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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#189;, 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&#39;&#8594;5&#39; 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...

Watch this Scientific Journal Video about Characterizing Single-Molecule Conformational Changes Under Shear Flow with Fluorescence Microscopy at JoVE.com

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 ...

characterizing-single-molecule-conformational-changes-under-shear

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

Parallel-plate Flow Chamber and Continuous Flow Circuit to Evaluate Endothelial Progenitor Cells under Laminar Flow Shear Stress

The overall goal of this method is to describe a technique to subject adherent cells to laminar flow conditions and evaluate their response to well quantifiable fluid shear stresses1.
Our flow chamber design and flow circuit (Fig. 1) contains a transparent viewing region that enables testing of cell adhesion and imaging of cell morphology immediately before flow (Fig. 11A, B), at various time points during flow (Fig. 11C), and after flow (Fig. 11D). These experiments are illustrated with human umbilical cord blood-derived endothelial progenitor cells (EPCs) and porcine EPCs2,3.
This method is also applicable to other adherent cell types, e.g. smooth muscle cells (SMCs) or fibroblasts.
The chamber and all parts of the circuit are easily sterilized with steam autoclaving. In contrast to other chambers, e.g. microfluidic chambers, large numbers of cells (&gt; 1 million depending on cell size) can be recovered after the flow experiment under sterile conditions for cell culture or other experiments, e.g. DNA or RNA extraction, or immunohistochemistry (Fig. 11E), or scanning electron microscopy5. The shear stress can be adjusted by varying the flow rate of the perfusate, the fluid viscosity, or the channel height and width. The latter can reduce fluid volume or cell needs while ensuring that one-dimensional flow is maintained. It is not necessary to measure chamber height between experiments, since the chamber height does not depend on the use of gaskets, which greatly increases the ease of multiple experiments. Furthermore, the circuit design easily enables the collection of perfusate samples for analysis and/or quantification of metabolites secreted by cells under fluid shear stress exposure, e.g. nitric oxide (Fig. 12)6.

We are describing a method to subject adherent cells to laminar flow shear stress in a sterile continuous flow circuit. The cells' adhesion, morphology can be studied through the transparent chamber, samples obtained from the circuit for metabolite analysis and cells harvested after shear exposure for future experiments or culture.

The overall goal of this method is to describe a technique to subject adherent cells to laminar flow conditions and evaluate their response to well ...

Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

The generation and detection of mechanical forces is a ubiquitous aspect of cell physiology, with direct relevance to cancer metastasis1, atherogenesis2 and wound healing3. In each of these examples, cells both exert force on their surroundings and simultaneously enzymatically remodel the extracellular matrix (ECM). The effect of forces on ECM has thus become an area of considerable interest due to its likely biological and medical importance4-7.
	Single molecule techniques such as optical trapping8, atomic force microscopy9, and magnetic tweezers10,11 allow researchers to probe the function of enzymes at a molecular level by exerting forces on individual proteins. Of these techniques, magnetic tweezers (MT) are notable for their low cost and high throughput. MT exert forces in the range of ~1-100 pN and can provide millisecond temporal resolution, qualities that are well matched to the study of enzyme mechanism at the single-molecule level12. Here we report a highly parallelizable MT assay to study the effect of force on the proteolysis of single protein molecules. We present the specific example of the proteolysis of a trimeric collagen peptide by matrix metalloproteinase 1 (MMP-1); however, this assay can be easily adapted to study other substrates and proteases.

In this article we describe the use of magnetic tweezers to study the effect of force on enzymatic proteolysis at the single molecule level in a highly parallelizable manner.

The  generation and detection of mechanical forces is a ubiquitous aspect of cell physiology,  with direct relevance to cancer metastasis1, atherogenesis2 ...

Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as growth factors. Here, we outline a method to create opposing gradients of adhesive and soluble cues in a microfluidic chamber, which is compatible with live cell imaging. A copolymer of poly-L-lysine and polyethylene glycol (PLL-PEG) is employed to passivate glass coverslips and prevent non-specific adsorption of biomolecules and cells. Next, microcontact printing or dip pen lithography are used to create tracks of streptavidin on the passivated surfaces to serve as anchoring points for the biotinylated peptide arginine-glycine-aspartic acid (RGD) as the adhesive cue. A microfluidic device is placed onto the modified surface and used to create the gradient of adhesive cues (100% RGD to 0% RGD) on the streptavidin tracks. Finally, the same microfluidic device is used to create a gradient of a chemoattractant such as fetal bovine serum (FBS), as the soluble cue in the opposite direction of the gradient of adhesive cues.

A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as ...

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 &#197;. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.

We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range ...

