Research in the Bianco laboratory is supported by NIH grants GM100156 and GM144414 to P.R.B.

1. Laser trap alignment and testing with polystyrene beads
NOTE: For the setup, refer to Figure 1A,B.
CAUTION: The experimentalist should wear appropriate protective eyewear or laser safety glasses during laser beam alignment. As the optical tweezer system described herein uses both HeNe and IR beams, two separate sets of laser safety glassware are required.
<ol>
	<li>HeNe Beam alignment
	<ol>
		<li>Position all optical components on the breadboard. Align the breadboard so it is as close to a 90&#176; angle as possible relative to the microscope.</li>
		<li>Attach the mating tube containing the objective and telan lens to the microscope (Figure 1A-11,12). 
		NOTE: The telan lens is a lens system that is used to focus the image at the intermediate image plane in the eyepieces.</li>
		<li>Turn on the HeNe laser and open the shutter (Figure 1A-18). The beam should strike filter (Figure 1A-3) at a height of 54 mm reflecting 50% of the light onto mirror 5 (Figure 1A-5). The remaining 50% of the beam strikes mirror 4 (Figure 1A-4) at the same height as mirror 3.</li>
		<li>Adjust mirror 4 (Figure 1A-4) so that the beam strikes mirror 6 (Figure 1A-6) at a height of 51 mm.</li>
		<li>Adjust mirror 6 (Figure 1A-6) so that the beam is directed to the lower galvanic mirror at a height of 51 mm. The beam that exits from the upper Galvanic mirror will be at 57.5 mm.</li>
		<li>Adjust mirror 3 (Figure 1A-3) so that this beam strikes mirror 5 (Figure 1A-5) at a height of 57.5 mm. Once reflected off mirror 5, the beam should strike mirror 9 (Figure 1A-9) at the same height. The beam is effectively recombined at mirror 9, passes through the objective, and telan lens system onto mirror 21 (Figure 1A-21), which reflects it into the objective.</li>
		<li>Place a clean microscope slide onto the stage of the microscope. Turn on the camera and image the beams of the fixed and scanning beams individually on the screen. Starting with mirror 3, then 5 and 9, finally 21, perform small adjustments to position the fixed beam in the center of the screen.</li>
		<li>To ensure that the angle of the beam exiting mirror 21 is 90&#176;, alter the Z-height of the microscope and position the beam imaged on the upper and lower surfaces of the slide. When they are identical, the beam is in the correct position, which is perpendicular to the stage. Shutter this beam using shutter 19 (Figure 1A-19).</li>
		<li>Starting with mirrors 4 and 6, perform small adjustments to position the fixed beam in the center of the screen. Shutter this beam.</li>
	</ol>
	</li>
	<li>Infrared (IR) Beam alignment 
	CAUTION: Perform all steps at low laser power for safety reasons.
	<ol>
		<li>Adjust wave plate 14 (Figure 1A-14) to adjust the ratio of transmitted and reflected beams being split at the beam splitter 2 (Figure 1A-2). Beam splitter 2 reflects the s-component of the laser beam and transmits the p-component.</li>
		<li>The beam that passes through 2, strikes mirror 7 (Figure 1A-7), which deflects the beam onto the lower galvanic mirror. It should follow the path of the HeNe beam onto mirror 21. Perform small movements of mirror 7 to achieve this positioning.</li>
		<li>Adjust mirror 2 so that the beam that is reflected by mirror 2, strikes mirror 10 (Figure 1A-10) at a height of 57.5 mm.</li>
		<li>The beam is reflected from mirror 10 onto mirror 8 (Figure 1A-8) at a height of 57.5 mm. The subsequent beam should strike mirror 9 at the same height. If not, make small adjustments to mirrors 10 and 8 as required.</li>
		<li>Remove the objective and place the head of the power meter over the port in the objective turret.</li>
		<li>Use plates 14, 15, and 16 (Figure 1A-14,15,16) to adjust the power of each laser beam so that they are equal as shown in Figure 1C.</li>
		<li>Put beam expander 1 (Figure 1A-1) into place. Adjust the expanded laser beam to be circular without any coma or astigmatism.</li>
		<li>To test the optical traps, make a solution of 1 &#956;m polystyrene beads suspended in water. Then, place 10 &#956;L of this solution on a microscope slide, place a coverslip on top and seal with nail polish. Place the slide on the microscope stage and test the optical traps by trapping these 1 &#956;m beads. Ensure that the beads are sucked into the traps and held stably when the stage is moved. Trapping should occur equally efficiently from all sides of the trap.</li>
	</ol>
	</li>
</ol>
2. Microfluidic chamber preparation
<ol>
	<li>If the flow cell is a commercially constructed device made from polydimethylsiloxane (PDMS) on a coverslip, proceed to step 3. If it is a glass device with connectors attached, proceed to step 4.</li>
	<li>If the flow cell does not have connectors attached, bond them to the entry holes on the microscope slide first. See step 3 for the steps to attach connectors.</li>
</ol>
3. Flow cell connections using a PDMS flow cell
NOTE: For the setup, refer to Figure 2D.
<ol>
	<li>Place the flow cell on a clean flat surface. Hold the PTFE tubing 3 mm from the free end with forceps. Push the tubing into the preformed port (the port can support 2 bar line pressure).</li>
