This work was supported by the European Union&#8217;s Horizon 2020 research and innovation programme under Grant Agreement No. 871124 Laserlab-Europe, by the Italian Ministry of University and Research (FIRB &#8220;Futuro in Ricerca&#8221; 2013 Grant No. RBFR13V4M2), and by Ente Cassa di Risparmio di Firenze. A.V. Kashchuk was supported by Human Frontier Science Program Cross-Disciplinary Fellowship LT008/2020-C.

1. Optical setup
NOTE: The experimental setup is composed of double optical tweezers with nanometer pointing stability and &#60; 1% laser intensity fluctuations. Under these conditions, stability of the dumbbell at the nanometer level is guaranteed under typical trap stiffness (0.1 pN/nm) and tension (1 pN - few tens of pN). Figure 2 shows a detailed scheme of the optical setup.
<ol>
	<li>Optical tweezers design and construction 9,10,11.
	<ol>
		<li>Place all the components of the setup on an optical table according to the scheme in Figure 2. Note that the optical table includes active isolators to minimize mechanical vibrations. Additionally, the microscope structure is mounted on elastomeric isolators to absorb acoustic noise and mechanical resonances.</li>
		<li>Insert an optical isolator close to the laser source (&#34;OI&#34; in Figure 2), to avoid random amplitude fluctuations due to optical feedback.</li>
		<li>Seal the whole path in a closed box to reduce air current and turbulences that could impact the laser pointing stability.</li>
		<li>Create the double optical tweezers by dividing the main laser source (Nd:YAG laser, 1,064 nm wavelength in Figure 2) into two branches with orthogonal polarizations by the use of polarizing beam splitters (PBS) . Time-shared traps should be avoided because they induce oscillation of the dumbbell under tension12.</li>
		<li>Use two acousto-optic deflectors (AODs in Figure 2) driven by Direct Digital Synthesizers (DDSs) to allow fine and rapid movements of the two traps and precise regulation of the actin tension by directly driving the DDSs through the digital outputs from the field-programmable gate array FPGA board (see Figure 2). 
		&#8203;NOTE: The overall feedback response time must be &#60;10 &#181;s to rapidly correct and maintain constant force on both traps during measurements, including the unbound state, myosin interaction and movement. To this end, position detectors must have a &#62;= 100 kHz bandwidth and data must be acquired at &#62;= 200 kHz sample rate. For each data point acquired (5 &#181;s acquisition time), proportional corrections for the two traps are calculated by the FPGA and sent to the two DDS driving the AODs. The AOD response time must be below 5 &#181;s to satisfy the required feedback response time.</li>
		<li>For nm detection of the trapped beads position, put two Quadrant Photodiode Detectors (QPDs in Figure 2) in the back focal plane of the condenser. Accurate alignment of the QPDs in a plane conjugated to the back focal plane of the condenser, as well as the AODs in a plane conjugated to the back focal plane of the objective, will assure that QPDs signals will be independent of the AODs frequency.</li>
		<li>Mount the AOD on a linear translator with micrometer drive and displace it until the crystal edge gets close to the laser beam. Then, replace the objective with an iris, centered on the threaded objective housing, and regulate its aperture to fit the objective back aperture size.</li>
		<li>Move the translator towards the laser beam until the portion of the beam blocked by the piezo crystal is visible after the iris, turn the translator slightly backwards to get the beam to completely fill the iris aperture again.</li>
		<li>Repeat steps 1.1.7-1.1.8 for the second AOD. Check the response time of the feedback loop by measuring the time lag from the QPDs while rapidly moving a beam on a bead stuck on the surface of the coverslip13. 
		&#8203;NOTE: The above three steps (1.1.7-1-1-9) lead to the careful alignment of AOD crystals. These steps are important to optimize the time response of both the beam deflection and the feedback13.</li>
	</ol>
	</li>
	<li>By means of a photodiode measure the intensity fluctuations of the trapping laser at the microscope entrance which must be below 1%. Note that the photodiode bandwidth must be larger than the imaging rate.</li>
	<li>Check the pointing stability of both traps.
	<ol>
		<li>Prepare silica beads in phosphate buffer (PB) by diluting 20 &#181;L of silica beads (1.2 &#181;m, 10% solids) in 1 mL of acetone, sonicate for 30 s, vortex briefly, and centrifuge for 2 min at 19,000 x g.</li>
		<li>Remove the supernatant, resuspend in 1 mL of acetone, and repeat the wash. Resuspend in 1 mL of 50 mM PB, wash 2 times. Finally, resuspend in 100 &#181;L of 50 mM PB.</li>
		<li>Perform optical tweezers calibration with silica beads by sticking a coverslip onto a microscope slide with a double-sided tape (about 60 &#181;m thick) to build a flow chamber. Fill the chamber with a 1 mg/mL BSA (Bovine Serum Albumin protein) and wait for 3 min.</li>
		<li>Flow a 1:1000 dilution of silica beads in PB into the chamber. Use a syringe filled with silicon grease to carefully seal the chamber. Trap a single bead in each trap and apply the power spectrum method14 to calibrate them.</li>
		<li>Prepare silica beads in pentyl acetate (at room temperature) by dissolving 20 &#181;L silica beads (1.2 &#181;m diameter, 10% solid) in 1-1.5 mL of acetone, vortex and sonicate for 30 s.</li>
		<li>Centrifuge at 18,500 x g for 2 min. Discard the supernatant and resuspend in 1 mL of acetone, then repeat the wash. Resuspend in 1 mL of pentyl acetate and repeat wash (centrifuge and resuspension) in pentyl acetate 2 times. Resuspend the pellet in 100 &#181;L of nitrocellulose 1% and 900 &#181;L pentyl acetate. Store at 4 &#176;C for 2 months.</li>
