This work was supported in part by a National Science Foundation grant DMS-1463234, National Institutes of Health grants HL082808 and AI133634, and Lehigh University internal funding.

1. Preparing VWF
<ol>
	<li>Reconstitute human plasma VWF to prepare it for the labeling reactions. Add 100 &#181;L of deionized (DI) water to 100 &#181;g of lyophilized VWF to create a 1 mg/mL VWF stock solution.</li>
	<li>Dialyze VWF stock solution in order to remove excess glycine, thereby increasing the biotin and fluorophore labeling efficiency.
	<ol>
		<li>Transfer 50 &#181;L of VWF stock solution into a 0.1 mL dialysis unit with a 10,000 molecular weight cut-off and seal with a cap. Store the remaining stock solution at -20 &#176;C. VWF stock will be stable for up to 1 year at -20 &#176;C.</li>
		<li>Run dialysis in 500 mL of 1x sterile phosphate-buffered saline (PBS) (0.01 M disodium phosphate, 0.0018 monopotassium phosphate, 0.0027 M potassium chloride, 0.137 M sodium chloride, pH 7.4 at 25 &#176;C) for 1 h at 4 &#176;C with slow stirring. Repeat dialysis for an additional hour using 500 mL of fresh PBS.</li>
	</ol>
	</li>
	<li>Start biotin labeling reaction. Prepare a 2 mM solution of NHS-PEG4-biotin by dissolving the solid in DI water immediately before the reaction. Allowing NHS-PEG4-biotin to remain in water for extended time will cause the NHS-ester group to hydrolyze, thereby decreasing labeling efficiency.
	<ol>
		<li>Add 2.5 &#181;L of 2 mM NHS-PEG4-biotin to the dialysis unit containing the VWF stock solution. This will result in a 20-fold molar excess of biotin compared to VWF monomers. Primary amines of VWF will react with the NHS-ester groups, thereby covalently binding to PEG4-biotin groups via amide linkages.</li>
		<li>Place the dialysis unit inside a 1.5 mL microcentrifuge tube. Seal the dialysis unit with its corresponding cap. Secure the tube-dialysis assembly with Parafilm. Keep upright and leave at room temperature for 40 min.</li>
	</ol>
	</li>
	<li>Start fluorophore labeling reaction. Prepare a 2.8 mM solution of Alexa 488 tetrafluorophenyl-ester (TFP-ester) fluorescent dye (excitationmax = 498 nm, emissionmax = 519 nm) by dissolving the fluorophore solid in DI water. Do this immediately before the reaction to prevent the TFP-ester group from hydrolyzing.
	<ol>
		<li>Add 2.9 &#181;L of the 2.8 mM 488 fluorophore to the dialysis unit. This will result in a 34-fold molar excess of fluorophore compared to VWF monomers. Remaining primary amines of VWF will react with the TFP-ester groups, thereby covalently binding to fluorophores via amide linkages.</li>
		<li>Add 2.0 &#181;L of 1 M sodium bicarbonate (dissolved in DI water) to dialysis unit. This adjusts the pH of the reaction closer to 8.0, which increases the efficiency of the TFP-ester and primary amine reaction.</li>
		<li>Secure the dialysis unit in a microcentrifuge tube just as in step 1.3.2. Store in the dark to prevent photobleaching and leave at room temperature for 1 h and 30 min.</li>
	</ol>
	</li>
	<li>Place the dialysis unit in 900 mL of 1x sterile PBS and dialyze overnight at 4 &#176;C. This will yield approximately 70 &#181;L of labeled VWF at a concentration of 0.71 mg/mL or 2.84 &#181;M (monomer concentration).</li>
	<li>Transfer labeled VWF into a microcentrifuge tube. Cover the tube with aluminum foil and protect from light. Store at 4 &#176;C. For long-term storage, add anti-microbial agent sodium azide to a final concentration of 0.02% (w/v). 
	NOTE: The protocol can be paused here.</li>
</ol>
2. Preparing Lambda DNA
<ol>
	<li>Biotinylate linear lambda DNA by filling its cohesive end sites (cos sites) with biotin-14-dCTP nucleotides according to standard protocols, repeated here in step 2.118. Fill in the remainder of the cos sites with dATP, dTTP and dGTP nucleotides.
	<ol>