	<li>Repeat this procedure for each of the remaining ports. Connect the inlet ports to the syringe pump and the outlet to a waste bottle (Figure 4A,B).</li>
	<li>Fill each glass syringe with 1 mL of spectrophotometric grade methanol. Attach each syringe to a switching valve. Ensure the valve has the outlet directed to waste and purge each line with 50 &#956;L of methanol (Figure 4C, closed position).</li>
	<li>Switch the outlet position to the flow cell (Figure 4C, open position). Pump 800 &#956;L of methanol through the flow cell at a flow rate of 100 &#956;L/h to wet the surfaces and eliminate bubbles.</li>
	<li>The next day, repeat this process using 800 &#956;L of ultrapure water. The flow cell is now ready for use.</li>
</ol>
4. Attachment of press fit tubing connectors
NOTE: For the setup, refer to Figure 2F.
<ol>
	<li>If press-fit tubing connectors are to be used, carefully remove the adhesive tape from one side of the connector and place it over the hole in the microscope slide. Press down for a few seconds. Repeat the process for the remaining connectors.</li>
	<li>Place the flow cell on a clean flat surface. Hold the PTFE tubing 3 mm from the free end with forceps. Push the tubing into the preformed hole in the port and repeat this procedure for each of the remaining ports.</li>
	<li>Attach tubing from the inlets to the switching valves. Proceed to step 7.</li>
</ol>
5. Attachment of permanent assemblies
NOTE: For the setup, refer to Figure 2A-C.
<ol>
	<li>Perform the attachment either on a clean, flat surface or on a custom-built manifold to hold the flow cell and connectors in place while bonding occurs.</li>
	<li>Place the flow cell on a clean flat surface or in the recessed section of the manifold (Figure 2B). Place a small amount of glass glue on the bottom of the assembly and insert the seal.</li>
	<li>Position the nanoport over one of the entry holes on the microscope slide. Gently push down and hold in place with no lateral movement. Repeat the process for the remaining ports.</li>
	<li>Allow to dry or clamp in place in the manifold. Gently remove the flow cell from the manifold and ensure that the assemblies align well with the entry ports in the flow cell. Proceed to step 7 when ready.</li>
</ol>
6. Flow cell connections and preparation
NOTE: For the setup, refer to Figure 4A,B.
<ol>
	<li>Place a flow cell on the microscope stage. Attach the tubing using the finger-tight connectors.</li>
	<li>Fill each glass syringe with 1 mL of spectrophotometric grade methanol. Attach each syringe to a switching valve. Ensure the valve has the outlet directed to waste and purge each line with 50 &#956;L of methanol (Figure 4C, closed position).</li>
	<li>Switch the outlet position to the flow cell (Figure 4C, open position). Pump 800 &#956;L of methanol through the flow cell at a flow rate of 100 &#956;L/h to wet the surfaces and eliminate bubbles.</li>
	<li>The next day, repeat this process using 800 &#956;L of ultrapure water. The flow cell is now ready for use.</li>
</ol>
7. Control of fluid flow
<ol>
	<li>Turn the switching valves to the closed position (Figure 4C). Remove the syringes, discard the residual water, and fill the syringes with experimental solutions, for example, DNA-beads; enzyme; and ATP. Return syringes to the pump and connect them to the switching valves.</li>
	<li>In this position, manually force the plungers down 5-10 &#956;L to remove any bubbles. Change the switching valve position to Open.</li>
	<li>Use the flow rate scheme to achieve a fluid flow speed optimal for trapping and visualization as outlined in Table 1. The optimal fluid flow for optical trapping should enable stable trapping of beads and once the DNA is stretched, it should be B-form (~3,000 bp/&#956;m).</li>
</ol>
8. Trapping of polystyrene beads with single DNA molecules attached
<ol>
	<li>At 10x magnification, locate the border between fluid streams close to the channel entry points. Add oil to the 100x objective and switch to the higher magnification.</li>
	<li>Open the shutters for the optical traps (either both or one at a time). The focused laser beams should trap beads and DNA should stretch out immediately.</li>
	<li>Once a complex has been trapped, translate the stage perpendicular to the flow (Figure 3A,C, inset). This will move the trapped complex from the DNA bead solution, into stream 2. Further translation will move the complex to stream 3.</li>
</ol>
9. Flow cell cleaning
NOTE: Perform this daily.
<ol>
	<li>Turn off the pump when the experiment is complete. Turn the switching valves to the closed position (Figure 4C).</li>
	<li>Remove syringes and empty them. Rinse three times with filtered distilled water.</li>
	<li>Fill the syringes with cleaning solution. Place syringes in the pump and connect to switching valves.</li>
	<li>Purge switching valves and change the position to open. Pump 800 &#956;L of cleaning solution at a flow rate of 100 &#956;L/h typically overnight.</li>
	<li>The next day, close the switching valves, and then remove and rinse syringes with water. Rinse the flow cell and lines with 4-800 &#956;L of water at a flow rate of 400-800 &#956;L/h. If more strenuous cleaning of flow cells is required, replace the cleaning solution with 6 M guanidium hydrochloride.</li>
</ol>