		<li>Take a 24 x 24 mm glass coverslip and clean it carefully with paper soaked with pure ethanol. Then, while holding it with clean tweezers, rinse it a second time by washing directly with pure ethanol. Let it dry under a gentle flow of nitrogen. If needed, repeat this operation to remove all visible residues on the glass surface.</li>
		<li>Take the silica beads stock, vortex it and sonicate it briefly for ~30 s.</li>
		<li>After cleaning a second coverslip (24 x 60 mm), use it to smear 2 &#181;L of silica beads solution on one surface of the coverslip, and wait for it to dry.</li>
		<li>Clean carefully a microscope slide (26 x 76 mm) that will be used to create the flow chamber.</li>
		<li>Cut two lines of double sticky tape (~3 mm large, 60-100 &#181;m thick) and attach them on one side of the microscope slide, as shown in Figure 3.</li>
		<li>By using clean tweezers, close the chamber (about 20 &#181;L final volume) by putting the coated coverslip (1.3.9) in contact with the sticky tape lines, with the nitrocellulose + beads layer facing the inside of the chamber, as shown in Figure 3a. Fill the flow chamber with 50 mM phosphate buffer and seal it with silicon grease.</li>
		<li>Image a single silica bead in brightfield microscopy at &#62;200x magnification using a charge-coupled device (CCD) or complementary metal-oxide-semiconductor (CMOS) camera with &#62;1.4 megapixels. Use a feedback software to move the piezo stage (with nanometer accuracy or better) to compensate for thermal drifts10.</li>
		<li>Overlap the center of the left trap with the center of the bead (x-y signal levels from the QPD should match those from the calibration). Then measure the position noise and standard deviation of the position signals for this trap.</li>
		<li>Repeat the previous step for the right trap15.</li>
	</ol>
	</li>
	<li>Trap position calibration: MHz to nm
	<ol>
		<li>Prepare a flow chamber with silica beads attached on the coverslip surface (1.3.5-1.3.12) and floating polystyrene beads (use &#945;-actinin conjugated beads prepared as in the following section 2.1).</li>
		<li>Focus a silica bead on the coverslip surface slightly decentered (~5 &#181;m) from the center of the field of view (FOV) and acquire an image of the FOV. Move the bead by 10 &#181;m towards the FOV center using the piezo stage and acquire a second image. Calculate the center of the bead in the two images using a centroid algorithm or similar and calculate the distance in pixel between the two beads to obtain the nm/pixel calibration of the brightfield camera.</li>
		<li>Trap a single floating particle in one trap. Then, move the trap using AOD in small steps (0.2 MHz) and acquire an image of the particle and the corresponding frequency of the AOD for each step. Calculate the position of the particle in the FOV by using the centroid algorithm as before and convert it in nm by using the nm/pixel calibration obtained in the previous step.</li>
		<li>Perform a linear fit to the frequency-position data and calculate the calibration constant in nm/MHz.</li>
		<li>Repeat calibration for the second trap</li>
	</ol>
	</li>
	<li>Trap power and stiffness calibration (MHz vs W), QPD (MHz vs pN/nm)
	<ol>
		<li>Prepare a flow chamber with silica beads attached on the coverslip surface (1.3.5-1.3.12) and floating polystyrene beads (use &#945;-actinin conjugated beads prepared as in the following section 2.1) and trap a single particle in one trap. Then displace both traps through AODs in small steps (0.2 MHz) and record a Brownian motion of the particle in both traps with QPD and the corresponding frequency of the AODs.</li>
		<li>Calculate an average power on the detectors at each position and obtain trap stiffness and QPD calibration constant beta by fitting a Lorentzian function to a power spectrum of the recorded Brownian motion13.</li>
	</ol>
	</li>
</ol>
2. Sample preparation
<ol>
	<li>Prepare &#945;-actinin conjugated fluorescent beads
	<ol>
		<li>Perform conjugation16: Take amino-functionalized polystyrene beads (1 &#181;m diameter, 2.5% solids), wash them twice in 500 &#956;L distilled water and resuspended in 500 &#956;L of PBS (pH 7.0). Add 1 mM HaloTag succinimidyl ester O2 ligand and incubate at room temperature for 1 h. Wash three times with 500 &#956;L of PBS and resuspended with 100-200 &#181;M HaloTag &#945;-actinin. Incubate for 1 h at 37 &#176;C and wash three times in 500 &#956;L of PBS (use beads within 1.5 weeks or flash-frozen in liquid nitrogen and store at &#8722;80 &#176;C).</li>
		<li>Labeling: Incubate 200 &#181;L of beads solution with Rhodamine-BSA at 5 &#181;g/mL final concentration for 10 min. Wash with 50 mM PB three times and resuspend in 500 &#181;L of PB 50 mM. This can be stored in aliquots at - 80 &#176;C for months).</li>
	</ol>
	</li>
	<li>Express and purify biotinylated Myosin-5B as described previously4,17.</li>
	<li>Polymerize and label F-actin13:
	<ol>
		<li>F-actin polymerization: Mix 69 &#181;L of ultrapure water, 10 &#181;L of actin polymerization buffer 10x (100 mM Tris HCl, 20 mM MgCl2, 500 mM KCl, 10 mM ATP, 50 mM guanidine carbonate pH 7.5), 20 &#181;L of G-actin 10 mg/mL, and 1 &#181;L of DL -Dithiothreitol (DTT) 1 M. Leave it on ice for more than 1 h.</li>
		<li>F-actin labeling with rhodamine (Ex/Em: 546/575 nm): Take 25 &#181;L of polymerized F-actin and add 19.5 &#181;L of ultrapure water, 2.5 &#181;L of actin polymerization buffer 10x (100 mM Tris HCl, 20 mM MgCl2, 500 mM KCl, 10 mM ATP, 50 mM guanidine carbonate pH 7.5), 1 &#181;L of 1 M DTT, and 2 &#181;L of 250 &#181;M rhodamine phalloidin. Leave it on ice overnight. For trapping experiments rhodamine F-actin can be stored on ice and used within a week.</li>
