">Rapp Polymer</td>
			<td style="width:119px;">13 5000-65-35</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">TCEP Disulfide reducing gel</td>
			<td style="width:139px;">Thermo Scientific, Pierce</td>
			<td align="right" style="width:119px;">77712</td>
			<td><a href="http://www.thermoscientific.com/pierce">www.thermoscientific.com/pierce</a></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Tris(hydroxymethyl)aminomethane</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">BioLever mini silicon nitride cantilevers</td>
			<td style="width:139px;">Olympus</td>
			<td style="width:119px;">BL-AC40TS-C2</td>
			<td>Soft batches</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">XYZ Piezoelectric actuators</td>
			<td style="width:139px;">Physik Instrumente GmbH</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Infrared &#8220;broad spectrum&#8221; IR laser</td>
			<td style="width:139px;">Superlum</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">MFP-3D AFM Controller</td>
			<td style="width:139px;">Asylum Research</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Igor Pro 6.31</td>
			<td style="width:139px;">Wavemetrics</td>
			<td style="width:119px;"></td>
			<td>Data acquisition and analysis</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium chloride</td>
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			<td height="21" style="height:21px;width:216px;">Calcium chloride</td>
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			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">pH Meter</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium borate</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tweezers</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Cover glasses</td>
			<td style="width:139px;">Thermo Scientific, Menzel-Gl&#228;ser</td>
			<td style="width:119px;"></td>
			<td>24 mm diameter, 0.5 mm thickness</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PTFE sample holder</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>custom made</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sonicator bath</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ethanol</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>analytical purity</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sulfuric acid (concentrated)</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>analytical purity</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Hydrogen peroxide (30%)</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>analytical purity</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Orbital shaker</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
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			<td height="21" style="height:21px;width:216px;">Toluene</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>analytical purity</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Filter paper</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Glass slides</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Microtubes</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micropipettes</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Centrifuge</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>suitable for microtubes</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Rotator</td>
			<td style="width:139px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Petri dishes</td>
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			<td style="width:119px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Beakers</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Optical microscope</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

investigating receptor-ligand systems of the cellulosome with afm-based single-molecule force spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

We used the described procedure to investigate a type I cohesin-dockerin pair from C. thermocellum. Upon successful binding of the cohesin-dockerin pair, the recorded force distance traces showed characteristic peak patterns. A typical trace is shown in Figure 4a. Every peak in the trace represents the unfolding of one protein subdomain with the last peak corresponding to the dissociation of the receptor-ligand complex.
For the CBM-cohesin-dockerin-xylanase complex in...

Watch this Scientific Journal Video about Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy at JoVE.com

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

investigating-receptor-ligand-systems-cellulosome-with-afm-based

Research

JoVE Journal

Bioengineering

7.2K Views.  Ludwig-Maximilians-Universität. Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Video: Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

The generation and detection of mechanical forces is a ubiquitous aspect of cell physiology, with direct relevance to cancer metastasis1, atherogenesis2 and wound healing3. In each of these examples, cells both exert force on their surroundings and simultaneously enzymatically remodel the extracellular matrix (ECM). The effect of forces on ECM has thus become an area of considerable interest due to its likely biological and medical importance4-7.
	Single molecule techniques such as optical trapping8, atomic force microscopy9, and magnetic tweezers10,11 allow researchers to probe the function of enzymes at a molecular level by exerting forces on individual proteins. Of these techniques, magnetic tweezers (MT) are notable for their low cost and high throughput. MT exert forces in the range of ~1-100 pN and can provide millisecond temporal resolution, qualities that are well matched to the study of enzyme mechanism at the single-molecule level12. Here we report a highly parallelizable MT assay to study the effect of force on the proteolysis of single protein molecules. We present the specific example of the proteolysis of a trimeric collagen peptide by matrix metalloproteinase 1 (MMP-1); however, this assay can be easily adapted to study other substrates and proteases.

In this article we describe the use of magnetic tweezers to study the effect of force on enzymatic proteolysis at the single molecule level in a highly parallelizable manner.

The  generation and detection of mechanical forces is a ubiquitous aspect of cell physiology,  with direct relevance to cancer metastasis1, atherogenesis2 ...