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM)

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and sensitivity in cells because the resolution of conventional light microscopy is diffraction-limited to ~250 nm. By combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM), PPIs can be visualized in cells with single molecule sensitivity and nanometer spatial resolution. BiFC is a commonly used technique for visualizing PPIs with fluorescence contrast, which involves splitting of a fluorescent protein into two non-fluorescent fragments. PALM is a recent superresolution microscopy technique for imaging biological samples at the nanometer and single molecule scales, which uses phototransformable fluorescent probes such as photoactivatable fluorescent proteins (PA-FPs). BiFC-PALM was demonstrated by splitting PAmCherry1, a PA-FP compatible with PALM, for its monomeric nature, good single molecule brightness, high contrast ratio, and utility for stoichiometry measurements. When split between amino acids 159 and 160, PAmCherry1 can be made into a BiFC probe that reconstitutes efficiently at 37 &#176;C with high specificity to PPIs and low non-specific reconstitution. Ras-Raf interaction is used as an example to show how BiFC-PALM helps to probe interactions at the nanometer scale and with single molecule resolution. Their diffusion can also be tracked in live cells using single molecule tracking (smt-) PALM. In this protocol, factors to consider when designing the fusion proteins for BiFC-PALM are discussed, sample preparation, image acquisition, and data analysis steps are explained, and a few exemplary results are showcased. Providing high spatial resolution, specificity, and sensitivity, BiFC-PALM is a useful tool for studying PPIs in intact biological samples.

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation BiFC-PALM

Protein-protein interactions are visualized in cells with nanometer spatial resolution by combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM). Described here is the use of BiFC-PALM for imaging Ras-Raf interactions in U2OS cells for visualizing the nanoscale clustering and diffusion of individual Ras-Raf complexes.

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

Label-free Single Molecule Detection Using Microtoroid Optical Resonators

Detecting small concentrations of molecules down to the single molecule limit has impact on areas such as early detection of disease, and fundamental studies on the behavior of molecules. Single molecule detection techniques commonly utilize labels such as fluorescent tags or quantum dots, however, labels are not always available, increase cost and complexity, and can perturb the events being studied. Optical resonators have emerged as a promising means to detect single molecules without the use of labels. Currently the smallest particle detected by a non-plasmonically-enhanced bare optical resonator system in solution is a 25 nm polystyrene sphere1. We have developed a technique known as Frequency Locking Optical Whispering Evanescent Resonator (FLOWER) that can surpass this limit and achieve label-free single molecule detection in aqueous solution2. As signal strength scales with particle volume, our work represents a &#62; 100x improvement in the signal to noise ratio (SNR) over the current state of the art. Here the procedures behind FLOWER are presented in an effort to increase its usage in the field.

We have developed a label-free biosensing system based on optical resonator technology known as Frequency Locking Optical Whispering Evanescent Resonator (FLOWER) that is capable of detecting single molecules in solution. Here the procedures behind this work are described and presented.

Detecting small concentrations of molecules down to the single molecule limit has impact on areas such as early detection of disease, and fundamental ...

Designing Microfluidic Devices for Studying Cellular Responses Under Single or Coexisting Chemical/Electrical/Shear Stress Stimuli

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other physical or chemical stimuli. They have been commonly used for in vitro cell studies by providing in vivo like surroundings. Especially, how cells response to chemical gradients, electrical fields, and shear stresses has drawn many interests since these phenomena are important in understanding cellular properties and functions. These microfluidic chips can be made of glass substrates, silicon wafers, polydimethylsiloxane (PDMS) polymers, polymethylmethacrylate (PMMA) substrates, or polyethyleneterephthalate (PET) substrates. Out of these materials, PMMA substrates are cheap and can be easily processed using laser ablation and writing. Although a few microfluidic devices have been designed and fabricated for generating multiple, coexisting chemical and electrical stimuli, none of them was considered efficient enough in reducing experimental repeats, particular for screening purposes. In this report, we describe our design and fabrication of two PMMA-based microfluidic chips for investigating cellular responses, in the production of reactive oxygen species and the migration, under single or coexisting chemical/electrical/shear stress stimuli. The first chip generates five relative concentrations of 0, 1/8, 1/2, 7/8, and 1 in the culture regions, together with a shear stress gradient produced inside each of these areas. The second chip generates the same relative concentrations, but with five different electric field strengths created within each culture area. These devices not only provide cells with a precise, controllable micro-environment but also greatly increase the experimental throughput.

Micro-fabricated devices integrated with fluidic components provide an in vitro platform for cell studies mimicking the in vivo micro-environment. We developed polymethylmethacrylate-based microfluidic chips for studying cellular responses under single or coexisting chemical/electrical/shear stress stimuli.

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other ...