This work is supported in part by the University of Mississippi Graduate Student Council Research Fellowship (OA), University of Mississippi Sally McDonnell-Barksdale Honors College (JCW, JER), the Mississippi Space Grant Consortium under grant number NNX15AH78H (JCW, DNR), and the American Heart Association under grant number 848586 (DNR).

1. Etching coverslips
<ol>
	<li>Dissolve 100 g of KOH in 300 mL of 100% ethanol in a 1,000 mL beaker. Stir with a stir bar until the majority of the KOH has dissolved. 
	CAUTION: Concentrated KOH solution can cause burns and damage to clothing. Wear gloves, eye protection, and a lab coat.</li>
	<li>Place coverslips individually in coverslip cleaning racks. 
	NOTE: Racks are designed with slits that hold single coverslips spaced apart to allow for etching and rinsing on each face of the coverslip, drain holes in the bottom, and made of material that can withstand the harsh etching conditions. They can be custom-made or purchased commercially.</li>
	<li>Prepare and label three 1,000 mL beakers: one with 300 mL of ethanol and two beakers with 300 mL of reverse osmosis (RO) water. 
	NOTE: Here, RO water was sourced from a lab water purifier, but it could also be purchased commercially if a local purifier is not available.</li>
	<li>Place each of the four beakers in a bath sonicator to degas for 5 min.</li>
	<li>Submerge a rack of coverslips in the beaker of KOH and ethanol and sonicate for 5 min.</li>
	<li>Transfer the rack of coverslips from the KOH/ethanol beaker to the ethanol-only beaker. Dip rack up and down in the beaker until there is no beading. 
	NOTE: Take care to not disturb the coverslips or forcefully drop the rack into the beaker. This will cause the coverslips to come out of the rack or cause chemical splashing.</li>
	<li>Carefully transfer the rack of coverslips from the ethanol beaker to a beaker of water, dipping up and down until there is no beading.</li>
	<li>Submerge the rack of coverslips in the beaker of water that has not been used yet and sonicate again for 5 min.</li>
	<li>Use a bottle to spray the rack of coverslips with water until it runs off the coverslips smoothly. Repeat with the ethanol.</li>
	<li>Place the racks to dry in an oven at 90 &#176;C for 20 min. Store the racks of etched coverslips at room temperature in closed containers to prevent contamination before use.</li>
</ol>
2. Actin filament polymerization
<ol>
	<li>Make Solution T
	<ol>
		<li>In a 50 mL conical tube, add 3.94 g of Tris-HCl and 0.147 g of CaCl2. Add RO water to make a total volume of 50 mL and mix well. 
		NOTE: The final concentrations of Solution T are 500 mM Tris-HCl and 20 mM CaCl2, respectively.</li>
		<li>Label the tube Solution T and store it at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Make TC Buffer
	<ol>
		<li>Mix 40 mL of RO water and 1.5 mL of Solution T in a 50 mL conical tube. Change the pH to 8.0 by adding small amounts of concentrated KOH. Add water to make 50 mL of the solution, and verify the pH. Adjust the pH if needed. 
		NOTE: The final TC buffer contains 5 mM Tris-HCl and 0.2 mM CaCl2 at pH 8.</li>
		<li>Label the tube TC and store it at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Make FC Buffer
	<ol>
		<li>Add 85 mL of RO water, 10 mL of Solution T, 3.73 g of KCl, and 0.041 g of MgCl2 to a 100 mL buffer bottle. Modify the pH to 7.5 by adding small volumes of concentrated KOH. Add water to make a final volume of 100 mL and verify the pH. 
		NOTE: The final FC buffer contains 500 mM Tris-HCl, 500 mM KCl, 2 mM MgCl2, and 2 mM CaCl2 at pH 7.5.</li>
		<li>Label the tube FC and store it at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare General Actin Buffer (GAB).
	<ol>
		<li>Mix 485 &#956;L of TC buffer, 10 &#956;L of 10 mM ATP, and 5 &#956;L of 50 mM DTT in a microcentrifuge tube. 
		NOTE: Final buffer conditions are 5 mM Tris-HCl, 0.2 mM CaCl2, 0.5 mM DTT, and 0.2 mM ATP.</li>
		<li>Label it as GAB and store it at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare Actin Polymerization Buffer (APB).
	<ol>
		<li>Mix 455 &#956;L of FC buffer, 25 &#956;L of 100 mM ATP, and 20 &#956;L of 50 mM DTT in a microcentrifuge tube. 
		NOTE: The final buffer conditions are 50 mM Tris-HCl, 500 mM KCl, 2 mM MgCl2, 2 mM CaCl2 2 mM DTT, and 5 mM ATP.</li>
		<li>Label the tube as APB and store it at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Reconstitute actin
	<ol>
		<li>Reconstitute rabbit skeletal muscle actin by adding 100 &#956;L of deionized water to a 1 mg vial of lyophilized actin. Mix well by gently pipetting up and down. Aliquot into 5 &#956;L samples, snap-freeze, and store the 10 mg/mL actin aliquots at -80 &#176;C.</li>