Quantifying the Mechanical Properties of the Endothelial Glycocalyx with Atomic Force Microscopy

Our understanding of the interaction of leukocytes and the vessel wall during leukocyte capture is limited by an incomplete understanding of the mechanical properties of the endothelial surface layer. It is known that adhesion molecules on leukocytes are distributed non-uniformly relative to surface topography 3, that topography limits adhesive bond formation with other surfaces 9, and that physiological contact forces (&asymp; 5.0 &minus; 10.0 pN per microvillus) can compress the microvilli to as little as a third of their resting length, increasing the accessibility of molecules to the opposing surface 3, 7. We consider the endothelium as a two-layered structure, the relatively rigid cell body, plus the glycocalyx, a soft protective sugar coating on the luminal surface 6. It has been shown that the glycocalyx can act as a barrier to reduce adhesion of leukocytes to the endothelial surface 4. In this report we begin to address the deformability of endothelial surfaces to understand how the endothelial mechanical stiffness might affect bond formation. Endothelial cells grown in static culture do not express a robust glycocalyx, but cells grown under physiological flow conditions begin to approximate the glycocalyx observed in vivo 2. The modulus of the endothelial cell body has been measured using atomic force microscopy (AFM) to be approximately 5 to 20 kPa 5. The thickness and structure of the glycocalyx have been studied using electron microscopy 8, and the modulus of the glycocalyx has been approximated using indirect methods, but to our knowledge, there have been no published reports of a direct measurement of the glycocalyx modulus in living cells. In this study, we present indentation experiments made with a novel AFM probe on cells that have been cultured in conditions to maximize their glycocalyx expression to make direct measurements of the modulus and thickness of the endothelial glycocalyx.

The mechanical characteristics of endothelial glycocalyx were measured by indentation using micron sized spheres on AFM cantilevers. Endothelial cells were cultured in a custom chamber under physiological flow conditions to induce glycocalyx expression. Data were analyzed using a thin film model to determine the glycocalyx thickness and modulus.

Our  understanding of the interaction of leukocytes and the vessel wall during  leukocyte capture is limited by an incomplete understanding of the ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions are likely affected by the mechanical environment in which binding and signaling take place. A recent study demonstrated that mechanical force can regulate antigen recognition by and triggering of the T-cell receptor (TCR). This was made possible by a new technology we developed and termed fluorescence biomembrane force probe (fBFP), which combines single-molecule force spectroscopy with fluorescence microscopy. Using an ultra-soft human red blood cell as the sensitive force sensor, a high-speed camera and real-time imaging tracking techniques, the fBFP is of ~1 pN (10-12 N), ~3 nm and ~0.5 msec in force, spatial and temporal resolution. With the fBFP, one can precisely measure single receptor-ligand binding kinetics under force regulation and simultaneously image binding-triggered intracellular calcium signaling on a single live cell. This new technology can be used to study other membrane receptor-ligand interaction and signaling in other cells under mechanical regulation.

We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

Label-free Single Molecule Detection Using Microtoroid Optical Resonators

Detecting small concentrations of molecules down to the single molecule limit has impact on areas such as early detection of disease, and fundamental studies on the behavior of molecules. Single molecule detection techniques commonly utilize labels such as fluorescent tags or quantum dots, however, labels are not always available, increase cost and complexity, and can perturb the events being studied. Optical resonators have emerged as a promising means to detect single molecules without the use of labels. Currently the smallest particle detected by a non-plasmonically-enhanced bare optical resonator system in solution is a 25 nm polystyrene sphere1. We have developed a technique known as Frequency Locking Optical Whispering Evanescent Resonator (FLOWER) that can surpass this limit and achieve label-free single molecule detection in aqueous solution2. As signal strength scales with particle volume, our work represents a &#62; 100x improvement in the signal to noise ratio (SNR) over the current state of the art. Here the procedures behind FLOWER are presented in an effort to increase its usage in the field.

We have developed a label-free biosensing system based on optical resonator technology known as Frequency Locking Optical Whispering Evanescent Resonator (FLOWER) that is capable of detecting single molecules in solution. Here the procedures behind this work are described and presented.