		<li>Prepare 1 mM solutions of dATP, dTTP, dGTP and biotin-14-dCTP in a 10x reaction buffer (500 mM sodium chloride, 100 mM Tris hydrochloride, 100 mM magnesium chloride, 10 mM dithiothreitol, pH 7.9 at 25 &#176;C).</li>
		<li>Place 48 &#181;L of 500 ng/&#181;L lambda DNA in a PCR tube and heat for 5 min at 65 &#176;C. The cos sites of circular lambda DNA will separate under heat, linearizing the molecule and making single-stranded overhangs ready for biotinylation. Immediately after, place on ice to prevent cos sites from re-annealing.</li>
		<li>Add 5 &#181;L of 1 mM dATP, dTTP and dGTP and 4 &#181;L of 1 mM biotin-14-dCTP to the lambda DNA. Also add 2.5 &#181;L of 5 U/&#181;L Klenow Fragment (3&#39;&#224;5&#39; exo-) to catalyze the DNA synthesis.</li>
		<li>Incubate the reaction mixture for 1 h at 37 &#176;C.</li>
		<li>Add 1.2 &#181;L of 0.5 M EDTA. Then heat the reaction mixture for 5 min at 70 &#176;C. This will deactivate the Klenow Fragment and biotinylation reaction.</li>
	</ol>
	</li>
	<li>Remove excess nucleotides from lambda DNA using a spin column that can hold 10-70 &#181;L and has a 6,000 molecular weight cut-off.
	<ol>
		<li>Place the column inside a 2 mL microcentrifuge tube. Centrifuge the column and tube at 1000 x g for 2 min. Dispose of flow-through that collects in the tube.</li>
		<li>Replace the column buffer with a 1x solution of the same reaction buffer from step 2.1.1. Do this by adding 500 &#181;L of 1x buffer to the column. Centrifuge for 1 min at 1000 x g. Dispose of flow-through. Repeat these 2 more times so that a total of 1500 &#181;L has been added to the column.</li>
		<li>Place the column in a 1.5 mL microcentrifuge tube. Carefully add the solution from step 2.1.5 to the top layer of the column. Centrifuge for 4 min at 1000 x g.</li>
		<li>Collect the flow-through (40-70 &#181;L) from the microcentrifuge tube and place in a PCR tube. This contains the purified, biotinylated lambda DNA.</li>
	</ol>
	</li>
	<li>Label lambda DNA with fluorescent YOYO-1 dye (excitationmax = 490 nm, emissionmax = 509 nm) according to standard protocols, repeated here in step 2.3.120.
	<ol>
		<li>Prepare a solution of YOYO-1 dye and lambda DNA with a dye to base pair molar ratio of 1:10. Assume no DNA was lost in the purification step to calculate the base pair concentration of lambda DNA. Full-length lambda DNA has 48,502 base pairs. 
		NOTE: For example, if 50 &#181;L of solution was recovered from step 2.2.4, add 7.4 &#181;L of 500 &#181;M YOYO-1 dye.</li>
		<li>Heat the solution for 2 h at 50 &#176;C in the dark to complete the reaction.</li>
		<li>Cover the tube with aluminum foil and protect from light. Store at 4 &#176;C. The solution is now ready to be injected into microfluidic devices. 
		NOTE: The protocol can be paused here.</li>
	</ol>
	</li>
</ol>
3. Creating microfluidic channel molds in silicon wafer
<ol>
	<li>Use photolithography to create microfluidic channels with appropriate dimensions (Figure 2) on a master silicon wafer according to standard protocols19.</li>
</ol>
4. Preparation of polydimethylsiloxane (PDMS) microfluidic device
<ol>
	<li>Add 5 parts silicone elastomer base to 1 part curing agent (by mass) in a weigh boat. Stir the contents thoroughly for 1 min to create pre-cured PDMS solution.</li>
	<li>Place the master silicon wafer in a plastic Petri dish. Pour the PDMS solution over the wafer to create a 5 mm layer. Cover the dish and leave in a desiccator under vacuum for 1 h to remove air bubbles.</li>
	<li>Incubate covered Petri dish at 60 &#176;C overnight to cure PDMS into a flexible solid. Curing will result in microfluidic channels molded into the PDMS at the PDMS-wafer interface.</li>
	<li>Cut 20 x 10 mm rectangles into the PDMS, around each microfluidic channel, using a razor. Remove the rectangular PDMS blocks with tweezers.</li>