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The author declares no conflicts of interest.

The careful assembly of the flow system is critical to the successful outcome of experiments4,6. One of the most challenging aspects of the protocol is the attachment of connectors to the glass surface. For this, we use the following two approaches: press-fit fit tubing connectors and nanoport assemblies. Press-fit connectors adhere easily to glass followed by pushing of PTFE tubing into the preformed holes using forceps. When a more stable attachment is required, the bonding of nanoport assemblies is preferred. With careful use and cleaning, a single glass flow cell with permanent assemblies can be used for more than 1 year. If flow cells need to be switched out more frequently, PDMS flow cells can be used as they are an excellent and cheaper alternative. They can be reused but PDMS is susceptible to swelling, which can alter the fluid flow behavior over time.
Both glass and PDMS flow cells are not indestructible. In the laboratory, the maximum flow rate used is 800 &#956;L/h. If this is exceeded daily and/or for extended periods of time, flow cell warping&#160;(glass flow cells) and distortion of PDMS can occur. While the initial wetting procedure with methanol and water takes time, it is well worth it as all bubbles are eliminated, and also leaks at connectors can be detected. The flow cells described here provide flexibility to the experiments being done. Using these approaches of different flow cells, tubing and attachment of different connectors, experimental conditions can be readily adapted and quickly optimized to ensure optimal data are obtained. Y-shaped channel flow cells are presented but a large range of microfluidic device designs are available from multiple companies both without and with connectors attached. This provides the investigator even greater flexibility in experimental design.
The use of four-way switching valves is a simple and effective method that ensures that once the flow system is assembled and tested, it effectively becomes a closed environment, where contamination is minimal, and bubbles are eliminated. The latter is critical as bubbles tend to compress and decompress randomly during an experiment and this perturbs fluid flow causing trapped DNA molecules to move in unexpected ways and even break. Great care should be taken to ensure they are prevented from forming in the system.
Finally, the system described here is optimized for visual biochemistry where careful analysis of DNA, beads attached to proteins or fluorescent-tagged proteins can be performed. Although the system is not designed to do force measurements with the dual traps and addition of quadrant photodetectors, forces can be measured by attaching beads of different diameter to DNA motor proteins, for example, and asking the motor to pull this bead against fluid flow. By varying solution viscosity, fluid flow, and bead diameter, a large range of opposing forces can be applied to provide detailed insight into motor protein function (Figure 5). This is a simple and inexpensive way to measure forces associated with biological transactions, but it lacks the precision associated with more sensitive approaches. When the visual biochemical approach is combined with sensitive force detection methods, more complete pictures of biochemical reactions can be provided.

The ability to visualize protein-DNA interactions at the single-molecule level and in real-time has provided significant insight into genome stability1,2. In addition to working with single molecules of DNA one at a time, the ability to view transactions between individual molecules nearby provides additional insight3,4,5. The manipulation of additional DNA molecules requires both additional optical traps as well as high-quality, multi-channel, microfluidic flow cells6.
There are several methods available to generate more than one optical trap. These include galvanometer scanning mirrors, acoustic optic modulators, and diffractive optics, which generate holographic optical tweezers4,7,8,9. Often, scanning mirrors and acoustic optic modulators produce traps that timeshare. In the setup described here, the beam of a single Nd:YAG laser is split on polarization, and then galvanometer laser scanning mirrors control the position of what is termed the mobile trap (Figure 1)4. To facilitate the positioning of mirrors and filters to direct the trapping beam to the back aperture of the microscope objective, a HeNe laser is used. This makes overall alignment easier as the HeNe beam is visible to the naked eye, whereas infrared beams are not. The HeNe beam is also safer to work with making the positioning of mirrors and other components less stressful. Initially, the beam path for this laser is separate from the 1064 nm beam, but is introduced into the same beam path, and then into the microscope objective. Once physical alignment is achieved, positioning the 1064 nm beam on top of the HeNe beam is done and this is facilitated by the use of an infrared viewer and various beam imaging tools to visualize beam position and quality. Then, the beam expander is introduced, and the resulting expanded infrared beam is aligned onto the back aperture of the objective. Finally, the objective is removed and the power in each polarized beam is measured and adjusted using &#955;/2 waveplates to be equal (Figure 1C). Power measurements are also done once the objective is returned and typically there is a 53% power loss. There is, however, sufficient power to form stable fixed and moving optical traps in the focal plane (Figure 1D).
