Name: use of dual optical tweezers and microfluidics for single-molecule studies
Uploaded: 2022-11-18T13:30:00
Description: Watch this Scientific Journal Video about Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies at JoVE.com

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>P...

<ol>
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H., Xie, X. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+and+ensemble+fluorescence+assays+for+a+functionally+important+conformational+change+in+T7+DNA+polymerase.">Single-molecule and ensemble fluorescence assays for a functionally important conformational change in T7 DNA polymerase.</a> Proceedings of the National Academy of Sciences of the United States of America. 104 (31), 12610-12615 (2007).</li><li>Sia, S. K., Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+devices+fabricated+in+poly(dimethylsiloxane)+for+biological+studies.">Microfluidic devices fabricated in poly(dimethylsiloxane) for biological studies.</a> Electrophoresis. 24 (21), 3563-3576 (2003).</li><li>Wuite, G. J. L., Davenport, R. 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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+imaging+of+single-molecules:+from+dynamics+of+a+single+DNA+chain+to+the+study+of+complex+DNA-protein+interactions.">Direct imaging of single-molecules: from dynamics of a single DNA chain to the study of complex DNA-protein interactions.</a> Science Progress. 84, 267-290 (2001).</li><li>Bianco, P. R., Bradfield, J. J., Castanza, L. R., Donnelly, A. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rad54+oligomers+translocate+and+cross-bridge+double-stranded+DNA+to+stimulate+synapsis.">Rad54 oligomers translocate and cross-bridge double-stranded DNA to stimulate synapsis.</a> Journal of Molecular Biology. 374 (3), 618-640 (2007).</li><li>Pezza, R. J., Camerini-Otero, R. D., Bianco, P. 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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 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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160; 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&#160; 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&#160;</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&#160; 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 &#62;3 OD units of light of wavelengths &#62;600 nm</td>
		</tr>
		<tr>
			<td>Laser safety glasses (IR)</td>
			<td>Thor labs</td>
			<td>LG11</td>
			<td>Blocks &#62;7 OD units of light of wavelengths &#179;1000 nm</td>
		</tr>
		<tr>
			<td>Mcherry&#160; 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&#160;</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>70-6000</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 thes...

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 ...

The attachment of ports to mate to syringes with tubing is critical because, when it's done right, there are no leaks, there are no bubbles in the flow cell, and experiments can be done repeatedly using the same flow cell over a period of one year with careful use. Here, two simple methods will be presented to attach the connectors to flow cells. These are easy, they're reproducible, and allow the investigator to easily adapt the flow cell to their needs without too much trouble.The use of these flow cells was done with single molecule experiments in mind. In particular, what we call visual biochemistry, but they can be used in any study where microfluidics are required and in situations where lamina fluid flow is to be used. So, to avoid mistakes, instead of using valuable flow cells to establish the techniques, I would suggest practicing on a glass slide or a cover slip first.This will enable one to get a feel for the glass or the PDMS materials to know how to handle them and how much force to apply when inserting tubing or bonding the connectors. This will require sacrificing some connectors but they're much cheaper than the flow cells. Begin by placing the flow cell on a clean flat surface.Hold the PTFE tubing three millimeters from the free end with forceps and push the tubing into the port. Repeat this procedure for each of the remaining ports. Connect the inlet ports to the syringe pump and the outlet to a waste bottle.Fill each glass syringe with one milliliter of spectro photometric, grade methanol and attach them to a switching valve. Ensure the valve has the outlet directed to waste and purge each line with 50 microliters of methanol. Switch the outlet position to the flow cell and pump 800 microliters of methanol through the flow cell at a flow rate of 100 microliters per hour to wet the surfaces and eliminate bubbles.The next day, repeat this process using 800 microliters of ultrapure water. The flow cell is now ready for use. 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. Then place the flow cell on a clean flat surface.Hold the PTFE tubing three millimeters from the free end and push the tubing into the hole in the port. Repeat this procedure for each of the remaining ports. Attach tubing from the inlets to the switching valves.Place the attachment either on a clean, flat surface or on a custom-built manifold to hold the flow cell and connectors in place while the bonding occurs. Place the flow cell on the recessed section of the manifold then place a small amount of glass glue on the bottom of the assembly and insert the seal. Position the nano port 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. 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. Place a flow cell on the microscope stage and attach the tubing using the finger tight connectors. Fill each glass syringe with one milliliter of spectro photometric grade methanol.Attach each syringe to a switching valve. Ensure the valve has the outlet directed to waste and purge each line with 50 microliters of methanol. Switch the outlet position to the flow cell and pump 800 microliters of methanol through the flow cell at a flow rate of 100 microliters per hour.The next day, repeat this process using 800 microliters of ultrapure water. Measurement of the power output is shown here. 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 100 x objective for each trap. The images demonstrate the stable optical trapping of one micrometer fluorescent beads. Here F is a fixed trap, and M is a mobile trap.With the mobile trap in the scanning mode moving through a small or large circle at 30 hertz. Fluid flow within the flow cell is laminar. The schematic shows the flow cell from the top to demonstrate that the inter channel diffusion is the main source of mixing between the adjacent fluid streams.Blue arrows indicate the flow direction 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 the adjacent fluid streams at 10 times magnification with fluorescent labeled DNA bead complexes in the lower stream and buffers only in the upper stream.The white spots in the upper stream are shot noise from the CCD camera. The flow profile and laminar flow cells is parabolic. The flow cell is viewed from the side and the direction of the flow is indicated above each cell.Linear bead velocity is affected by pump speed and sucrose concentration. The opposing force on a bead is affected by solution viscosity. Bead diameter influences the force on beads under flow.When attempting this procedure, make sure that the area you are working in is clean and that you have cleaned the glass surfaces with spectrometric grade methanol or acetone. At this stage, you should have a flow cell that is made it to your syringe pump and you are now ready to introduce your solutions and begin optical tweezer experiments. The use of flow cells, which can be used for extended periods of time, has enabled researchers to investigate a variety of single molecule reactions where either forces are measured, reactions are visualized by fluorescence or a combination of both.

use-dual-optical-tweezers-microfluidics-for-single-molecule

Research

JoVE Journal

Biochemistry

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...