	<li>Use a 25 G blunt end needle with sharpened edges to punch a hole 0.5 mm in diameter at one end of the channel, making sure the hole goes completely through the PDMS block (Figure 2). Use a thin needle to punch out PDMS from hole. Repeat this at the other end of the channel. This will create an inlet and outlet for flow through the channel.</li>
	<li>Clean the surface of the PDMS block with vinyl cleanroom tape. Blow compressed nitrogen gas over a No. 1 &#189;, 22 x 50 mm coverslip to remove debris.</li>
	<li>Place the PDMS block with the channel side up and the coverslip into the chamber of a plasma bonding machine. Start the treatment.</li>
	<li>When treatment is complete, quickly place the PDMS block on a coverslip so that the channel is in contact with the slip. Apply pressure along the edges of the block. Place the coverslip-PDMS assembly on a hot plate at 115 &#176;C for 15 min to reinforce the permanent bond.</li>
	<li>Insert 10 cm-long, 0.25 mm inner diameter tubing into the outlet hole at the top of the PDMS block. This allows fluid to easily flow out of the channel. The device is now complete.</li>
</ol>
5. Treating surface of microfluidic device
<ol>
	<li>Inject &#60;10 &#181;L of 10 &#181;g/mL biotinylated bovine serum albumin (BSA-biotin) dissolved in sterile 1x PBS into the inlet of microfluidic device for VWF experiments. Inject &#60;10 &#181;L of 1 mg/mL BSA-biotin for lambda DNA experiments. Withhold a few microliters of BSA-biotin in the pipette tip after injection and allow the tip to remain embedded in the inlet.
	<ol>
		<li>Always keep a droplet of DI water around the tip. This will prevent air bubbles from entering the channel. Apply this technique every time a new solution is injected into the channel.</li>
		<li>Allow BSA-biotin to incubate in the device for 2 h. The BSA will nonspecifically bind to the coverslip surface (Figure 3A).</li>
	</ol>
	</li>
	<li>Remove the pipette tip. Inject &#60;10 &#181;L of casein blocking solution into the channel and allow it to incubate for 30 min. The casein will block any free sites, reducing nonspecific binding of biomolecules to the surface (Figure 3B).</li>
	<li>Remove the tip and inject &#60;10 &#181;L of 10 &#181;g/mL streptavidin dissolved in sterile 1x PBS into the channel for VWF experiments. Use 100 &#181;g/mL streptavidin for lambda DNA experiments. Incubate for 10 min. The streptavidin will bind to the biotin groups of the BSA-biotin (Figure 3C).</li>
	<li>Remove the tip and inject &#60;10 &#181;L of 1x detergent solution (0.05% Tween 20 in PBS) into the channel to wash away excess streptavidin.</li>
	<li>Remove the tip and inject &#60;10 &#181;L of either 28.4 nM VWF diluted in casein solution or lambda DNA from step 2.3.3. Incubate VWF for 3 min. Incubate lambda DNA for 45 min (Figure 3D).</li>
	<li>Remove the tip and inject &#60;10 &#181;L of 5 mM free biotin diluted in casein solution. Free biotin will block excess streptavidin binding sites on channel surface (Figure 3E).</li>
</ol>
6. Visualizing VWF and Lambda DNA under fluorescence microscopy
<ol>
	<li>Prepare 1 mL of casein blocking solution with 2.2 mM protocatechuic acid and 37 nM protocatechuate-3,4-dioxygenase (to minimize photobleaching). Load into a syringe and secure in a syringe pump. Take 30 cm long, 0.25 mm inner diameter tubing and attach one end to the syringe needle. Flow in the solution to remove air bubbles. Attach the other end of the tube to the inlet of the microfluidic device. It is recommended to do this 3 minutes after step 5.6.</li>
	<li>Select the highest magnification objective (i.e., 60-100X) of a total internal reflection (TIRF) or confocal fluorescence microscope. Add a drop of immersion oil on its objective if needed. Place the microfluidic device on the microscope stage so that the coverslip is flush with the objective.</li>
	<li>Start brightfield microscopy. Adjust focus so that any features, like debris and bubbles, are visible. Then adjust the stage in the X and Y direction until the edge of the microfluidic channel is visible and bisects the frame.</li>