		<li>Reconstitute biotinylated rabbit skeletal muscle actin by adding 20 &#956;L of RO water. Aliquot into 5 &#956;L samples, snap-freeze, and store the 1 mg/mL biotinylated actin aliquots at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Non-labeled actin polymerization with rhodamine phalloidin stabilization
	<ol>
		<li>Thaw one vial of 10 mg/mL actin and keep it on ice.</li>
		<li>Prepare fresh GAB buffer, add 100 &#956;L of GAB to the actin aliquot, and mix by gently pipetting up and down. Incubate the solution on ice for 1 h.</li>
		<li>Prepare fresh APB during the incubation. After incubation, polymerize the actin into filaments by adding 11 &#956;L of APB to the actin solution. Mix well by gently pipetting up and down. Place on ice for 20 min.</li>
		<li>Add 5 &#956;L of rhodamine-labeled phalloidin to the freshly polymerized actin filament solution. Leave on ice in the dark for 1 h.</li>
		<li>Store the rhodamine actin vial wrapped in aluminum foil in the dark at 4 &#176;C. 
		NOTE: It is suggested to use these filaments for a maximum period of 1 week. AF quality can be confirmed each day through a quick imaging of a flow cell containing only AFs and viewing consistent filaments day to day.</li>
	</ol>
	</li>
	<li>Biotinylated actin polymerization with Alexa Fluor 488 phalloidin stabilization
	<ol>
		<li>Thaw one vial of 10 mg/mL actin and 1 vial of 1 mg/mL biotinylated actin and keep them on ice.</li>
		<li>Make fresh GAB buffer.</li>
		<li>Combine the two vials (step 2.8.1) in a 10:1 actin:biotinylated actin ratio. Add 100 &#956;L of GAB to the actin mixture and mix well by gently pipetting up and down. Incubate on ice for 1 h.</li>
		<li>Make fresh APB during the incubation.</li>
		<li>After the incubation step, polymerize the actin by adding 11 &#956;L of APB to the actin solution. Mix well by pipetting up and down gently. Incubate on ice for 20 min.</li>
		<li>Add 5 &#956;L of Alexa Fluor 488-labeled phalloidin and incubate on ice in the dark for 1 h.</li>
		<li>Store the biotinylated actin vial wrapped in aluminum foil in the dark at 4 &#176;C. 
		&#8203;NOTE: These filaments can be used for a maximum period of 1 week.</li>
	</ol>
	</li>
</ol>
3. Myosin and bead preparation
<ol>
	<li>Reconstitute Myosin II
	<ol>
		<li>Briefly spin down (~5 s) lyophilized skeletal myosin II to collect it at the bottom of the tube using a standard minicentrifuge.</li>
		<li>Reconstitute the myosin to 10 mg/mL by adding 100 &#956;L of 1 mM DTT prepared in RO water.</li>
		<li>Dilute the stock myosin solution 10x by adding 10 &#956;L of 10 mg/mL myosin to 90 &#956;L of 1 mM DTT in RO water. Make small-volume (1-5 &#956;L) aliquots, snap-freeze, and store at -80 &#176;C. 
		NOTE: Myosin activity can be confirmed by performing a standard gliding filament assay as published previously46,47. See the discussion for a brief description.</li>
	</ol>
	</li>
	<li>Cleaning streptavidin-coated beads
	<ol>
		<li>Dilute 20 &#956;L of 1 &#956;m streptavidin beads into 80 &#956;L of RO water. Wash four times by spinning down at 9,600 &#215; g and reconstituting in 100 &#956;L of RO water.</li>
		<li>Sonicate for 2 min at 40% amplitude and store the washed beads on a rotator at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
4. Flow cell preparation
<ol>
	<li>Prepare a poly-l-lysine solution (PLL) by adding 30 mL of 100% ethanol to a 50 mL tube and adding 200 &#956;L of 0.1% w/v poly-l-lysine in water and mix well.</li>
	<li>Add an etched coverslip to the PLL solution and allow it to soak for 15 min. Remove the coverslip with tweezers, taking care to only touch the edge of the coverslip as it is pulled up from the tube (see Figure 1A-C). Grab the coverslips by their edges with a gloved hand.</li>
	<li>Dry the coverslip with a filtered airline until there is no ethanol left and no residue on the coverslip.</li>
	<li>Apply two pieces of double-sided sticky tape to the middle of a microscope slide, 3-4 mm apart from each other. Tear or cut off the excess tape that hangs off the edge of the slide.</li>
	<li>Add the PLL-coated coverslip on top of the tape perpendicular to the long axis of the microscope slide (forming a T) to form a channel.</li>
	<li>Use a small tube to compress the coverslip onto the tape and microscope slide thoroughly until the tape is transparent (Figure 1A). Ensure there are no bubbles in the tape as this can cause leakage from the flow channel. 
	&#8203;NOTE: The flow cell can hold a volume of 10-15 &#956;L.</li>