To image DNA transactions, microfluidic flow cells play a key role as they permit controlled measurements at the single-molecule level with high spatial and temporal resolution (Figure 2). The term microfluidic refers to the ability to manipulate fluids in one or more channels with dimensions ranging from 5-500 &#956;m10,11. The term stream refers to the actual fluid within a channel and the channel refers to the physical channel in which a fluid stream or streams move. Single-channel flow cell design has a common, physical channel where reactions are observed and there is typically only one fluid stream present. Thus, these designs are known as single-stream flow cells. In contrast, multi-stream flow cells are defined as a microfluidic device in which two or more entry channels converge into a single, common, physical channel (Figure 2A). Within the common channel, the fluid streams that originate from the individual channels flow parallel to one another, and remain separated with only minimal mixing between them occurring due to diffusion (Figure 2B). In the majority of experimental setups, a single pump pushes fluids into each channel at the same speed. In contrast, when boundary steering is used, three or more, independently controlled pumps push fluids through the channels. However, each pump operates at a different speed but the net flow rate in the common channel is constant12. This&#160;permits the rapid exchange of main channel components simply by altering pump speed.
In addition to laminar flow, another critical factor is the parabolic velocity profile within the laminar fluid stream. The highest flow velocity occurs in the middle of the stream and the slowest occurs&#160;next to the surfaces (Figure 2C)13. This profile must be considered to fully stretch a DNA molecule that is attached to a bead held in the stream for precise visualization of fluorescent DNA and accurate single-molecule analyses. Here, the DNA is stretched to B-form and is held in place under 0 pN of force. To achieve this, focusing of the optical trap position should be to a position 10-20 &#956;m from the bottom coverslip surface (Figure 2D). Care must be taken so that the DNA molecule is not stretched beyond B-form as this can inhibit enzyme reactions. Under typical buffer conditions, 1 &#956;m = 3,000 bp of DNA14. Furthermore, by trapping 10-20 &#956;m from the coverslip, the DNA complex is positioned far from the surface thereby minimizing surface interactions.
Many methods have been used to create microfluidic device channels and these can be done in the laboratory or flow cells can be purchased from commercial sources6,15,16,17. The optimal materials used to construct the flow cells must be mechanically rigid, optically transparent with low fluorescence, and impervious to organic solvents6. Frequently, borosilicate float glass, or fused silica are used to provide a stable flow environment for an extended time that is suitable for optical trapping, visualization, and force detection. These materials also permit the use of non-aqueous solvents (e.g., spectrophotometric grade methanol) to simplify surface wetting and removal of air bubbles, and denaturants (e.g., 6M guanidinium hydrochloride) or detergents to clean the flow cell. Finally, the methods used to introduce fluids into flow cells vary from complex vacuum pump systems to single syringe pumps14,18,19,20,21,22,23,24,25,26,27. In the approach described here, a syringe pump that can accommodate up to 10 syringes is used (Figure 3A). This provides flexibility to use single-channel flow cells or flow cells with multiple inlet channels. Here, a three-channel flow cell is used and is mated to syringes held in place on the syringe pump using poly-ether-ether-ketone (PEEK) tubing (Figure 3A-C). The flow of fluids is controlled by four-way switching valves and thus serve to minimize the introduction of bubbles into the flow cell (Figure 3A,D). In addition, Hamilton gastight syringes which have stiff glass walls and polytetrafluoroethylene (PTFE)-coated plungers, are recommended as they provide exceptionally smooth plunger motion that is essential to obtaining a smooth flow14,27.
In the experimental system described, flow cells with two to five inlet channels have been used. The number of inlet channels is dictated by the experiment being done. For the study of RecBCD and Hop2-Mnd1, two stream channels were sufficient14,28. For the helicase, the enzyme was bound to the free end of the DNA and translated into a stream containing magnesium and ATP to initiate translocation and unwinding. For Hop2-Mnd1, optically trapped DNA was translated into the adjacent fluid stream containing proteins and buffer &#177; divalent metal ions. The use of three-channel flow cells enables one to trap DNA in stream 1, translate the DNA into stream 2 to allow protein binding to occur, and then to stream 3 where ATP is present, for example, to initiate reactions. A variation on the above is to use fluorescent-tagged protein in channel 2, which results in the fluid stream being completely white and precludes visualization of the DNA. When this molecule is translated into stream 3, both proteins and DNA are now visible when reactions are initiated.
The use of four-way switching valves to control fluid flow is a critical component of the system to eliminate bubbles in the flow cells. Bubbles are detrimental to stable fluid flow as they contract and expand in unpredictable ways resulting in rapid changes in flow velocity and the introduction of turbulence. When the valves are positioned between syringes and inlet tubing, the flow path is disconnected by switching the valve position when syringes are changed out. When the new syringe is put in place, the plunger can be manually depressed so that &#62;6 &#956;L is ejected (the dead volume of the valve) and this eliminates bubbles almost entirely.
The attachment of connectors to flow cells is frequently the rate-limiting step in flow cell use. We describe the use of two types of connectors: removable known as press-fit and permanent ones (nanoport assemblies). The removable connectors are simple to adhere to a flow cell and different types of flexible tubing in addition to the recommended PTFE can be tested with these connectors. This is a rapid and cost-effective way to test tubing and connectors without sacrificing more expensive glass flow cells. In contrast, nanoport assemblies are attached permanently, withstand pressures up to 1,000 psi, and, in our hands, their use is restricted to PEEK tubing of different diameters. This is not a disadvantage as PEEK tubing is preferably used. A single glass flow cell with permanent assemblies attached can be reused for more than 1 year with careful use.