	<li>Switch to the 488 channel (FITC). Adjust Z-level and TIRF angle as needed until individual green, globular molecules can be distinguished. These are either VWF or lambda DNA molecules.</li>
	<li>Adjust exposure time and laser intensity to visualize fluorescent molecules without photobleaching them too quickly. Adjust contrast to also visualize molecules more clearly.</li>
	<li>Start flow from the syringe pump so that casein blocking solution flows into the channel and out of the outlet. Do this exactly 5 minutes after step 5.6. Stop and start flow to observe the changes in the conformation of molecules. When applying flow, use rates between 5,000 and 30,000 &#181;L/h. Repeat this throughout various areas of the microfluidic device. Continue this process to locate molecules that can extend and relax upon multiple cycles of stopping and starting flow.</li>
	<li>Note how long it takes for molecules to reach maximum extension and completely relax into globules. Record videos of the continuous behavior of molecules under shear flow, selecting the best exposure time, exposure frequency and video duration that will capture the full range of extensional behavior and minimize photobleaching.</li>
	<li>Save the videos as .AVI files with a scalebar. 
	NOTE: The protocol can be paused here.</li>
</ol>
7. Image analysis of conformational changes
<ol>
	<li>Calculate the wall shear rate (<img alt="Symbol 1" src="/files/ftp_upload/60784/60784sy1.jpg" />) applied to macromolecules using the flow rate (Q) and the height (h) and width (w) of the rectangular microfluidic channel. Use the following equation to do so: 
	<img alt="Equation 1" src="/files/ftp_upload/60784/60784eq1.jpg" /></li>
	<li>Determine the length of any biomolecule under various shear rates using a customized MATLAB code (see Supplementary Files). Create a folder titled videos analysis which includes the following MATLAB codes: main.m, save_each_frame.m, get_length.m and get_length.fig. Create a subfolder within videos analysis titled videos and add .AVI files to be analyzed into it.</li>
	<li>Open main.m using MATLAB 2019a and run the code. Type in the name of the video file to be analyzed in the command window under Please input the data file to analyze:.</li>
	<li>In the opened graphical user interface (GUI), set threshold (text in the box on the top right of the window) to 20 and click on the Set threshold button to confirm.</li>
	<li>Use data cursor in the top tool bar of the window to choose one pixel anywhere on the scale bar. Click Start point in the Scale bar section on the right of the window. The (x,y) position of the chosen pixel will appear on the right of the button. Click on the Pixel size (&#181;m) button. The pixel size needs to be measured only once in each video.</li>
	<li>Choose any pixel on the molecule of interest. Click on Start point in the VWF section. After the position of the chosen pixel appears on the text box on the right, click on Left end, Right end and String length to get the molecular length in the image. 
	NOTE: This step can be used to analyze the conformational changes of any biomolecule, despite the code having a specific section named VWF.</li>
	<li>Double-check the left and right end of the molecule. Zoom in and use the data cursor to check the pixel position of interest. Manually choose the pixel as an end and recalculate the length (in pixels) when necessary.</li>
	<li>Record the pixel size in &#181;m and string length in pixel number into an Excel sheet and calculate the string length in &#181;m.</li>
	<li>Repeat the steps above for every image. Use Last, Next button in the bottom right corner of the GUI to switch among images in the same video file. Click on Close to close the GUI window.</li>
</ol>