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 ...

Single-Molecule Imaging of EWS-FLI1 Condensates Assembling on DNA

The fusion genes resulting from chromosomal translocation have been found in many solid tumors or leukemia. EWS-FLI1, which belongs to the FUS/EWS/TAF15 (FET) family of fusion oncoproteins, is one of the most frequently involved fusion genes in Ewing sarcoma. These FET family fusion proteins typically harbor a low-complexity domain (LCD) of FET protein at their N-terminus and a DNA-binding domain (DBD) at their C-terminus. EWS-FLI1 has been confirmed to form biomolecular condensates at its target binding loci due to LCD-LCD and LCD-DBD interactions, and these condensates can recruit RNA polymerase II to enhance gene transcription. However, how these condensates are assembled at their binding sites remains unclear. Recently, a single-molecule biophysics method-DNA Curtains-was applied to visualize these assembling processes of EWS-FLI1 condensates. Here, the detailed experimental protocol and data analysis approaches are discussed for the application of DNA Curtains in studying the biomolecular condensates assembling on target DNA.

Here, we describe the use of the single-molecule imaging method, DNA Curtains, to study the biophysical mechanism of EWS-FLI1 condensates assembling on DNA.

The fusion genes resulting from chromosomal translocation have been found in many solid tumors or leukemia. EWS-FLI1, which belongs to the FUS/EWS/TAF15 ...

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

Imaging the Motion of Fully Reconstituted Cdc45/Mcm2-7/GINS (CMG)

Hybrid Ensemble and Single-molecule Assay to Image the Motion of Fully Reconstituted CMG

Eukaryotes have one replicative helicase known as CMG, which centrally organizes and drives the replisome, and leads the way at the front of replication forks. Obtaining a deep mechanistic understanding of the dynamics of CMG is critical to elucidating how cells achieve the enormous task of efficiently and accurately replicating their entire genome once per cell cycle. Single-molecule techniques are uniquely suited to quantify the dynamics of CMG due to their unparalleled temporal and spatial resolution. Nevertheless, single-molecule studies of CMG motion have thus far relied on pre-formed CMG purified from cells as a complex, which precludes the study of the steps leading up to its activation. Here, we describe a hybrid ensemble and single-molecule assay that allowed imaging at the single-molecule level of the motion of fluorescently labeled CMG after fully reconstituting its assembly and activation from 36 different purified S. cerevisiae polypeptides. This assay relies on the double functionalization of the ends of a linear DNA substrate with two orthogonal attachment moieties, and can be adapted to study similarly complex DNA-processing mechanisms at the single-molecule level.

Author Spotlight: Investigating the Motion Dynamics of the Eukaryotic Replisome Components at the Single-Molecule Level

We report a hybrid ensemble and single-molecule assay to directly image and quantify the motion of fluorescently labeled, fully reconstituted Cdc45/Mcm2-7/GINS (CMG) helicase on linear DNA molecules held in place in an optical trap.

Eukaryotes have one replicative helicase known as CMG, which centrally organizes and drives the replisome, and leads the way at the front of replication ...

Studying TRF1/2 Interactions with Telomeric DNA Using Single-Molecule Assay

Analyzing Telomeric Protein-DNA Interactions Using Single-Molecule Magnetic Tweezers

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper function depends on tightly regulated processes of replication, elongation, and damage response. The shelterin complex, especially Telomere Repeat-binding Factor 1 (TRF1) and TRF2, plays a pivotal role in telomere protection and has emerged as a potential anti-cancer target for drug discovery. These proteins bind to the repetitive telomeric DNA motif TTAGGG, facilitating the formation of protective structures and recruitment of other telomeric proteins. Structural methods and advanced imaging techniques have provided insights into telomeric protein-DNA interactions, but probing the dynamic processes requires single-molecule approaches. Tools like magnetic tweezers, optical tweezers, and atomic force microscopy (AFM) have been employed to study telomeric protein-DNA interactions, revealing important details such as TRF2-dependent DNA distortion and telomerase catalysis. However, the preparation of single-molecule constructs with telomeric repetitive motifs continues to be a challenging task, potentially limiting the breadth of studies utilizing single-molecule mechanical methods. To address this, we developed a method to study interactions using full-length human telomeric DNA with magnetic tweezers. This protocol describes how to express and purify TRF2, prepare telomeric DNA, set up single-molecule mechanical assays, and analyze data. This detailed guide will benefit researchers in telomere biology and telomere-targeted drug discovery.

Author Spotlight: Advanced Single-Molecule Techniques for Investigating Telomeric Protein-DNA Interactions

This protocol demonstrates using single-molecule magnetic tweezers to study interactions between telomeric DNA-binding proteins (Telomere Repeat-binding Factor 1 [TRF1] and TRF2) and long telomeres extracted from human cells. It describes the preparatory steps for telomeres and telomeric repeat-binding factors, the execution of single-molecule experiments, and the data collection and analysis methods.