</ol>
5. Actomyosin bundle preparation
<ol>
	<li>In separate tubes, dilute each type of actin filament (rhodamine- and biotinylated 488-labeled) 600x by mixing 0.5 &#956;L of the respective, labeled actin with 300 &#956;L of APB. Add an additional 5 &#956;L of the correspondingly labeled phalloidin to each tube and incubate on ice in the dark for 15 min.</li>
	<li>To the biotinylated actin solution, add an oxygen scavenging system of 1 &#956;L of beta-D-glucose at 500 mg/mL, 1 &#956;L of glucose oxidase at 25 mg/mL, and 1 &#956;L of catalase at 500 units/mL. Add 1 &#956;L of 100 mM ATP and 1 &#956;L of 100x diluted, cleaned streptavidin beads. Gently stir with a pipette tip. Put the suspension on a rotator at 4 &#176;C while the rest of the actomyosin bundle is being assembled.</li>
	<li>Add 15 &#956;L of the diluted rhodamine actin to the PLL flow cell (Figure 1D). Wick the excess solution through the flow cell but do not allow the flow channel to become dry. Incubate for 10 min in a humidity chamber. 
	NOTE: Humidity chambers can be made from empty pipette tip boxes with water added to the bottom and the lid covered in aluminum foil to block light.</li>
	<li>Prepare a 1 mg/mL casein solution in APB.</li>
	<li>Add 15 &#956;L of 1 mg/mL casein to prevent non-specific binding of the subsequent components (Figure 1E). Incubate for 5 min in a humidity chamber.</li>
	<li>Add the desired concentration of myosin to the biotinylated actin and bead suspension from step 5.2. Gently stir with the pipette tip, and then immediately add 15 &#956;L of the step 5.2 suspension + the desired myosin concentration to the flow cell (Figure 1F,G). Incubate for 20 min. Seal the open ends of the flow cell with nail polish to prevent evaporation during imaging and optical trapping experiments. 
	NOTE: A myosin solution concentration of 1 &#956;M yields robust bundling and can be used as a starting point for the desired customization of the assay (see Figure 2).</li>
</ol>
6. Force measurements using Optical Trap (NT2 Nanotracker2)
NOTE: While the protocol below is specifically for the NT2 system, this assay can be used with other optical trapping systems, including those that are custom-built, that also have fluorescence capabilities. The general workflow remains the same of getting the surface of the slide in focus, performing bead calibrations, and acquiring data by finding fluorescent actin bundles. For the NT2 system, Supplemental Figure S1, Supplemental Figure S2, Supplemental Figure S3, Supplemental Figure S4, Supplemental Figure S5, Supplemental Figure S6, and Supplemental Figure S7 provide details of the optical trapping system and the software interface.
<ol>
	<li>Turn on the control box and laser (Supplemental Figure S1).</li>
	<li>Start the optical trap computer software by clicking on the JPK Nanotracker icon on the desktop.</li>
	<li>Wake up the remote controller by clicking on the Logitech button at the center (Supplemental Figure S2).</li>
	<li>Turn on the fluorescence module by toggling the on/off switch (Supplemental Figure S3).</li>
	<li>Turn the filter cube turret for brightfield imaging (Supplemental Figure S4).</li>
	<li>Once the system is ready, turn on the laser using the Laser Power button at the left-bottom corner of the screen to 50 mW and let it stabilize for 30 min (Supplemental Figure S5).</li>
	<li>Sequentially click on the Illumination, Camera, Objective, and Stage Movement buttons within the software to bring up those windows for viewing and manipulation during the experiment. Turn the microscope illumination on by clicking on the On/Off button and setting it to maximum power by clicking and dragging the bar all the way to the right (Supplemental Figure S5).</li>
	<li>Open the sample area and remove the sample holder from the microscope stage. Add the flow cell, secure it with the metal sample holders, and make sure that the slide with the coverslip is on the bottom.</li>
	<li>Add 30 &#956;L of RO water to the center of the bottom objective. Do not let the pipette tip touch the lens. Reinsert the sample stage. 
	NOTE: As the NT2 system uses a water immersion objective as the trapping objective, the immersion media may be different depending on the trapping objective in the user&#39;s setup.</li>
	<li>Raise the lower objective using the on-screen control arrows or L2 on the remote controller until the bead of water touches the coverslip (Supplemental Figure S5).</li>
	<li>Lower the top objective until about half the distance to the flow cell is reached using the on-screen arrows or R2 on the remote controller. Add 170 &#956;L of RO water to the top of the flow cell directly under the top objective. Lower the top objective until it breaks the surface tension of the water and forms a meniscus.</li>
	<li>Move the microscope stage using the arrow pad on the remote controller until the edge of the tape adjacent to the flow channel is reached. Close the sample door. 
	NOTE: A &#34;click&#34; upon closing the sample door indicates that the laser shutter is now open. This is a safety feature that only allows the shutter to open if the door is closed.</li>
	<li>Using the Objective window in the screen, bring the edge of the tape in focus by bringing the bottom objective named Laser Objective up by clicking on the upper arrow using the on-screen controls. Do the same for the top objective by clicking bottom arrow (Supplemental Figure S5). 
	NOTE: The double arrows move the objective or stage faster. The edge of the tape is used for focusing because it is a large, easy-to-find object that is close to the coverslip surface. Air bubbles within the tape are another option. However, this is not required if the user has an automated routine to find the surface focus or a preferred in-house method.</li>
	<li>Once the tape is in focus, partially close the iris at the top of the optical trap. Bring the top objective down until the polygon shape of the iris is visible. Bring those edges in focus, reopen the iris, and then couple the objectives using by clicking on the Padlock icon (Supplemental Figure S5).</li>
	<li>Find a floating bead and trap it by clicking on the Trap Shutter button, which will open the shutter and allow the trapping laser to hit the sample. Click on the Trap cursor on the screen and drag it to move the location of the trapping laser. Once trapped, calibrate the bead to correlate voltage measurements to force and displacement.</li>
	<li>Click on the Calibration button. Adjust the calibration routine based on power spectra analysis and fit the corner frequency within the software for the X, Y, and Z directions (Supplementary Figure S6).</li>
	<li>Click on Settings. Type in the diameter of the bead (1,000 nm), and type in the temperature of the stage found in the bottom left of the software window. (see Supplemental Figure S6).</li>
	<li>Click on Trap 1. Click on X Signal. Click on Run to perform the corner frequency fit. Click and drag within the window to optimize the function fit. Click on Use It for sensitivity and stiffness values. Click on Accept Values. Repeat for the Y and Z signals. Close the window. (see Supplemental Figure S6). 
	NOTE: Bead calibration routines on other optical trapping systems or custom-built systems that have been robustly tested by the user, such as the equipartition method or drag force method, are also acceptable57,58.</li>
	<li>Find an actomyosin bundle by searching for beads bound to AFs on the surface of the coverslip.</li>
	<li>When a bead uncrowded by other floating beads is detected, observe the AFs around it by fluorescence imaging to verify the presence of a bundle.</li>
	<li>Verify that a bundle is present by looking for both fluorescent AFs colocalized. Turn on the white light source and use the appropriate filter cube to image each actin filament by turning the turret (488 nm and 532 nm excitation filter cubes for Alexa Fluor 488 and rhodamine excitation, respectively). See Supplemental Figure S4. 
	NOTE: A control experiment to verify the fluorescence intensity of single AFs can be useful in identifying bundles that are composed of a single 488- and single rhodamine-labeled filaments, or applicable to whichever set of fluorophores the user chooses to use.</li>
	<li>Once verified, trap the bead attached to the top filament of the bundle by clicking on the Trap Shutter button.</li>
	<li>Use the on-screen controls to record the data by clicking on the Oscilloscope button (Supplemental Figure S7). To visualize measurements without recording the data, click on Start. To save all data, click on Autosave. To record measurements, click on Start Record. Choose which data are to be visualized in real-time (position, force, x-direction, y-direction) by choosing from the drop down menu X signal or Y signal. Remember that xdirection is left to right, and y-direction is up and down on the screen. See Supplemental Figure S7. 
	NOTE: Data will be saved as .out files and includes time, voltage, displacement, and force for each direction. These files can be exported into other software for visualization and analysis.</li>
</ol>

The authors have no conflicts of interest to declare.

An in vitro study using optical tweezers combined with fluorescence imaging was performed to investigate the dynamics of myosin ensembles interacting with actin filaments. Actin-myosin-actin bundles were assembled using muscle myosin II, rhodamine actin at the bottom of the bundle and on the coverslip surface, and 488-labeled, biotinylated actin filaments on the top of the bundle. Actin protein from rabbit muscle was polymerized and stabilized using general actin buffers (GAB) and actin polymerizing buffers (APB...

Motor proteins are essential to life, converting chemical energy into mechanical work1,2,3. Myosin motors interact with actin filaments by taking steps along the filaments similar to a track, and the dynamics of actin-myosin networks carry out muscle contraction, cell motility, the contractile ring during cytokinesis, and movement of cargo inside the cell, among other essential tasks3,4,5,6,7,8. Since myosins have so many essential roles, failure in the functionality of the myosin-actin network can lead to disease development, such as mutations in the myosin heavy chain that cause heart hypercontractility in hypertrophic cardiomyopathy (HCM)9,10,11,12,13,14. In muscle contraction, individual myosin motors cooperate with each other by working as an ensemble to provide the required mechanical energy that carries out the relative sliding of AFs4,15,16,17,18. Myosin motors form crossbridges between AFs and use conformational changes due to its mechanochemical cycle to collectively move toward the barbed end of the aligned filaments17,18,19,20,21.
Development of quantitative in vitro motility assays at the SM level using techniques such as optical trapping has facilitated gathering unprecedented detail of how individual myosin motors function, including measuring SM force generation and step sizes22,23,24,25,26,27,28,29,30. Finer et al. developed the &#34;three-bead&#34; or &#34;dumbbell&#34; optical trapping assay to probe the force-generation mechanics of single myosin II motors23,31. As muscle myosin II works in teams to contract AFs but is non-processive at the SM level, the optical trapping assay orientation had to be rearranged from the classic motor-bound bead approach32. To form the dumbbell assay, two optical traps were used to hold an AF over a myosin motor bound to a coverslip-attached bead, and force output by the single motor was measured through movements of the AF within the trap23.
However, SM forces and using a single motor/single filament assay orientation do not give a full image about system-level force generation since many motor proteins, including myosin II, do not work in isolation and often do not function as a sum of their parts15,16,17,32,33,34,35,36. More complex structures that include more than one motor interacting with more than one filament are necessary to better understand the synergy of myosin and actin filaments&#39; networks15,32. The dumbbell assay orientation has been exploited to investigate small ensemble force generation by having multiple myosins attached to a bead or using a myosin-thick filament attached to a surface and allowing the motors to interact with the suspended AF4,23,34,37,38,39,40.
Other small ensemble assays include an in vitro filament gliding assay wherein myosin motors are coated onto a coverslip surface, and a bead bound to an AF is used to probe the force generated by the team of motors4,35,36,38,39,40,41,42,43. In both these cases, the myosins are bound to a rigid surface &#8211; bead or coverslip &#8211; and utilize one AF. In these cases, the motors are not able to move freely or communicate with each other, nor does having myosins rigidly bound reflect the compliant, hierarchical environment in which the motors would work together in the sarcomere32. Previous studies have suggested that myosin II can sense its environment and adapt accordingly to changing viscoelastic or motor concentration conditions by altering characteristics such as force generation and duty ratio41,44,45. Thus, there is a need to develop an optical trapping assay that fosters and captures motor communication and system compliancy to paint a more realistic picture of the mechanistic underpinnings of myosin II ensemble force generation.
Here, we developed a method to couple hierarchical structure in vitro with optical trapping by forming actomyosin bundles or sandwiches consisting of multiple myosin motors interacting between two actin filaments. This modular assay geometry has the ability to directly probe how molecular and environmental factors influence ensemble myosin force generation. Further, investigating force generation mechanisms through these actin-myosin ensembles have the potential to aid in modeling and understanding how large-scale cellular tasks, such as muscle contraction, propagate up from the molecular level9,10,13.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Actin protein (biotin): skeletal muscle</td>
			<td>Cytoskeleton</td>
			<td>AB07-A</td>
			<td>Biotinylated actin protein</td>
		</tr>
		<tr>
			<td>Actin protein, rabbit skeletal muscle</td>
			<td>Cytoskeleton</td>
			<td>AKL99-A</td>
			<td>Actin protein</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Phalloidin</td>
			<td>Invitrogen</td>
			<td>A12379</td>
			<td>Actin stabilizer and Alexa Fluor 488 stain</td>
		</tr>
		<tr>
			<td>ATP</td>
			<td>Fisher scientific</td>
			<td>BP413-25</td>
			<td>Required for actin assembly and myosin motility</td>
		</tr>
		<tr>
			<td>Beta-D-glucose</td>
			<td>Fisher scientific</td>
			<td>MP218069110</td>
			<td>Part of oxygen scavenging system used to reduce photobleaching during fluorescence imaging</td>
		</tr>
		<tr>
			<td>Blotting Grade Blocker (casein)</td>
			<td>Biorad</td>
			<td>1706404</td>
			<td>Used to block surface from non-specific binding</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Fisher scientific</td>
			<td>C79500</td>
			<td>Calcium chloride, provides the necessary control over the dynamics of actin myosin network</td>
		</tr>
		<tr>
			<td>Catalase</td>
			<td>Fisher scientific</td>
			<td>ICN10040280</td>
			<td>Part of oxygen scavenging system used to reduce photobleaching during fluorescence imaging</td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Fisher scientific</td>
			<td>12544C</td>
			<td>Used to make flow cells</td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>Fisher scientific</td>
			<td>AC327190010</td>
			<td>Used for buffer preparation</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Fisher scientific</td>
			<td>A4094</td>
			<td>Regent used for cleaning coverslips</td>
		</tr>
		<tr>
			<td>Glucose oxidase</td>
			<td>Fisher scientific</td>
			<td>34-538-610KU</td>
			<td>Part of oxygen scavenging system used to reduce photobleaching during fluorescence imaging</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher scientific</td>
			<td>P217-500</td>
			<td>Used for buffer preparation</td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>Fisher scientific</td>
			<td>P250-1</td>
			<td>Used to etch coverslips and adjust buffer pH</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Fisher scientific</td>
			<td>M33-500</td>
			<td>Used for buffer preparation</td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>Fisher scientific</td>
			<td>12-544-2</td>
			<td>Used to make flow cells</td>
		</tr>
		<tr>
			<td>Myosin II protein: rabbit skeletal muscle</td>
			<td>Cytoskeleton</td>
			<td>MY02</td>
			<td>Full length myosin motor protein isolated from rabbit skeletal muscle</td>
		</tr>
		<tr>
			<td>Nanotracker2</td>
			<td>Bruker/JPK</td>
			<td>NT2</td>
			<td>Optical trapping instrument</td>
		</tr>
		<tr>
			<td>Poly-l-lysine</td>
			<td>Sigma-Aldrich</td>
			<td>P8920</td>
			<td>Facilities adhesion of actin filaments onto glass surface of the coverslip</td>
		</tr>
		<tr>
			<td>Rhodamine Phalloidin</td>
			<td>Cytoskeleton</td>
			<td>PHDR1</td>
			<td>Actin stabilizer and rhodamine fluorescent stain</td>
		</tr>
		<tr>
			<td>Streptavidin beads, 1 &#956;m</td>
			<td>Spherotech</td>
			<td>SVP-10-5</td>
			<td>Optical trapping handle</td>
		</tr>
		<tr>
			<td>Tris-HCl</td>
			<td>Fisher scientific</td>
			<td>PR H5121</td>
			<td>Used for buffer preparation</td>
		</tr>
	</tbody>
</table>

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.

Flow cells containing the actomyosin bundle systems are of a standard design, consisting of a microscope slide and an etched coverslip separated by a channel made from double-sided sticky tape (Figure 1). The assay is then built from the coverslip up using staged introductions as described in the protocol. The final assay consists of template rhodamine-labeled actin filaments; the desired myosin concentration (1 &#956;M was used for the representative results in Figure 2...

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Probing Myosin Ensemble Mechanics in Actin Filament Bundles Using Optical Tweezers

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 ...

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2.1K Views.  University of Mississippi. Formation of actomyosin bundles in vitro and measuring myosin ensemble force generation using optical tweezers is presented and discussed.

Video: Probing Myosin Ensemble Mechanics in Actin Filament Bundles Using Optical Tweezers

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not have been accomplished by other approaches1. With the use of the calcium- and voltage-sensitive dyes, optical mapping allows measurement of transmembrane action potentials and calcium transients with high spatial resolution without the physical contact with the tissue. This makes measurements of the cardiac electrical activity possible under many conditions where the use of electrodes is inconvenient or impossible1. For example, optical recordings provide accurate morphological changes of membrane potential during and immediately after stimulation and defibrillation, while conventional electrode techniques suffer from stimulus-induced artifacts during and after stimuli due to electrode polarization1. 
The Langendorff-perfused rabbit heart is one of the most studied models of human heart physiology and pathophysiology. Many types of arrhythmias observed clinically could be recapitulated in the rabbit heart model. It was shown that wave patterns in the rabbit heart during ventricular arrhythmias, determined by effective size of the heart and the wavelength of reentry, are very similar to that in the human heart2. It was also shown that critical aspects of excitation-contraction (EC) coupling in rabbit myocardium, such as the relative contribution of sarcoplasmic reticulum (SR), is very similar to human EC coupling3. Here we present the basic procedures of optical mapping experiments in Langendorff-perfused rabbit hearts, including the Langendorff perfusion system setup, the optical mapping systems setup, the isolation and cannulation of the heart, perfusion and dye-staining of the heart, excitation-contraction uncoupling, and collection of optical signals. These methods could be also applied to the heart from species other than rabbit with adjustments to flow rates, optics, solutions, etc.
Two optical mapping systems are described. The panoramic mapping system is used to map the entire epicardium of the rabbit heart4-7. This system provides a global view of the evolution of reentrant circuits during arrhythmogenesis and defibrillation, and has been used to study the mechanisms of arrhythmias and antiarrhythmia therapy8,9. The dual mapping system is used to map the action potential (AP) and calcium transient (CaT) simultaneously from the same field of view10-13. This approach has enhanced our understanding of the important role of calcium in the electrical alternans and the induction of arrhythmia14-16.

This article describes the basic procedures for conducting optical mapping experiments in the Langendorff-perfused rabbit heart using the panoramic imaging system, and the dual (voltage and calcium) imaging modality.

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not ...

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Stretching Short Sequences of DNA with Constant Force Axial Optical Tweezers

Single-molecule techniques for stretching DNA of contour lengths less than a kilobase are fraught with experimental difficulties. However, many interesting biological events such as histone binding and protein-mediated looping of DNA1,2, occur on this length scale. In recent years, the mechanical properties of DNA have been shown to play a significant role in fundamental cellular processes like the packaging of DNA into compact nucleosomes and chromatin fibers3,4. Clearly, it is then important to understand the mechanical properties of short stretches of DNA. In this paper, we provide a practical guide to a single-molecule optical tweezing technique that we have developed to study the mechanical behavior of DNA with contour lengths as short as a few hundred basepairs.
The major hurdle in stretching short segments of DNA is that conventional optical tweezers are generally designed to apply force in a direction lateral to the stage5,6, (see Fig. 1). In this geometry, the angle between the bead and the coverslip, to which the DNA is tethered, becomes very steep for submicron length DNA. The axial position must now be accounted for, which can be a challenge, and, since the extension drags the microsphere closer to the coverslip, steric effects are enhanced. Furthermore, as a result of the asymmetry of the microspheres, lateral extensions will generate varying levels of torque due to rotation of the microsphere within the optical trap since the direction of the reactive force changes during the extension.
Alternate methods for stretching submicron DNA run up against their own unique hurdles. For instance, a dual-beam optical trap is limited to stretching DNA of around a wavelength, at which point interference effects between the two traps and from light scattering between the microspheres begin to pose a significant problem. Replacing one of the traps with a micropipette would most likely suffer from similar challenges. While one could directly use the axial potential to stretch the DNA, an active feedback scheme would be needed to apply a constant force and the bandwidth of this will be quite limited, especially at low forces.
We circumvent these fundamental problems by directly pulling the DNA away from the coverslip by using a constant force axial optical tweezers7,8. This is achieved by trapping the bead in a linear region of the optical potential, where the optical force is constant-the strength of which can be tuned by adjusting the laser power. Trapping within the linear region also serves as an all optical force-clamp on the DNA that extends for nearly 350 nm in the axial direction. We simultaneously compensate for thermal and mechanical drift by finely adjusting the position of the stage so that a reference microsphere stuck to the coverslip remains at the same position and focus, allowing for a virtually limitless observation period.

We illustrate the use of a constant force axial optical tweezers to explore the mechanical properties of short DNA molecules. By stretching DNA axially, we minimize steric hindrances and artifacts arising in conventional lateral manipulation, allowing us to study DNA molecules as short as ~100 nm.

Single-molecule techniques for stretching DNA of contour lengths  less than a kilobase are fraught with experimental difficulties. However, many ...

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 ...

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the arterial wall lead to tissue injury, which initiates restenosis2-7. This hypothesis needs further investigations including better quantifications of non-uniform strain distribution on the artery following stent implantation. A non-contact surface strain measurement method for the stented artery is presented in this work. ARAMIS stereo optical surface strain measurement system uses two optical high speed cameras to capture the motion of each reference point, and resolve three dimensional strains over the deforming surface8,9. As a mesh stent is deployed into a latex vessel with a random contrasting pattern sprayed or drawn on its outer surface, the surface strain is recorded at every instant of the deformation. The calculated strain distributions can then be used to understand the local lesion response, validate the computational models, and formulate hypotheses for further in vivo study.