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The authors declare no competing interests.

To obtain high quality data of single-molecule conformational changes using fluorescence microscopy as described in this method, it is critical to incubate the molecule for the appropriate amount of time, minimize its nonspecific interactions with the surface and establish microscope settings that reduce photobleaching. The ability of the molecule to freely change conformation is related to the number of biotin-streptavidin interactions formed between the molecule and the surface. As mentioned previously, this must be co...

Mechanisms of how biomolecules respond to environmental stimuli have been studied widely. In a flow environment in particular, shear and elongational forces regulate the conformational changes and potentially the function of biomolecules. Typical examples include shear-induced unraveling of lambda DNA and von Willebrand factor (VWF). Lambda DNA has been used as a tool to understand conformational dynamics of individual, flexible polymer chains and the rheology of polymer solutions1,2,3,4. VWF is a natural flow sensor that aggregates platelets at wound sites of blood vessels with abnormal shear rates and flow patterns. Unraveling of VWF is essential in activating the binding of platelets to the A1 domain and collagen binding to the A3 domain. In addition, high shear-induced A2 domain unfolding allows the cleavage of VWF, which regulates its molecular weight distribution in circulation5,6. Thus, direct visualization of how these molecules behave under flow can greatly enhance our fundamental understanding of their biomechanics and function, which in turn can enable novel diagnostic and therapeutic applications.
Typical methodologies to characterize single-molecule conformations include optical/magnetic tweezers, atomic force microscopy (AFM) and single-molecule F&#246;rster resonance energy transfer (FRET)7. Single-molecule force spectroscopy is a powerful tool to investigate the force and motion associated with the conformational changes of biomolecules. However, it lacks the ability to map overall molecular conformations8. AFM is capable of imaging with high spatial resolution but is limited in temporal resolution9,10. In addition, contact between the tip and the sample may confound the response induced by flow. Other methods like FRET and nanopore analytics determine single-molecule protein folding and unfolding states based on the detection of intramolecular distance and excluded volumes. However, these methods are still in their infancy and limited in their direct observation of single-molecule conformations11,12,13,14.
On the other hand, directly observing macromolecules with high temporal and spatial resolution under fluorescence microscopy has improved our understanding of single-molecule dynamics in many biological processes15,16. For example, Fu et al. recently achieved simultaneous visualization of VWF elongation and platelet receptor binding for the first time. In their work, VWF molecules were immobilized on the surface of a microfluidic channel through biotin-streptavidin interactions and imaged under total internal reflection fluorescence (TIRF) microscopy at varying shear flow environments17. Applying a similar method as Fu&#39;s, we here demonstrate that conformations of VWF and lambda DNA can be directly observed under both TIRF and confocal fluorescence microscopy. As shown in Figure 1, microfluidic devices are used to create and control shear flow, and biomolecules are immobilized on the channel surface. Upon the application of varying shear rates, conformations of the same molecule are recorded to measure the extensional length, also shown in Figure 1. The method could be widely applied to explore other polymer behaviors under complex flow environments for both rheological and biological studies.

<table><tbody><tr><td>Alexa Fluor 488 Labeling Kit</td><td>Invitrogen</td><td>A30006</td><td></td></tr><tr><td>Bio-Spin P-6 Gel Columns</td><td>Bio-Rad</td><td>7326221</td><td></td></tr><tr><td>Biotin</td><td>Sigma-Aldrich</td><td>B4501</td><td>Use as free biotin in Step 5.6</td></tr><tr><td>Biotin-14-dCTP</td><td>AAT Bioquest</td><td>17019</td><td></td></tr><tr><td>BSA-Biotin</td><td>Sigma-Aldrich</td><td>A8549</td><td></td></tr><tr><td>Coverslips</td><td>VWR</td><td>48393-195</td><td>No. 1 &amp;frac12;, 22 x 50 mm</td></tr><tr><td>dNTP Set</td><td>Invitrogen</td><td>10297018</td><td></td></tr><tr><td>Float Buoys for Mini Dialysis Device</td><td>Thermo Scientific</td><td>69588</td><td></td></tr><tr><td>Klenow Fragment (3&#039;&amp;rarr;5&#039; exo-)</td><td>New England BioLabs</td><td>M0212S</td><td>Use for 10X reaction buffer in Step 2.1.1 and 1X reaction buffer in Step 2.2.2</td></tr><tr><td>Lambda DNA</td><td>New England BioLabs</td><td>N3011S</td><td></td></tr><tr><td>Mini Dialysis Device</td><td>Thermo Scientific</td><td>69570</td><td>10K MWCO, 0.1 mL volume</td></tr><tr><td>NEBuffer 4</td><td>New England BioLabs</td><td>B7004S</td><td></td></tr><tr><td>NHS-PEG4-Biotin</td><td>Thermo Scientific</td><td>21330</td><td></td></tr><tr><td>Protocatechuate 3,4-Dioxygenase</td><td>Sigma-Aldrich</td><td>P8279</td><td></td></tr><tr><td>Protocatechuic acid</td><td>Santa Cruz Biotechnology</td><td>sc-205818</td><td></td></tr><tr><td>Silicone Elastomer Kit for PDMS Fabrication</td><td>The Dow Chemical Company</td><td>4019862</td><td></td></tr><tr><td>Streptavidin</td><td>Sigma-Aldrich</td><td>85878</td><td></td></tr><tr><td>The Blocking Solution</td><td>CANDOR Bioscience</td><td>110 050</td><td>Use as casein blocking solution throughout protocol</td></tr><tr><td>Vinyl Cleanroom Tape</td><td>Fisher Scientific</td><td>19-120-3217</td><td></td></tr><tr><td>von Willebrand Factor, Human Plasma</td><td>Millipore Sigma</td><td>681300</td><td></td></tr><tr><td>YOYO-1 Dye</td><td>AAT Bioquest</td><td>17580</td><td></td></tr><tr><td>0.25 mm Inner Diameter Tubing</td><td>Cole-Parmer</td><td>EW-06419-00</td><td></td></tr><tr><td>25 Gauge Needle</td><td>Thomas Scientific</td><td>JG2505X</td><td></td></tr></tbody></table>

characterizing single-molecule conformational changes under shear flow with fluorescence microscopy