Detecting small concentrations of molecules down to the single molecule limit has impact on areas such as early detection of disease, and fundamental ...

A Guide to Build a Highly Inclined Swept Tile Microscope for Extended Field-of-view Single-molecule Imaging

Single-molecule imaging has greatly advanced our understanding of molecular mechanisms in biological studies. However, it has been challenging to obtain large field-of-view, high-contrast images in thick cells and tissues. Here, we introduce highly inclined swept tile (HIST) microscopy that overcomes this problem. A pair of cylindrical lenses was implemented to generate an elongated excitation beam that was scanned over a large imaging area via a fast galvo mirror. A 4f configuration was used to position optical components. A scientific complementary metal-oxide semiconductor camera detected the fluorescence signal and blocked the out-of-focus background with a dynamic confocal slit synchronized with the beam sweeping. We present a step-by-step instruction on building the HIST microscope with all basic components.

A detailed instruction is described on how to build a highly inclined swept tile (HIST) microscope and its usage for single-molecule imaging.

Single-molecule imaging has greatly advanced our understanding of molecular mechanisms in biological studies. However, it has been challenging to obtain ...

Characterizing Single-Molecule Conformational Changes Under Shear Flow with Fluorescence Microscopy

Single-molecule behavior under mechanical perturbation has been characterized widely to understand many biological processes. However, methods such as atomic force microscopy have limited temporal resolution, while F&#246;rster resonance energy transfer (FRET) only allow conformations to be inferred. Fluorescence microscopy, on the other hand, allows real-time in situ visualization of single molecules in various flow conditions. Our protocol describes the steps to capture conformational changes of single biomolecules under different shear flow environments using fluorescence microscopy. The shear flow is created inside microfluidic channels and controlled by a syringe pump. As demonstrations of the method, von Willebrand factor (VWF) and lambda DNA are labeled with biotin and fluorophore and then immobilized on the channel surface. Their conformations are continuously monitored under variable shear flow using total internal reflection (TIRF) and confocal fluorescence microscopy. The reversible unraveling dynamics of VWF are useful for understanding how its function is regulated in human blood, while the conformation of lambda DNA offers insights into the biophysics of macromolecules. The protocol can also be widely applied to study the behavior of polymers, especially biopolymers, in varying flow conditions and to investigate the rheology of complex fluids.

We present a protocol for immobilizing single macromolecules in microfluidic devices and quantifying changes in their conformations under shear flow. This protocol is useful for characterizing the biomechanical and functional properties of biomolecules such as proteins and DNA in a flow environment.

Single-molecule behavior under mechanical perturbation has been characterized widely to understand many biological processes. However, methods such as ...

Fabrication of Zero Mode Waveguides for High Concentration Single Molecule Microscopy

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to nanomolar ranges, several orders of magnitude less than many physiological ligand concentrations. Optical nanostructures called zero mode waveguides (ZMWs), which are 100&#8722;200 nm in diameter apertures fabricated in a thin conducting metal such as aluminum or gold, allow imaging of individual molecules at micromolar concentrations of fluorophores by confining visible light excitation to zeptoliter effective volumes. However, the need for expensive and specialized nanofabrication equipment has precluded the widespread use of ZMWs. Typically, nanostructures such as ZMWs are obtained by direct writing using electron beam lithography, which is sequential and slow. Here, colloidal, or nanosphere, lithography is used as an alternative strategy to create nanometer-scale masks for waveguide fabrication. This report describes the approach in detail, with practical considerations for each phase. The method allows thousands of aluminum or gold ZMWs to be made in parallel, with final waveguide diameters and depths of 100&#8722;200 nm. Only common lab equipment and a thermal evaporator for metal deposition are required. By making ZMWs more accessible to the biochemical community, this method can facilitate the study of molecular processes at cellular concentrations and rates.

Described here is a nanosphere lithography method for parallel fabrication of zero mode waveguides, which are arrays of nanoapertures in a metal-clad glass microscopy coverslip for single molecule imaging at nano- to micromolar concentrations of fluorophores. The method takes advantage of colloidal crystal self-assembly to create a waveguide template.

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to ...