<table><tbody><tr><td>100x objective</td><td>Leica</td><td>506318 or 506038</td><td>Oil immersion lenses; Imaging and optical trapping only; Plan APO objectives optimized for fluorescence imaging</td></tr><tr><td>10X Objective</td><td>Leica</td><td>506263</td><td>Used to locate laser beams spots during alignment; to find focus and X-Y position in flow cell</td></tr><tr><td>1 mm fluorescent beads</td><td>Bangs Labs</td><td>FSDG004</td><td>Used for tap performance, focal position determination</td></tr><tr><td>1 mm polystyrene beads</td><td>Bangs Labs</td><td>CPO1004</td><td>Used for trap performance evaluation and binding to biotinylated molecules</td></tr><tr><td>63x objective</td><td>Leica</td><td>506081</td><td>Used to locate laser beams spots during alignment and to find focus and X-y position in flow cell; can be used for optical trapping as it has an identical back aperture diameter to the 100X; oil immersion lens</td></tr><tr><td>Alignment laser</td><td>Lumentum</td><td>1100 series</td><td>10mW HeNe laser that is visible to the naked eye that is used to position optics</td></tr><tr><td>Beam alignment camera</td><td>Amscope</td><td>MU303</td><td>A simple, inexpensive and software controlled camera for imaging of the beam position</td></tr><tr><td>Camera control and Image capture software</td><td>Hamamatsu</td><td>HCImage</td><td>Coordinates activities of the Lambda DG4 with the camera to facilitate rapid wavelength switching</td></tr><tr><td>Camera; Orca flash 4</td><td>Hamamatsu</td><td>c13440-20cu</td><td>CCD camera for imaging of single-molecule experiments</td></tr><tr><td>C-mount for the beam alignment camera</td><td>Spot imaging solutions</td><td>DE50CMT</td><td>Provides optimal positioning of the camera for imaging of laser beams during alignment</td></tr><tr><td>C-mount for the Orca Flash 4 camera</td><td></td><td></td><td>Has a retainer ring to hold an IR blocking filter in place. This eliminates reflected IR beam from the optical traps and facilitates clearer imaging of trapped objects.</td></tr><tr><td>Cy5&amp;nbsp; fluorescence filter cube</td><td>Semrock</td><td>cy5-404a-lsc-zero</td><td>Used in conjunction with Lambda DG4 to image Cy5 only</td></tr><tr><td>Fitc-Txred&amp;nbsp; fluorescence filter cube</td><td>Semrock</td><td>fitc/txred-2x-b-000</td><td>Used in conjunction with Lambda DG4 to image FITC and TXRed</td></tr><tr><td>Fluidics tubing</td><td>Grace Bio</td><td>46004</td><td>PTFE tubing as an alternate to PEEK; works well on some flow cells. Can be used with PDMS flow cells or glass flow cells when Grace Bio fit tubing connectors are used</td></tr><tr><td>GFP fluorescence filter cube</td><td>Semrock</td><td>gfp-3035b-lsc-zero</td><td>Used in conjunction with Lambda DG4 to image GFP only</td></tr><tr><td>Glass flow cells</td><td>Translume</td><td>Custom</td><td>Clear flow channels for imaging (Fig. 2E)</td></tr><tr><td>Glass glue</td><td>Loctite</td><td>233841</td><td>Securely and easily bonds Nanoport assemblies to glass flow cells</td></tr><tr><td>Glass/PDMS sandwich flow cells</td><td>CIDRA Precision services</td><td>Custom design</td><td>Flow cells built according to your specifications; imaging channels are clear (Fig. 2C)</td></tr><tr><td>Hamilton Cleaning solution</td><td>Hamilton</td><td>18311</td><td>Gentle but efficient cleaning solution for glass flow cells; does not bubble when used carefully</td></tr><tr><td>Illumination system</td><td>Sutter Instrument</td><td>Lamda DG4</td><td>Discontinued so recommend Lambda 721</td></tr><tr><td>Illumination system</td><td>Sutter Instrument</td><td>Lamda DG4</td><td>Discontinued so recommend Lambda 721</td></tr><tr><td>Image analysis software</td><td>Media cybernetics</td><td>Image Pro Premiere</td><td>Analysis of images and single molecule tracking</td></tr><tr><td>Image analysis software</td><td>Fiji/NIH Image/Image J</td><td>Shareware</td><td>Analysis of images and single molecule tracking</td></tr><tr><td>Image display card</td><td>Melles Griot</td><td>06 DLA 001</td><td>Alternate product from Thorlabs: VRC5</td></tr><tr><td>Immersion oil</td><td>Zeiss</td><td>444960</td><td>Immersol 518 F fluorescence free</td></tr><tr><td>Laser beam alignment tools</td><td>Thor labs</td><td>FMP05/M; dgo5-1500-h1; BHM1&amp;nbsp;</td><td>Used to ensure beams are horizontal and at the correct height</td></tr><tr><td>Laser beam viewer</td><td>Canadian Photonics labs</td><td>IR 3150</td><td>Used to image IR beam spots on mirrors and&amp;nbsp; targets</td></tr><tr><td>Laser power meter</td><td>Thor labs</td><td></td><td>Measurement of laser output as well as trap strength</td></tr><tr><td>Laser safety glasses (HeNe)</td><td>Thor labs</td><td>LG7 or 8</td><td>Blocks &gt;3 OD units of light of wavelengths &gt;600 nm</td></tr><tr><td>Laser safety glasses (IR)</td><td>Thor labs</td><td>LG11</td><td>Blocks &gt;7 OD units of light of wavelengths &amp;sup3;1000 nm</td></tr><tr><td>Mcherry&amp;nbsp; fluorescence filter cube</td><td>Semrock</td><td>mcherry-a-lsc-zero</td><td>Used in conjunction with Lambda DG4 to image mcherry only</td></tr><tr><td>Microscope</td><td>Leica</td><td>DMIRE2</td><td>DIC port removed to accommodate Dichroic trapping/alignment mirror</td></tr><tr><td>Microscope control software&amp;nbsp;</td><td>UCSF/shareware</td><td>uManager</td><td>Controls the microscope, permits focal alignment of objectives as well as stage control</td></tr><tr><td>Nanoport assembly</td><td>IDEX</td><td>N333</td><td>Connectors that are bonded to flow cells</td></tr><tr><td>Optical table support</td><td>Thor Labs</td><td>PA52502</td><td>Active isolation table support</td></tr><tr><td>Optics and lenses</td><td>Solar TII</td><td>Various</td><td>Interference mirrors, telescopes and lenses custom designed for the system</td></tr><tr><td>PDMS flow cells</td><td>ufluidix</td><td>Custom</td><td>Flow cells built according to your specifications; imaging channels are clear (Figs. 2B and D)</td></tr><tr><td>PEEK tubing</td><td>IDEX</td><td>1532</td><td>Provides excellent connection to flow cells and switching valves</td></tr><tr><td>Pinkel fluorescence filter cube</td><td>Semrock</td><td>lf488/543/635-3x-a-000</td><td>Used in conjunction with Lambda DG4 to image multiple fluorophores rapidly</td></tr><tr><td>Press fit tubing connectors</td><td>GraceBio</td><td>46003</td><td>Clear silicone connector with adhesive that binds well to glass</td></tr><tr><td>Scanning mirrors</td><td>GSI Lumonics</td><td>VM500</td><td>Used to provide control of the second optical trap. GSI Lumonics no longer exists. Similar mirrors can be purchased from Cambridge Scientific</td></tr><tr><td>Stage</td><td>Leica</td><td></td><td></td></tr><tr><td>Stage micrometer</td><td>Electron Microscopy Sciences</td><td>68042-08</td><td>Provides on screen ruler for positioning of the beam and system calibration</td></tr><tr><td>Switching valves</td><td>IDEX</td><td>V-101T</td><td>Control direction of fluid flow and eliminate introduction of bubbles into flow cells</td></tr><tr><td>Syringe and valve manifold</td><td>Machine shop</td><td>None</td><td>Custom built</td></tr><tr><td>Syringe pump</td><td>Harvard Apparatus</td><td>PHD 2000</td><td>Controls fluid flow through flow cells</td></tr><tr><td>Syringe pump software</td><td>Harvard Apparatus</td><td>&lt;strong&gt;70-6000&lt;/strong&gt;</td><td>Flow control provides seamless, programmable control of fluid flow</td></tr><tr><td>Syringes</td><td>Hamilton</td><td>81320</td><td>Gas-tight, PTFE Luer Lock, glass barrels with Teflon-coated plungers</td></tr><tr><td>Table top</td><td>Thor Labs</td><td>T36H</td><td>Optical table top or breadboard</td></tr><tr><td>Trapping laser</td><td>Newport/Spectra Physics</td><td>J-series; BL106C</td><td>Nd:YAG laser; 1064 nm; 5W laser</td></tr></tbody></table>

use of dual optical tweezers and microfluidics for single-molecule studies

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the averaging that takes place in bulk-phase studies. To achieve visualization, dual optical tweezers, where one trap is fixed and the other is mobile, are focused into one channel of a multi-stream microfluidic chamber positioned on the stage of an inverted fluorescence microscope. The optical tweezers trap single molecules of fluorescently labeled DNA and fluid flow through the chamber and past the trapped beads, stretches the DNA to B-form (under minimal force, i.e., 0 pN) with the nucleic acid being observed as a white string against a black background. DNA molecules are moved from one stream to the next, by translating the stage perpendicular to the flow to enable the initiation of reactions in a controlled manner. To achieve success, microfluidic devices with optically clear channels are mated to glass syringes held in place in a syringe pump. Optimal results use connectors permanently bonded to the flow cell, tubing that is mechanically rigid and chemically resistant, and which is connected to switching valves that eliminate bubbles that prohibit laminar flow.

The initial testing of the trap alignment and strength is done with 1 &#956;m, non-fluorescent polystyrene beads. Since most of the research done in the laboratory uses fluorescence, we further test trap strength using 1 &#956;m, Dragon green polystyrene beads (Figure 1D,E). Thereafter work changes to optical trapping of DNA-bead complexes where the DNA is stained with the bis-intercalating dye YOYO-114,29. When these complexes are trapped 10-15 &#956;m above the coverslip surface in a microfluidic flow cell, fluid flow past the bead stretches out the DNA (Figures 3B,C). It is important to measure the length of the fluid-stretched DNA to ensure that it is stretched to its B-form length, which is the expected length. For example, for bacteriophage lambda DNA, the full length is slightly longer than 16 &#956;m, which corresponds to 48,504 bp14.
To characterize fluid flow, fluorescent beads were introduced into flow cells with a focus height of 15 &#956;m above the coverslip surface using different pump speeds and in solutions of different viscosity. Movies of bead position were captured, and bead position was tracked using image analysis software. The results show that for 1 &#956;m diameter beads, linear velocity can be accurately tracked at a pump flow rate of 100 &#956;L/h in sucrose solutions ranging from 10%-50% (Figure 5A). Under similar conditions, the force on the bead can be calculated and found to range from 10 to 400 pN, depending on the pump flow rate and solution viscosity (Figure 5B). Finally, the effect of bead diameter ranging from 120 to 970 nm on force were evaluated in different concentrations of sucrose. The data reveal that forces ranging from 1-27 pN could be measured by attaching beads of different diameter to a motor protein and asking the motor to pull that bead against the fluid flow (Figure 5C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64023/64023fig01.jpg" /> 
Figure 1: Optical layout and demonstration of trapping activity. (A) Schematic of the optical layout with the IR beam in red and HeNe beam in black. (B) Photograph of optical components mounted on a breadboard. The component numbers in panels A and B are as follows: 1), Beam expander; 2), Interference mirror (Rsmax = 1064 nm; Tpmax = 1064 nm); 3), Interference mirrors (633 nm; R = 50%; T = 50%); 4-6), Interference mirrors (Rmax = 633 nm); 7), Interference mirror (Rmax = 1064 nm; Tmax= 633 nm); 8), Interference mirror (Rsmax = 1064 nm; Tpmax = 1064 nm, Tmax = 633 nm); 9), Interference mirror (Rmax = 1064 nm, 633 nm; Tmax = 1064 nm, 633 nm); 10), Interference mirror (Rmax = 1064 nm); 11), Objective lens, f = 110 nm; 12), Telan lens; 13), Galvanic mirrors; 14-16), Rotators (/2); 17), IR scanning channel shutter; 18), Visible channel shutter; 19), IR Fixed channel shutter; 20), Beam stops and 21), Interference mirror (Rmax = 1064 nm; Tmax = 633 nm). (C) Measurement of power output. The power output from the laser is measured before installing the laser head into the optical layout. Once trap alignment is done, the beam power is measured after the 100x objective for each trap. (D) and (E) Images demonstrating stable, optical trapping of 1 &#956;m fluorescent beads. F, fixed trap and M, mobile trap with the mobile trap in scanning mode moving through a small (D) or large (E) circle at 30 mHz. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64023/64023fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64023/64023fig02.jpg" /> 
Figure 2: Microfluidic flow cells. (A) Assembly of a three-channel flow cell made from borosilicate glass. (B) A manifold to stably position flow cells during connector bonding. Top, the base of the aluminum manifold with a 1 mm recessed section to laterally position the flow cell. The four posts (one in each corner) guide the lid of the manifold into position, while the more centrally placed threaded bolts are used to clamp the lid in place. Bottom, manifold lid with drilled holes to match the posts and bolts on the manifold base. (1), nuts that attach to the manifold base bolts; (2) springs that are placed between the nuts and the manifold lid to provide light tension during the bonding process. (C) A close-up view of the convergence of the inlet channels into a common single fluid stream in a glass flow cell with connectors already attached. A quarter is placed next to the flow cell as a reference. (D) An image of the PDMS flow cell. The PDMS layers is 3 mm thick. (E) A fused silica flow cell with luer connectors. (F) A borosilicate glass flow cell with press-fit connectors in place. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64023/64023fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64023/64023fig03.jpg" /> 