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

Observing the dynamic behavior of biomolecules such as VWF and lambda DNA is highly dependent on optimizing their binding to the device surface. Incubating surface treatments for the recommended times in the microfluidic device is crucial to obtaining binding with a few anchorage points, so that molecules can freely extend and relax upon changing flow. If the proteins or DNA are bound too strongly with multiple linkages, they will either extend to limited lengths or not extend at all. This occurs particularly with VWF wh...

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

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

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

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Research

JoVE Journal

Bioengineering

5.7K Views.  Lehigh University. We present a protocol for immobilizing single macromolecules in microfluidic devices and quantifying changes in their conformations under shear flow. This protocol is useful for characterizing the biomechanical and functional properties of biomolecules such as proteins and DNA in a flow environment.

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

Submillisecond Conformational Changes in Proteins Resolved by Photothermal Beam Deflection

Photothermal beam deflection together with photo-acoustic calorimetry and thermal grating belongs to the family of photothermal methods that monitor the time-profile volume and enthalpy changes of light induced conformational changes in proteins on microsecond to millisecond time-scales that are not accessible using traditional stop-flow instruments. In addition, since overall changes in volume and/or enthalpy are probed, these techniques can be applied to proteins and other biomacromolecules that lack a fluorophore and or a chromophore label. To monitor dynamics and energetics of structural changes associated with Ca2+ binding to calcium transducers, such neuronal calcium sensors, a caged calcium compound, DM-nitrophen, is employed to photo-trigger a fast (&#964; &#60; 20 &#956;sec) increase in free calcium concentration and the associated volume and enthalpy changes are probed using photothermal beam deflection technique.

Here we report an application of the photothermal beam deflection technique in combination with a caged calcium compound, DM-nitrophen, to monitor microsecond and millisecond dynamics and energetics of structural changes associated with the calcium association to a neuronal calcium sensor, Downstream Regulatory Element Antagonist Modulator.

Photothermal beam deflection together with photo-acoustic calorimetry and thermal grating belongs to the family of photothermal methods that monitor the ...

Chemistry

Flexural Rigidity Measurements of Biopolymers Using Gliding Assays

Microtubules are cytoskeletal polymers which play a role in cell division, cell mechanics, and intracellular transport. Each of these functions requires microtubules that are stiff and straight enough to span a significant fraction of the cell diameter. As a result, the microtubule persistence length, a measure of stiffness, has been actively studied for the past two decades1. Nonetheless, open questions remain: short microtubules are 10-50 times less stiff than long microtubules2-4, and even long microtubules have measured persistence lengths which vary by an order of magnitude5-9.
	Here, we present a method to measure microtubule persistence length. The method is based on a kinesin-driven microtubule gliding assay10. By combining sparse fluorescent labeling of individual microtubules with single particle tracking of individual fluorophores attached to the microtubule, the gliding trajectories of single microtubules are tracked with nanometer-level precision. The persistence length of the trajectories is the same as the persistence length of the microtubule under the conditions used11. An automated tracking routine is used to create microtubule trajectories from fluorophores attached to individual microtubules, and the persistence length of this trajectory is calculated using routines written in IDL.
	This technique is rapidly implementable, and capable of measuring the persistence length of 100 microtubules in one day of experimentation. The method can be extended to measure persistence length under a variety of conditions, including persistence length as a function of length along microtubules. Moreover, the analysis routines used can be extended to myosin-based acting gliding assays, to measure the persistence length of actin filaments as well.

A method to measure the persistence length or flexural rigidity of biopolymers is described. The method uses a kinesin-driven microtubule gliding assay to experimentally determine the persistence length of individual microtubules and is adaptable to actin-based gliding assays.

Microtubules are cytoskeletal polymers which  play a role in cell division, cell mechanics, and intracellular transport. Each of these functions requires ...