	</ol>
	</li>
	<li>Sample assembly
	<ol>
		<li>Take the flow chamber (1.3.5-1.3.12) and incubate 1 mg/mL biotinylated BSA for 5 min. After washing with AB buffer (25 mM MOPS ,25 mM KCl, 4mM MgCl2, 1 mM EGTA, 1 mM DTT, pH 7.2) incubate 1 mg/mL streptavidin for 5 min and wash again with AB buffer. Incubate biotinylated myosin-5B heavy meromyosin at 3 nM concentration in M5B buffer (10 mM MOPS pH 7.3, 0.5 M NaCl, 0.1 mM EGTA, 3 mM NaN3) with 2 &#181;M Calmodulin (CaM) for 5 min. Wash with three volumes of 1 mg/mL biotinylated BSA supplemented with 2 &#181;M CaM in AB and incubate for 3 min.</li>
		<li>While incubating prepare the Reaction Mix (RM): 0.005% &#945;-actinin functionalized beads (section 2.1), 1 nM rhodamine F-Actin (section 2.3) in Imaging Buffer (IB: AB buffer with 1.2 &#181;M Glucose-Oxidase, 0.2 &#181;M catalase, 17 mM glucose, 20 mM DTT, 2 &#181;M CaM and ATP at the concentration needed for the experiment).</li>
		<li>Wash with RM and seal the chamber with silicone grease. The sample is now ready to be watched under the microscope.</li>
	</ol>
	</li>
</ol>
3. Measurement
<ol>
	<li>Assemble the dumbbell.
	<ol>
		<li>Look for the floating &#945;-actinin beads by moving the sample using long-range translators, switch on one trap and trap one bead.</li>
		<li>Once the first trap is occupied, move the translator to position the trapped bead close to the coverslip surface to avoid trapping of multiple beads, and trap another bead in the second trap.</li>
		<li>Adjust the two traps to equal stiffnesses by adjusting the power of the AODs acoustic waves. Stiffnesses are usually set between 0.03 and 0.14 pN/nm. The smaller the stiffness the smaller the force noise, in particular at low forces.</li>
		<li>Then flip the motorized mirror M (Figure 2) to switch to fluorescence microscopy, and look for an actin filament floating in solution by displacing the sample through long-range translators13. Prefer long filaments (&#62;5 &#181;m) since the processive myosin will displace it and move it for a few microns before detaching.</li>
		<li>Move the sample to let one of the trapped beads approaching one end of the filament until they attach to each other. Then, adjust the beads distance to the approximate filament length and create a flow in the direction of the unbound second bead by moving the stage in its direction. The filament will be stretched by the flow and it will eventually bind to it13. The bead-actin-bead complex is called &#34;dumbbell&#34;.</li>
	</ol>
	</li>
	<li>Establish actin-myosin contact.
	<ol>
		<li>Gently separate the two traps far apart to pre-tension the filament up to about 3 pN and probe the rigidity of the dumbbell by making one trap oscillate in a triangular wave by changing the frequency of one of the two AODs and verifying the consequent transmission of the motion to the trailing bead through its position signal.</li>
		<li>Move the stage to put the dumbbell in proximity of a pedestal silica bead and allow the contact between the filament and the protein attached onto bead surface by adjusting the height of the trapped bead centers slightly below the silica bead diameter. Then position the center of the silica bead in between the trapped beads.</li>
	</ol>
	</li>
	<li>Force clamp and nm stabilization feedback:
	<ol>
		<li>Switch on the ultrafast force-clamp with 2-3 pN force and 200 nm oscillation and scan the pedestal bead in discrete steps of about 20-30 nm in the direction perpendicular to the actin filament. Wait for interactions to occur at each position (few seconds), then step ahead if no interaction is observed. As the protein-filament interaction is established, look for the position where interactions are more frequent.</li>
		<li>Ensure that when the processive myosin moves towards one end of the actin filament (usually the + end), the trapped bead attached to the + end moves towards the silica bead. Move the stage towards the -end of the filament so that the silica bead lies as close as possible to the trapped bead attached to the -end of the filament when myosin is not bound and start the nanometer-stabilization feedback. In doing so, the probability that the trapped bead attached to the +end of the filament crashes into the silica bead is minimized.</li>
	</ol>
	</li>
	<li>Record data.</li>
</ol>
4. Data analysis4
NOTE: The analysis method that is described allows for the detection and measurement of processive runs and fast stepping events based on changes in the dumbbell velocity, as caused by myosin stepping. Analysis of processive runs is performed based on a data analysis method for non-processive motors described in references3,4,13.
<ol>
	<li>Set a threshold for velocity changes to allow for stepping event detection. Since in this case both forward and backward steps are expected, the crossing of the threshold is accepted in both directions.</li>
	<li>Assign each step to the corresponding run: if the time interval between two consequent steps is shorter than 3 ms and the amplitude of the steps is &#60; 90 nm, steps are assigned to the same run, otherwise steps are assigned to different runs4.</li>
	<li>Correct run length for assistive forces4.
	<ol>
		<li>Correct run lengths under assisting forces by calculating the real run length value RL from the measured average run length value &#60;RLm&#62; from the following equation, where D is the oscillation range: 
		<img alt="Equation 1" src="/files/ftp_upload/62388/62388eq1v3.jpg" /> 
		NOTE: Details on the derivation of this equation can be found in reference4.</li>
	</ol>
	</li>
</ol>