Figure 3: Fluid flow characteristics through a glass flow cell. (A) Fluid flow within the flow cell is laminar. The schematic shows the flow cell from the top to demonstrate that inter-channel diffusion is the main source of mixing between adjacent fluid streams. The flow direction is indicated by blue arrows and the individual streams are colored white, light gray, and white. The widening regions of transverse diffusion between channels are indicated in red. The inset shows a fluorescence image of adjacent fluid streams at 10x magnification with fluorescent-labeled DNA-bead complexes in the lower stream and buffer only in the upper stream. The white spots in the upper stream are shot noise from the CCD camera. (B) The flow profile (light purple) in laminar flow cells is parabolic. The flow cell is viewed from the side and the direction of the flow is indicated above each cell. The fastest fluid velocities occur in the flow cell located in the center while the slowest fluid velocities occur next to the flow cell surfaces. Consequently, for an optically trapped DNA-bead complex positioned at the center of the flow cell, the fluid flow stretches the bacteriophage&#160;&#955; DNA molecule (48,504 bp) to 16 &#956;m (B-form) so that clear visualization can be made (left image). In contrast, the same molecule positioned near the surface of the flow cell experience low flow rates and cannot be stretched out. (C) A single DNA bead complex is positioned within a flow cell. Different depth flow cells can be used but, in each case, the beam alignments can be set so that optimal trapping occurs 10-20 &#956;m above the coverslip surface, which eliminates surface effects and has stable flow so that DNA molecules can be stretched to B-form. As indicated in the left image in (C), the DNA is stretched to enable clear visualization. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64023/64023fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64023/64023fig04.jpg" /> 
Figure 4: Flow and reaction control of optically trapped DNA within a flow cell in the microscope stage. (A) Mating of the flow cell to a syringe pump. A photograph of a three-channel flow cell connected to a syringe pump via three-way switching valves is presented. The flow cell is held in place on a motorized stage in an inverted microscope. The flow cell is connected to a waste line (left) and switching valves (dashed cyan circle) via orange, 500 &#181;m tubing (indicated with green tape). The valves are stably mounted onto a custom-built manifold adapted to the syringe pump apparatus, which houses 1 mL gastight syringes (yellow arrows demarcate plungers). The waste lines are used to remove air bubbles that can be trapped in the switching valve during syringe changes. (B) Close-up view of the three-channel flow cell. To facilitate visualization of fluid streams, the outer ones have been colored with food coloring with the central stream containing water only. The zoomed region indicates how an optically trapped bead-DNA molecule complex is translated from one fluid stream to the next (blue arrow) by translating the stage perpendicular to the flow (black arrow). (C) Schematics of the three-way switching valves indicating their operational positions. The letter designations F, S, and W correspond to the direction fluid and can flow to the flow cell, syringe, and waste, respectively. One port is permanently closed as indicated by the black plug on the top of each schematic. In the Open or operational position (left panel), fluid flow passes directly through the switching valve from the syringe (S) to the flow cell (F). In the Closed position (right panel), the dial is turned so that the flow cell is sealed from the environment. In this position, the flow is directed to waste and syringes can be switched out without introducing air bubbles into the flow cell. As the valves have a 6 &#956;L dead volume, very little solution is wasted when purging lines before reverting to the Open configuration. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64023/64023fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64023/64023fig05.jpg" /> 
Figure 5: Bead motion and opposing force are affected by fluid flow. (A) Linear bead velocity is affected by pump speed and sucrose concentration. (B) The opposing force on a bead is affected by solution viscosity. (C) Bead diameter influences the force on beads under flow. In all experiments, fluorescent beads were used (Dragon green). In (A) and (B), 0.97 &#181;m diameter beads were used and in (C), beads of different diameter were used. Sucrose solutions were made in ultrapure water, filtered through 0.2 &#956;m filters and refractive index measured using a handheld refractometer, with viscosity determined in centipoise using ISCO Tables and finally converted to SI units in mPa.s. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64023/64023fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Flow rate (&#956;L/h)b</td>
			<td>Time per step (min)</td>
			<td>Volume dispensed (&#956;L)</td>
		</tr>
		<tr>
			<td>800</td>
			<td>5</td>
			<td>67</td>
		</tr>
		<tr>
			<td>400</td>
			<td>5</td>
			<td>33</td>
		</tr>
		<tr>
			<td>200</td>
			<td>5</td>
			<td>17</td>
		</tr>
		<tr>
			<td>100</td>
			<td>5</td>
			<td>8</td>
		</tr>
		<tr>
			<td>50</td>
			<td>5</td>
			<td>4</td>
		</tr>
		<tr>
			<td>25</td>
			<td>5</td>
			<td>2</td>
		</tr>
		<tr>
			<td>15</td>
			<td>5 or more</td>
			<td>1 or more</td>
		</tr>
		<tr>
			<td colspan="3">a. These are fluid flow rates calculated by the syringe pump software and are unique for each syringe diameter used. The flow rates enable rapid filling of the lines and flow chamber (800 &#956;l/hr), followed by a gradual decrease to the ideal flow rate for trapping and visualization</td>
		</tr>
		<tr>
			<td colspan="3">b. The syringe pump can be controlled from the keypad or using software provided by the vendor.</td>
		</tr>
	</tbody>
</table>
Table 1: Fluid flow rates to minimize bubbles and achieve stable trapping.

Watch this Scientific Journal Video about Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies at JoVE.com

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the ...

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2.1K Views.  University of Nebraska Medical Center. Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Video: Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

A Microfluidic-based Hydrodynamic Trap for Single Particles

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science. Methods for particle trapping based on optical, magnetic, electrokinetic, and acoustic techniques have led to major advancements in physics and biology ranging from the molecular to cellular level. In this article, we introduce a new microfluidic-based technique for particle trapping and manipulation based solely on hydrodynamic fluid flow. Using this method, we demonstrate trapping of micro- and nano-scale particles in aqueous solutions for long time scales. The hydrodynamic trap consists of an integrated microfluidic device with a cross-slot channel geometry where two opposing laminar streams converge, thereby generating a planar extensional flow with a fluid stagnation point (zero-velocity point). In this device, particles are confined at the trap center by active control of the flow field to maintain particle position at the fluid stagnation point. In this manner, particles are effectively trapped in free solution using a feedback control algorithm implemented with a custom-built LabVIEW code. The control algorithm consists of image acquisition for a particle in the microfluidic device, followed by particle tracking, determination of particle centroid position, and active adjustment of fluid flow by regulating the pressure applied to an on-chip pneumatic valve using a pressure regulator. In this way, the on-chip dynamic metering valve functions to regulate the relative flow rates in the outlet channels, thereby enabling fine-scale control of stagnation point position and particle trapping. The microfluidic-based hydrodynamic trap exhibits several advantages as a method for particle trapping. Hydrodynamic trapping is possible for any arbitrary particle without specific requirements on the physical or chemical properties of the trapped object. In addition, hydrodynamic trapping enables confinement of a "single" target object in concentrated or crowded particle suspensions, which is difficult using alternative force field-based trapping methods. The hydrodynamic trap is user-friendly, straightforward to implement and may be added to existing microfluidic devices to facilitate trapping and long-time analysis of particles. Overall, the hydrodynamic trap is a new platform for confinement, micromanipulation, and observation of particles without surface immobilization and eliminates the need for potentially perturbative optical, magnetic, and electric fields in the free-solution trapping of small particles.

In this article, we present a microfluidic-based method for particle confinement based on hydrodynamic flow. We demonstrate stable particle trapping at a fluid stagnation point using a feedback control mechanism, thereby enabling confinement and micromanipulation of arbitrary particles in an integrated microdevice.

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science.  Methods for ...

Bioengineering

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 ...

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 ...

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to understand morphogenetic processes is to directly measure cellular forces and mechanical properties in vivo during embryogenesis. Here, we present a setup of optical tweezers coupled to a light sheet microscope, which allows to directly apply forces on cell-cell contacts of the early Drosophila embryo, while imaging at a speed of several frames per second. This technique has the advantage that it does not require the injection of beads into the embryo, usually used as intermediate probes on which optical forces are exerted. We detail step by step the implementation of the setup, and propose tools to extract mechanical information from the experiments. By monitoring the displacements of cell-cell contacts in real time, one can perform tension measurements and investigate cell contacts&#39; rheology.

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

We present a setup of optical tweezers coupled to a light sheet microscope, and its implementation to probe cell mechanics without beads in the Drosophila embryo.

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to ...

Developmental Biology

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

RNA adopts diverse structural folds, which are essential for its functions and thereby can impact diverse processes in the cell. In addition, the structure and function of an RNA can be modulated by various trans-acting factors, such as proteins, metabolites or other RNAs. Frameshifting RNA molecules, for instance, are regulatory RNAs located in coding regions, which direct translating ribosomes into an alternative open reading frame, and thereby act as gene switches. They may also adopt different folds after binding to proteins or other trans-factors. To dissect the role of RNA-binding proteins in translation and how they modulate RNA structure and stability, it is crucial to study the interplay and mechanical features of these RNA-protein complexes simultaneously.&#160;This work illustrates how to employ single-molecule-fluorescence-coupled optical tweezers to explore the conformational and thermodynamic landscape of RNA-protein complexes at a high resolution. As an example, the interaction of the SARS-CoV-2 programmed ribosomal frameshifting element with the trans-acting factor short isoform of zinc-finger antiviral protein is elaborated. In addition, fluorescence-labeled ribosomes were monitored using the confocal unit, which would ultimately enable the study of translation elongation. The fluorescence coupled OT assay can be widely applied to explore diverse RNA-protein complexes or trans-acting factors regulating translation and could facilitate studies of RNA-based gene regulation.

This protocol presents a complete experimental workflow for studying RNA-protein interactions using optical tweezers. Several possible experimental setups are outlined including the combination of optical tweezers with confocal microscopy.

RNA adopts diverse structural folds, which are essential for its functions and thereby can impact diverse processes in the cell. In addition, the ...

Magnetic Tweezers in a Microplate Format

Mechanobiology describes how the physical forces and mechanical properties of biological material contribute to physiology and disease. Typically, these approaches are limited single-molecule methods, which restricts their availability. To address this need, a microplate assay was developed that enables mechanical manipulation while performing standard biochemical assays. This is achieved using magnets incorporated into a microplate lid to create multiple magnetic tweezers. In this format, force is exerted across biomolecules connected to paramagnetic beads, equivalent to a typical magnetic tweezer. The study demonstrates the application of this tool with FRET-based assays to monitor protein conformations. However, this approach is widely applicable to different biological systems ranging from measuring enzymatic activity through to the activation of signaling pathways in live cells.

Here, we describe the use of a novel microplate assay to enable mechanical manipulation of biomolecules while performing ensemble biochemical assays. This is achieved using a microplate lid modified with magnets to create multiple static magnetic tweezers across the microplate.

Mechanobiology describes how the physical forces and mechanical properties of biological material contribute to physiology and disease. Typically, these ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

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 ...

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

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Rozdziały w tym wideo

A Microfluidic-based Hydrodynamic Trap for Single Particles

Stretching Short Sequences of DNA with Constant Force Axial Optical Tweezers

Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

Magnetic Tweezers for the Measurement of Twist and Torque

Nanomanipulation of Single RNA Molecules by Optical Tweezers

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

Magnetic Tweezers in a Microplate Format

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

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