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One &#945;-Synuclein Monomer at a Time

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural dynamics and its contributions to biology. While the FRET-based ruler reports on inter-dye distances in the 3-10 nm range, other spectroscopic techniques, such as protein-induced fluorescence enhancement (PIFE), report on the proximity between a dye and a protein surface in the shorter 0-3 nm range. Regardless of the method of choice, its use in measuring freely-diffusing biomolecules one at a time retrieves histograms of the experimental parameter yielding separate centrally-distributed sub-populations of biomolecules, where each sub-population represents either a single conformation that stayed unchanged within milliseconds, or multiple conformations that interconvert much faster than milliseconds, and hence an averaged-out sub-population. In single-molecule FRET, where the reported parameter in histograms is the inter-dye FRET efficiency, an intrinsically disordered protein, such as the &#945;-Synuclein monomer in buffer, was previously reported as exhibiting a single averaged-out sub-population of multiple conformations interconverting rapidly. While these past findings depend on the 3-10 nm range of the FRET-based ruler, we sought to put this protein to the test using single-molecule PIFE, where we track the fluorescence lifetime of site-specific sCy3-labeled &#945;-Synuclein proteins one at a time. Interestingly, using this shorter range spectroscopic proximity sensor, sCy3-labeled &#945;-Synuclein exhibits several lifetime sub-populations with distinctly different mean lifetimes that interconvert in 10-100 ms. These results show that while &#945;-Synuclein might be disordered globally, it nonetheless attains stable local structures. In summary, in this work we highlight the advantage of using different spectroscopic proximity sensors that track local or global structural changes one biomolecule at a time.

Time-resolved single-molecule protein-induced fluorescence enhancement is a useful fluorescence spectroscopic proximity sensor sensitive to local structural changes in proteins. Here we show it can be used to uncover stable local conformations in &#945;-Synuclein, which is otherwise known as globularly unstructured and unstable when measured using the longer range FRET ruler.

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural ...

Biochemistry

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Monitoring Conformational Dynamics of Single Unmodified Proteins using Plasmonic Nanotweezers

Current single-molecule techniques to characterize proteins typically require labels, tethers, or the use of non-native solution conditions. Such changes can alter protein biophysics and reduce the usefulness of the data acquired. Plasmonic nanotweezers is a technique that uses localized surface plasmon resonance (LSPR) on gold nanostructures to enhance the electric field within a confined hotspot region. This field enhancement allows for the use of low laser powers to trap single nanoparticles far smaller than conventional optical tweezers, down to only a few nanometers in diameter, such as single proteins. Trapping of single protein molecules within the hotspot region induces a shift in the local refractive index (nprotein &#62; nwater), altering light scattering as a product of the molecule&#39;s polarisability, which is affected by its volume, shape anisotropy, and refractive index. An avalanche photodiode (APD) collects the subsequent changes in light scattering. These alterations can then be analyzed to determine changes in the trapped molecule, including its size, global conformation, and dynamics of conformational change over time. The incorporation of microfluidics within the system allows for controlled environmental changes and real-time monitoring of their subsequent effects on the molecule. In this protocol, we demonstrate the steps to trap single protein molecules, alter their environmental solution conditions, and monitor their corresponding conformational changes using a plasmonic nanotweezers system.

Plasmonic nanotweezers use localized surface plasmon resonance in gold nanostructures to trap single nanoparticles, including proteins, within a nanometer-scale optical field. Changes in the scattered signal reveal protein presence and conformational dynamics, enabling monitoring without fluorophore modifications or surface tethering.

Current single-molecule techniques to characterize proteins typically require labels, tethers, or the use of non-native solution conditions. Such changes ...

Engineering

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Biology

Monitoring GTPase Atlastin in Different Nucleotide Loading States

Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

The rigid-body rotation of the three-helical middle domain (3HB) relative to the GTPase domain of dynamin-like protein atlastin (ATL) is a crucial driver of homotypic membrane fusion within the endoplasmic reticulum (ER). Disruptions in this process have been associated with hereditary spastic paraplegia (HSP), a neurodegenerative disorder. Structural and biochemical studies suggest that the conformational changes in ATL are linked to GTP hydrolysis, but real-time visualization of these conformational dynamics during the GTP hydrolysis cycle remains challenging. To better understand the mechanical mechanisms behind ATL function, single-molecule F&#246;rster resonance energy transfer (smFRET) was utilized. Three specific strategies were employed to immobilize the N-terminal cytosolic region of human ATL1 (ATL1cyto) in a streptavidin-coated microfluidic chamber, facilitating the application of intramolecular and intermolecular smFRET imaging. This allowed precise monitoring of protein conformations in various nucleotide-loading states, providing direct insights into individual molecular behaviors. This method can be applied to study other mechanochemical proteins as well.