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The authors declare no competing interests.

Although single molecule techniques, such as the three-bead assay, are technically challenging and low throughput, UFFCS improves the detection of molecular interactions thanks to the high signal-to-noise ratio of the data. UFFCS allows the study of the load-dependence of motor proteins, with the main advantages of applying the force very rapidly upon binding of the motor to the filament to probe early and very rapid events in force production and weak binding states under controlled force; maintaining the force constant...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62388">Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultra Force-Clamp Spectroscopy</a>. The title was updated.
The title was updated from:
Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultra Force-Clamp Spectroscopy
to:
Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62388">Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultra Force-Clamp Spectroscopy</a>. The title was updated.

Erratum: Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultra Force-Clamp Spectroscopy

In the last decades optical tweezers have been a valuable tool to elucidate the mechanochemistry of protein interactions at the single molecule level, due to the striking possibility of concurrent manipulation and measurement of conformational changes and enzymatic kinetics 1,2. In particular, the capability to apply and measure forces in the range of those exerted by molecular motors in the cell, together with the capacity to measure sub-nanometer conformational changes, made optical tweezers a unique single-molecule tool for unraveling the chemomechanical properties of motor proteins and their mechanical regulation.
Ultrafast force-clamp spectroscopy (UFFCS) is a single-molecule force-spectroscopy technique based on optical tweezers, developed to study the fast kinetics of molecular motors under load in a three-bead geometry (Figure 1a)3,4. UFFCS reduces the time lag for force application to the motor protein to the physical limit of optical tweezers, i.e., the mechanical relaxation time of the system, thus allowing the application of the force rapidly after the beginning of a myosin run (few tens of microseconds)3. This capability has been exploited to investigate the early mechanical events in fast skeletal 3 and cardiac5 muscle myosin to reveal the load dependence of the powerstroke, the weak- and strong-binding states, as well as the order of biochemical (Pi) and mechanical (powerstroke) events.
The three-bead geometry is usually employed to study non-processive motors, a single bead geometry with a force-clamp has been commonly used to investigate processive non-conventional myosins such as myosin Va6. However, there are several reasons to prefer a three-bead UFFCS assay also for processive myosins. First, the rapid application of load right after actin-myosin binding allows the measurement of the early events in force development as in non-processive motors. In addition, in the case of processive motors it also allows an accurate measurement of the motor&#39;s run lengths and run durations under constant force all through their progression (Figure 1b). Moreover, because of the high rate of the force feedback, the system can maintain the force constant during fast changes in position, such as the myosin working stroke, thereby guaranteeing a constant load during motor stepping. The high-temporal resolution of the system allows the detection of sub-ms interactions, opening the possibility of investigating weak binding of myosin to actin. Finally, the assay geometry guarantees that the force is applied along the actin filament, with negligible transverse and vertical components of the force. This point is of particular relevance since the vertical force component has been shown to influence significantly the load-dependence of motor&#39;s kinetics7,8. By using this technique, we could apply a range of assistive and resistive loads to processive myosin-5B and directly measure the load dependence of its processivity for a wide range of forces4.
As shown in Figure 1a, in this system a single actin filament is suspended between two polystyrene beads trapped in the focus of double optical tweezers (the &#34;dumbbell&#34;). An imbalanced net force F= F1-F2 is imposed on the filament, through a fast feedback system, which makes the filament move at constant velocity in one direction until it reaches a user-defined inversion point where the net force is reversed in the opposite direction. When the motor protein is not interacting with the filament, the dumbbell is free to move back and forth in a triangular wave shape (Figure 1b, bottom panel) spanning the pedestal bead on which a single motor protein is attached. Once the interaction is established the force carried by the dumbbell is very rapidly transferred to the motor protein and the motor starts displacing the filament by stepping under the force intensity and direction that was applied by the feedback system at the time of the interaction, until myosin detaches from actin. Being the displacement produced by the stepping of the motor dependent on the polarity of the trapped actin filament, according to the direction of the applied force the load can be either assistive, i.e., pushing in the same direction of the motor displacement (push in Figure 1b upper panel), or resistive, i.e., pulling in the opposite direction with respect to the motor displacement (pull in Figure 1b upper panel) making it possible to study the chemomechanical regulation of the motor processivity by both the intensity and the directionality of the applied load.
In the next sections all the steps to measure actin-myosin-5B interactions under different loads with an ultrafast force-clamp spectroscopy setup are fully described, including 1) the setting up of the optical setup, optical traps alignment and calibration procedures, 2) the preparations of all the components and their assembly in the sample chamber, 3) the measurement procedure, 4) representative data and data analysis to extract important physical parameters, such as the run length, the step size and the velocity of the motor protein.

<table><tbody><tr><td>&amp;nbsp;Aliphatic Amine Latex Beads</td><td>ThermoFisher</td><td>A37362</td><td>1.0-&amp;mu;m diameter, 2% (w/v)</td></tr><tr><td>Acetone</td><td>Sigma</td><td>32201</td><td></td></tr><tr><td>Actin polymerization buffer</td><td>Cytoskeleton</td><td>BSA02</td><td>10X</td></tr><tr><td>AODs (acousto-optic deflectors)</td><td>AA Opto Electronic</td><td>DTS-XY 250</td><td>Laser beam deflectors</td></tr><tr><td>ATP</td><td>Sigma</td><td>A7699</td><td></td></tr><tr><td>Biotinylated-BSA</td><td>ThermoFisher</td><td>29130</td><td></td></tr><tr><td>BSA</td><td>Sigma</td><td>B4287</td><td></td></tr><tr><td>Calmodulin from porcine brain (CaM)</td><td>Merck Millipore</td><td>208783</td><td></td></tr><tr><td>Catalase from bovine liver</td><td>Sigma</td><td>C40</td><td></td></tr><tr><td>Condenser</td><td>Olympus</td><td>OlympusU-AAC, Aplanat, Achromat</td><td>NA 1.4, oil immersion</td></tr><tr><td>Creatine phosphate disodium salt tetrahydrate</td><td>Sigma</td><td>27920</td><td></td></tr><tr><td>Creatine Phosphokinase from rabbit muscle</td><td>Sigma</td><td>C3755</td><td></td></tr><tr><td>DDs</td><td>AA Opto Electronic</td><td>AA.DDS.XX</td><td>Two-channel digital synthesizer</td></tr><tr><td>DL-Dithiothreitol (DTT)/td&gt;</td><td>Sigma</td><td>43819</td><td></td></tr><tr><td>EGTA</td><td>Sigma</td><td>E4378</td><td></td></tr><tr><td>G-actin protein</td><td>Cytoskeleton</td><td>AKL99</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>G7528</td><td></td></tr><tr><td>Glucose Oxidase from Aspergillus niger</td><td>Sigma</td><td>&amp;nbsp;G7141</td><td></td></tr><tr><td>HaloTag succinimidyl ester O2 ligand</td><td>Promega</td><td>P1691</td><td></td></tr><tr><td>High vacuum silicone grease heavy</td><td>Merck Millipore</td><td>107921</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P9541</td><td></td></tr><tr><td>KH2PO4/K2HPO4</td><td>Sigma</td><td>P5379/ P8281</td><td></td></tr><tr><td>Labview</td><td>National Instruments</td><td>version 8.1</td><td>Data acquisition</td></tr><tr><td>Labview FPGA module</td><td>National Instruments</td><td>version 8.1</td><td>Fast Force-Clamp</td></tr><tr><td>Matlab</td><td>MathWorks</td><td>2016</td><td>Data analysis</td></tr><tr><td>MgCl2</td><td>Fluka</td><td>63020</td><td></td></tr><tr><td>Microscope Objective</td><td>Nikon</td><td>Plan-Apo 60X</td><td>NA 1.2, WD 0.2 mm, water imm.</td></tr><tr><td>MOPS</td><td>Sigma</td><td>M1254</td><td></td></tr><tr><td>Nitrocellulose</td><td>Sigma</td><td>N8267</td><td>0.45 pore size</td></tr><tr><td>Pentyl acetate solution</td><td>Sigma</td><td>46022</td><td></td></tr><tr><td>Pure Ethanol</td><td>&amp;nbsp;Sigma</td><td>2860</td><td></td></tr><tr><td>QPDs</td><td>UDT</td><td>DLS-20</td><td>D Position Detecto</td></tr><tr><td>Rhodamine BSA</td><td>Molecular Probes</td><td>A23016</td><td></td></tr><tr><td>Rhodamine Phalloidin</td><td>&amp;nbsp;Sigma</td><td>P1951</td><td></td></tr><tr><td>Silica beads</td><td>Bangslabs</td><td>SS04N</td><td>1.21 mm, 10% solids</td></tr><tr><td>Sodium azide</td><td>&amp;nbsp;Sigma</td><td>S2002</td><td></td></tr><tr><td>Streptavidin protein</td><td>&amp;nbsp;Sigma</td><td>189730</td><td></td></tr></tbody></table>

dissecting mechanoenzymatic properties of processive myosins with ultrafast force-clamp spectroscopy

Ultrafast force-clamp spectroscopy (UFFCS) is a single molecule technique based on laser tweezers that allows the investigation of the chemomechanics of both conventional and unconventional myosins under load with unprecedented time resolution. In particular, the possibility to probe myosin motors under constant force right after the actin-myosin bond formation, together with the high rate of the force feedback (200 kHz), has shown UFFCS to be a valuable tool to study the load dependence of fast dynamics such as the myosin working stroke. Moreover, UFFCS enables the study of how processive and non-processive myosin-actin interactions are influenced by the intensity and direction of the applied force.
By following this protocol, it will be possible to perform ultrafast force-clamp experiments on processive myosin-5 motors and on a variety of unconventional myosins. By some adjustments, the protocol could also be easily extended to the study of other classes of processive motors such as kinesins and dyneins. The protocol includes all the necessary steps, from the setup of the experimental apparatus to sample preparation, calibration procedures, data acquisition and analysis.

Representative data consist in position records over time as shown in Figure 4. In the position record two kinds of displacement are visible. Firstly, when the myosin motor is not interacting with the actin filament the trapped beads are moving at constant velocity against the viscous drag force of the solution showing a linear displacement oscillating within the oscillation range set by the operator in a triangular wave3 (not visible in <strong class...

Watch this Scientific Journal Video about Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy at JoVE.com

Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy

Presented here is a comprehensive protocol to perform ultrafast force-clamp experiments on processive myosin-5 motors, which could be easily extended to the study of other classes of processive motors. The protocol details all the necessary steps, from the setup of the experimental apparatus to sample preparation, data acquisition and analysis.

Ultrafast force-clamp spectroscopy (UFFCS) is a single molecule technique based on laser tweezers that allows the investigation of the chemomechanics of ...

dissecting-mechanoenzymatic-properties-processive-myosins-with

Research

JoVE Journal

Biology

1.3K Views.  National Institute of Optics, National Research Council. Presented here is a comprehensive protocol to perform ultrafast force-clamp experiments on processive myosin-5 motors, which could be easily extended to the study of other classes of processive motors. The protocol details all the necessary steps, from the setup of the experimental apparatus to sample preparation, data acquisition and analysis.

Video: Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy

Demonstrating the Uses of the Novel Gravitational Force Spectrometer to Stretch and Measure Fibrous Proteins

The study of macromolecular structure has become critical to the elucidation of molecular mechanisms and function. There are several limited, but important bioinstruments capable of testing the force dependence of structural features in proteins. Scale has been a limiting parameter on how accurately researchers can peer into the nanomechanical world of molecules, such as nucleic acids, enzymes, and motor proteins that perform life-sustaining work. Atomic force microscopy (AFM) is well tuned to determine native structures of fibrous proteins with a distance resolution on par with electron microscopy. However, in AFM force studies, the forces are typically much higher than a single molecule might experience 1, 2. Optical traps (OT) are very good at determining the relative distance between the trapped beads and they can impart very small forces 3. However, they do not yield accurate absolute lengths of the molecules under study. Molecular simulations provide supportive information to such experiments, but are limited in the ability to handle the same large molecular sizes, long time frames, and convince some researchers in the absence of other supporting evidence2, 4.
The gravitational force spectrometer (GFS) fills a critical niche in the arsenal of an investigator by providing a unique combination of abilities. This instrument is capable of generating forces typically with 98% or better accuracy from the femtonewton range to the nanonewton range. The distance measurements currently are capable of resolving the absolute molecular length down to five nanometers, and relative bead pair separation distances with a precision similar to an optical trap. Also, the GFS can determine stretching or uncoiling where the force is near equilibrium, or provide a graded force to juxtapose against any measured structural changes. It is even possible to determine how many amino acid residues are involved in uncoiling events under physiological force loads 2. Unlike in other methods where there is extensive force calibration that must precede any assay, the GFS requires no such force calibration 5. By complementing the strengths of other methods, the GFS will bridge gaps in understanding the nanomechanics of vital proteins and other macromolecules.

This is a step-by step guide showing the purpose, operation, and representative results from the novel gravitational force spectrometer.

The study of macromolecular structure has become critical to the elucidation of molecular mechanisms and function. There are several limited, but ...

Bioengineering

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 ...

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.

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. ...

Biochemistry

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic properties are adapted for the particular function they execute in cells. Myosin 5a processively walks on F-actin to transport melanosomes and vesicles in cells. Conversely, nonmuscle myosin 2b operates as a bipolar filament containing approximately 30 molecules. It moves F-actin of opposite polarity toward the center of the filament, where the myosin molecules work asynchronously to bind actin, impart a power stroke, and dissociate before repeating the cycle. Nonmuscle myosin 2b, along with its other nonmuscle myosin 2 isoforms, has roles that include&#160;cell adhesion, cytokinesis, and tension maintenance. The mechanochemistry of myosins can be studied by performing in vitro motility assays using purified proteins. In the gliding actin filament assay, the myosins are bound to a microscope coverslip surface and translocate fluorescently labeled F-actin, which can be tracked. In the single molecule/ensemble motility assay, however, F-actin is bound to a coverslip and the movement of fluorescently labeled myosin molecules on the F-actin is observed. In this report, the purification of recombinant myosin 5a from Sf9 cells using affinity chromatography is outlined. Following this, we outline two fluorescence microscopy-based assays: the gliding actin filament assay and the inverted motility assay. From these assays, parameters such as actin translocation velocities and single molecule run lengths and velocities can be extracted using the image analysis software. These techniques can also be applied to study the movement of single filaments of the nonmuscle myosin 2 isoforms, discussed herein in the context of nonmuscle myosin 2b. This workflow represents a protocol and a set of quantitative tools that can be used to study the single molecule and ensemble dynamics of nonmuscle myosins.

Presented here is a procedure to express and purify myosin 5a followed by a discussion of its characterization, using both ensemble and single molecule in vitro fluorescence microscopy-based assays, and how these methods can be modified for the characterization of nonmuscle myosin 2b.

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic ...

High-Speed Magnetic Tweezers for Nanomechanical Measurements on Force-Sensitive Elements

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are therefore poised to be useful in the field of mechanobiology. Since the method commonly relies on image-based tracking of magnetic beads, the speed limit in recording and analyzing images, as well as the thermal fluctuations of the beads, has long hampered its application in observing small and fast structural changes in target molecules. This article describes detailed methods for the construction and operation of a high-resolution MT setup that can resolve nanoscale, millisecond dynamics of biomolecules and their complexes. As application examples, experiments with DNA hairpins and SNARE complexes (membrane-fusion machinery) are demonstrated, focusing on how their transient states and transitions can be detected in the presence of piconewton-scale forces. We expect that high-speed MTs will continue to enable high-precision nanomechanical measurements on molecules that sense, transmit, and generate forces in cells, and thereby deepen our molecular-level understanding of mechanobiology.

Here, we describe a high-speed magnetic tweezer setup that performs nanomechanical measurements on force-sensitive biomolecules at the maximum rate of 1.2 kHz. We introduce its application to DNA hairpins and SNARE complexes as model systems, but it will be also applicable to other molecules involved in mechanobiological events.

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are ...

A Typical Workflow to Simulate Cytoskeletal Systems

Many cytoskeletal systems are now sufficiently well known to permit their precise quantitative modeling. Microtubule and actin filaments are well characterized, and the associated proteins are often known, as well as their abundance and the interactions between these elements. Thus, computer simulations can be used to investigate the collective behavior of the system precisely, in a way that is complementary to experiments. Cytosim is an Open Source cytoskeleton simulation suite designed to handle large systems of flexible filaments with associated proteins such as molecular motors. It also offers the possibility to simulate passive crosslinkers, diffusible crosslinkers, nucleators, cutters, and discrete versions of the motors that only step on unoccupied lattice sites on a filament. Other objects complement the filaments by offering spherical or more complicated geometry that can be used to represent chromosomes, the nucleus, or vesicles in the cell.
Cytosim offers simple command-line tools for running a simulation and displaying its results, which are versatile and do not require programming skills. In this workflow, step-by-step instructions are given to i) install the necessary environment on a new computer, ii) configure Cytosim to simulate the contraction of a 2D actomyosin network, and iii) produce a visual representation of the system. Next, the system is probed by systematically varying a key parameter: the number of crosslinkers. Finally, the visual representation of the system is complemented by the numerical quantification of contractility to view, in a graph, how contractility depends on the composition of the system. Overall, these different steps constitute a typical workflow that can be applied with few modifications to tackle many other problems in the cytoskeletal field.

This protocol demonstrates how to use Cytosim, an Open Source cytoskeleton simulation, to investigate the behavior of a network of filaments connected by molecular motors and passive crosslinkers. A generic workflow with step-by-step instructions is followed to vary the number of crosslinkers and plot the resulting network contractility.

Many cytoskeletal systems are now sufficiently well known to permit their precise quantitative modeling. Microtubule and actin filaments are well ...

Developmental Biology

Recombinant Human Myosin-7a Production and Functional Characterization Protocol

MultiBac System-Based Purification and Biophysical Characterization of Human Myosin-7a

Myosin-7a is an actin-based motor protein vital for auditory and visual processes. Mutations in myosin-7a lead to Usher syndrome type 1, the most common and severe form of deaf-blindness in humans. It is hypothesized that myosin-7a forms a transmembrane adhesion complex with other Usher proteins, essential for the structural-functional integrity of photoreceptor and cochlear hair cells. However, due to the challenges in obtaining pure, intact protein, the exact functional mechanisms of human myosin-7a remain elusive, with limited structural and biomechanical studies available. Recent studies have shown that mammalian myosin-7a is a multimeric motor complex consisting of a heavy chain and three types of light chains: regulatory light chain (RLC), calmodulin, and calmodulin-like protein 4 (CALML4). Unlike calmodulin, CALML4 does not bind to calcium ions. Both the calcium-sensitive, and insensitive calmodulins are critical for mammalian myosin-7a for proper fine-tuning of its mechanical properties. Here, we describe a detailed method to produce recombinant human myosin-7a holoenzyme using the MultiBac Baculovirus protein expression system. This yields milligram quantities of high-purity full-length protein, allowing for its biochemical and biophysical characterization. We further present a protocol for assessing its mechanical and motile properties using tailored in vitro motility assays and fluorescence microscopy. The availability of the intact human myosin-7a protein, along with the detailed functional characterization protocol described here, paves the way for further investigations into the molecular aspects of myosin-7a in vision and hearing.

Author Spotlight: Unraveling the Role of Myosin-7a and Usher Proteins in Hearing and Human Disease

This protocol details the procedures for recombinantly producing the human myosin-7a holoenzyme using the MultiBac Baculovirus system and for studying its motility using a tailored in vitro filament gliding assay.

Myosin-7a is an actin-based motor protein vital for auditory and visual processes. Mutations in myosin-7a lead to Usher syndrome type 1, the most common ...

Probing Myosin Ensemble Mechanics in Actin Filament Bundles Using Optical Tweezers

Myosins are motor proteins that hydrolyze ATP to step along actin filament (AF) tracks and are essential in cellular processes such as motility and muscle contraction. To understand their force-generating mechanisms, myosin II has been investigated both at the single-molecule (SM) level and as teams of motors in vitro using biophysical methods such as optical trapping.
These studies showed that myosin force-generating behavior can differ greatly when moving from the single-molecule level in a three-bead arrangement to groups of motors working together on a rigid bead or coverslip surface in a gliding arrangement. However, these assay constructions do not permit evaluating the group dynamics of myosin within viscoelastic structural hierarchy as they would within a cell. We have developed a method using optical tweezers to investigate the mechanics of force generation by myosin ensembles interacting with multiple actin filaments.
These actomyosin bundles facilitate investigation in a hierarchical and compliant environment that captures motor communication and ensemble force output. The customizable nature of the assay allows for altering experimental conditions to understand how modifications to the myosin ensemble, actin filament bundle, or the surrounding environment result in differing force outputs.

Formation of actomyosin bundles in vitro and measuring myosin ensemble force generation using optical tweezers is presented and discussed.

Myosins are motor proteins that hydrolyze ATP to step along actin filament (AF) tracks and are essential in cellular processes such as motility and muscle ...

Ensemble Force Spectroscopy by Shear Forces

Single-molecule techniques based on fluorescence and mechanochemical principles provide superior sensitivity in biological sensing. However, due to the lack of high throughput capabilities, the application of these techniques is limited in biophysics. Ensemble force spectroscopy (EFS) has demonstrated high throughput in the investigation of a massive set of molecular structures by converting mechanochemical studies of individual molecules into those of molecular ensembles. In this protocol, the DNA secondary structures (i-motifs) were unfolded in the shear flow between the rotor and stator of a homogenizer tip at shear rates up to 77796/s. The effects of flow rates and molecular sizes on the shear forces experienced by the i-motif were demonstrated. The EFS technique also revealed the binding affinity between DNA i-motifs and ligands. Furthermore, we have demonstrated a click chemistry reaction that can be actuated by shear force (i.e., mechano-click chemistry). These results establish the effectiveness of using shear force to control the conformation of molecular structures.

Ensemble force spectroscopy (EFS) is a robust technique for mechanical unfolding and real-time sensing of an ensemble set of biomolecular structures in biophysical and biosensing fields.

Single-molecule techniques based on fluorescence and mechanochemical principles provide superior sensitivity in biological sensing. However, due to the ...

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This feature is attributed to the mechanical and dynamic properties of the cell cytoskeleton, notably, the actomyosin cytoskeleton, which is an active gel of polar actin filaments, myosin motors, and accessory proteins that exhibit intrinsic contraction properties. The usually accepted view is that the cytoskeleton behaves as a viscoelastic material. However, this model cannot always explain the experimental results, which are more consistent with a picture describing the cytoskeleton as a poroelastic active material-an elastic network embedded with cytosol. Contractility gradients generated by the myosin motors drive the flow of the cytosol across the gel pores, which infers that the mechanics of the cytoskeleton and the cytosol are tightly coupled. One main feature of poroelasticity is the diffusive relaxation of stresses in the network, characterized by an effective diffusion constant that depends on the gel elastic modulus, porosity, and cytosol (solvent) viscosity. As cells have many ways to regulate their structure and material properties, our current understanding of how cytoskeleton mechanics and cytosol flow dynamics are coupled remains poorly understood. Here, an in vitro reconstitution approach is employed to characterize the material properties of poroelastic actomyosin gels as a model system for the cell cytoskeleton. Gel contraction is driven by myosin motor contractility, which leads to the emergence of a flow of the penetrating solvent. The paper describes how to prepare these gels and run experiments. We also discuss how to measure and analyze the solvent flow and gel contraction both at the local and global scales. The various scaling relations used for data quantification are given. Finally, the experimental challenges and common pitfalls are discussed, including their relevance to cell cytoskeleton mechanics.

The Mechanics of Poro-Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

In this work, an in vitro reconstitution approach is employed to study the poroelasticity of actomyosin gels under controlled conditions. The dynamics of the actomyosin gel and the embedded solvent are quantified, through which the network poroelasticity is demonstrated. We also discuss the experimental challenges, common pitfalls, and relevance to cell cytoskeleton mechanics.

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This ...