Stent-induced arterial strain distributions are characterized using an optical surface strain measurement system. This visualization technique is used to gain insights into the impact of stent implantation on the host vessel.

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the ...

In vitro Synthesis of Native, Fibrous Long Spacing and Segmental Long Spacing Collagen

Collagen fibrils are present in the extracellular matrix of animal tissue to provide structural scaffolding and mechanical strength. These native collagen fibrils have a characteristic banding periodicity of ~67 nm and are formed in vivo through the hierarchical assembly of Type I collagen monomers, which are 300 nm in length and 1.4 nm in diameter. In vitro, by varying the conditions to which the monomer building blocks are exposed, unique structures ranging in length scales up to 50 microns can be constructed, including not only native type fibrils, but also fibrous long spacing and segmental long spacing collagen. Herein, we present procedures for forming the three different collagen structures from a common commercially available collagen monomer. Using the protocols that we and others have published in the past to make these three types typically lead to mixtures of structures. In particular, unbanded fibrils were commonly found when making native collagen, and native fibrils were often present when making fibrous long spacing collagen. These new procedures have the advantage of producing the desired collagen fibril type almost exclusively. The formation of the desired structures is verified by imaging using an atomic force microscope.

In vitro Synthesis of Native, Fibrous Long Spacing and Segmental Long Spacing Collagen

Simple and reproducible procedures are described for making three structurally distinct collagen assemblies from a common commercially available Type I collagen monomer. Native type, fibrous long spacing or segmental long spacing collagen can be constructed by varying the conditions to which the 300 nm long and 1.4 nm diameter monomer building block is exposed.

Collagen fibrils are present in the  extracellular matrix of animal tissue to provide structural scaffolding and  mechanical strength. These native ...

Magnetic Tweezers for the Measurement of Twist and Torque

Single-molecule techniques make it possible to investigate the behavior of individual biological molecules in solution in real time. These techniques include so-called force spectroscopy approaches such as atomic force microscopy, optical tweezers, flow stretching, and magnetic tweezers. Amongst these approaches, magnetic tweezers have distinguished themselves by their ability to apply torque while maintaining a constant stretching force. Here, it is illustrated how such a &#8220;conventional&#8221; magnetic tweezers experimental configuration can, through a straightforward modification of its field configuration to minimize the magnitude of the transverse field, be adapted to measure the degree of twist in a biological molecule. The resulting configuration is termed the freely-orbiting magnetic tweezers. Additionally, it is shown how further modification of the field configuration can yield a transverse field with a magnitude intermediate between that of the &#8220;conventional&#8221; magnetic tweezers and the freely-orbiting magnetic tweezers, which makes it possible to directly measure the torque stored in a biological molecule. This configuration is termed the magnetic torque tweezers. The accompanying video explains in detail how the conversion of conventional magnetic tweezers into freely-orbiting magnetic tweezers and magnetic torque tweezers can be accomplished, and demonstrates the use of these techniques. These adaptations maintain all the strengths of conventional magnetic tweezers while greatly expanding the versatility of this powerful instrument.

Magnetic tweezers, a powerful single-molecule manipulation technique, can be adapted for the direct measurements of the twist (using a configuration called freely-orbiting magnetic tweezers) and torque (using a configuration termed magnetic torque tweezers) in biological macromolecules. Guidelines for performing such measurements are given, including applications to the study of DNA and associated nucleo-protein filaments.

Single-molecule techniques make it possible to investigate the behavior of individual biological molecules in solution in real time. These techniques ...

Nanomanipulation of Single RNA Molecules by Optical Tweezers

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be increasingly important. As RNA function often depends on its ability to adopt alternative structures, it is difficult to predict RNA three-dimensional structures directly from sequence. Single-molecule approaches show potentials to solve the problem of RNA structural polymorphism by monitoring molecular structures one molecule at a time. This work presents a method to precisely manipulate the folding and structure of single RNA molecules using optical tweezers. First, methods to synthesize molecules suitable for single-molecule mechanical work are described. Next, various calibration procedures to ensure the proper operations of the optical tweezers are discussed. Next, various experiments are explained. To demonstrate the utility of the technique, results of mechanically unfolding RNA hairpins and a single RNA kissing complex are used as evidence. In these examples, the nanomanipulation technique was used to study folding of each structural domain, including secondary and tertiary, independently. Lastly, the limitations and future applications of the method are discussed.

Optical tweezers have been used to study RNA folding by stretching individual molecules from their 5&#8217; and 3&#8217; ends. Here common procedures are described to synthesize RNA molecules for tweezing, calibration of the instrument, and methods to manipulate single molecules.

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be ...

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 ...