The functions of dynamin superfamily proteins depend on conformational changes coupled with GTP hydrolysis. A system is described using the single-molecule FRET (smFRET) technique to monitor the conformational dynamics of dynamin-like GTPase atlastin in different nucleotide-loading states.

The rigid-body rotation of the three-helical middle domain (3HB) relative to the GTPase domain of dynamin-like protein atlastin (ATL) is a crucial driver ...

Time-Resolved Fluorescence Anisotropy from Single Molecules for Characterizing Local Flexibility in Biomolecules

We describe a protocol for conducting time-resolved fluorescence anisotropy at the single-molecule level using confocal microscopy to investigate the local flexibility and dynamics of the deoxyribonucleic acid (DNA)-binding forkhead (FKH) domain of the FoxP1 transcription factor. FoxP1 dimerizes through a three-dimensional domain-swapping (3D-DS) mechanism, forming a disordered intermediate with or without DNA. Since 3D-DS involves an intrinsically disordered region, understanding its behavior is crucial for elucidating the structural and functional properties of FoxP1. Using a single-cysteine-labeled FoxP1, we conducted single-molecule fluorescence anisotropy (smFA) experiments, applying dynamic anisotropy Photon Distribution Analysis (daPDA) and time-resolved anisotropy Burst Variance Analysis (traBVA) approaches to probe local flexibility and dynamics. This protocol provides a detailed, step-by-step guide for smFA measurements, emphasizing time-resolved analyses, variance, and probability distribution techniques to capture structural dynamics across different timescales. This approach enabled us to relate dynamics and heterogeneity to FoxP1 dimerization and DNA binding, highlighting the complex action mechanism that characterizes this transcription factor.

Here, we present the protocol to study the local flexibility and dynamics of biomolecules using time-resolved fluorescence anisotropy at the single-molecule level in confocal microscopy mode.

We describe a protocol for conducting time-resolved fluorescence anisotropy at the single-molecule level using confocal microscopy to investigate the ...

Ensemble Force Spectroscopy by Shear Forces

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

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

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

Live Cell Single-Molecule Imaging-Based Quantification of Protein Interactions

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) have been considered critical in cellular organization and an increasing number of cellular functions. Characterizing LLPS in live cells is also important because aberrant condensation has been linked to numerous diseases, including cancers and neurodegenerative disorders. LLPS is often driven by selective, transient, and multivalent interactions between intrinsically disordered proteins. Of great interest are the interaction dynamics of proteins participating in LLPS, which are well-summarized by measurements of their binding residence time (RT), that is, the amount of time they spend bound within condensates. Here, we present a method based on live-cell single-molecule imaging that allows us to measure the mean RT of a specific protein within condensates. We simultaneously visualize individual protein molecules and the condensates with which they associate, use single-particle tracking (SPT) to plot single-molecule trajectories, and then fit the trajectories to a model of protein-droplet binding to extract the mean RT of the protein. Finally, we show representative results where this single-molecule imaging method was applied to compare the mean RTs of a protein at its LLPS condensates when fused and unfused to an oligomerizing domain. This protocol is broadly applicable to measuring the interaction dynamics of any protein that participates in LLPS.

Author Spotlight: Evaluation of Protein-Condensate Dynamics in Live Human Cells

Many intrinsically disordered proteins have been shown to participate in the formation of highly dynamic biomolecular condensates, a behavior important for numerous cellular processes. Here, we present a single-molecule imaging-based method for quantifying the dynamics by which proteins interact with each other in biomolecular condensates in live cells.

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) have been considered critical in cellular organization and an increasing number ...

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

Summary

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Chapters in this video

Submillisecond Conformational Changes in Proteins Resolved by Photothermal Beam Deflection

Flexural Rigidity Measurements of Biopolymers Using Gliding Assays

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One α-Synuclein Monomer at a Time

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

Monitoring Conformational Dynamics of Single Unmodified Proteins using Plasmonic Nanotweezers

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

Time-Resolved Fluorescence Anisotropy from Single Molecules for Characterizing Local Flexibility in Biomolecules

Ensemble Force Spectroscopy by Shear Forces

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates