Name: characterizing single-molecule conformational changes under shear flow with fluorescence microscopy
Uploaded: 2020-01-25T14:00:00
Description: Watch this Scientific Journal Video about Characterizing Single-Molecule Conformational Changes Under Shear Flow with Fluorescence Microscopy at JoVE.com

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 solu...

<ol>
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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+folding+studied+by+single-molecule+FRET.">Protein folding studied by single-molecule FRET.</a> Current Opinion in Structural Biology. 18 (1), 16-26 (2008).</li><li>Howorka, S., Siwy, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanopore+analytics:+sensing+of+single+molecules.">Nanopore analytics: sensing of single molecules.</a> Chemical Society Reviews. 38 (8), 2360-2384 (2008).</li><li>Talaga, D. S., Li, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+protein+unfolding+in+solid+state+nanopores.">Single-molecule protein unfolding in solid state nanopores.</a> Journal of the American Chemical Society. 131 (26), 9287-9297 (2009).</li><li>Moerner, W. E., Fromm, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+of+single-molecule+fluorescence+spectroscopy+and+microscopy.">Methods of single-molecule fluorescence spectroscopy and microscopy.</a> Review of Scientific Instruments. 74 (8), 3597-3619 (2003).</li><li>Xia, T., Li, N., Fang, X. 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B., Cui, Y., Bustamante, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Overstretching+B-DNA:+The+Elastic+Response+of+Individual+Double-Stranded+and+Single-Stranded+DNA+Molecules.">Overstretching B-DNA: The Elastic Response of Individual Double-Stranded and Single-Stranded DNA Molecules.</a> Science. 271 (5250), 795-799 (1996).</li><li>Wang, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shear-Induced+Extensional+Response+Behaviors+of+Tethered+von+Willebrand+Factor.">Shear-Induced Extensional Response Behaviors of Tethered von Willebrand Factor.</a> Biophysical Journal. 116 (11), 29092 (2019).</li><li>Carlsson, C., Johnsson, M., &#197;kerman, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Double+bands+in+DNA+gel+electrophoresis+caused+by+bis-intercalating+dyes.">Double bands in DNA gel electrophoresis caused by bis-intercalating dyes.</a> Nucleic Acids Research. 23 (13), 2413-2420 (1995).</li><li>Collins, B. E., Ye, L. F., Duzdevich, D., Greene, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+curtains:+Novel+tools+for+imaging+protein&#8211;nucleic+acid+interactions+at+the+single-molecule+level.">DNA curtains: Novel tools for imaging protein&#8211;nucleic acid interactions at the single-molecule level.</a> Methods in Cell Biology. 123, 217-234 (2014).</li><li>Green, E. C., Wind, S., Fazio, T., Gorman, J., Visnapuu, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+Curtains+for+High-Throughput+Single-Molecule+Optical+Imaging.">DNA Curtains for High-Throughput Single-Molecule Optical Imaging.</a> Methods in Enzymology. 472, 293-315 (2010).</li></ol>

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 ...

Quantifying the conformational changes in single bio-molecules under shear flow is useful for understanding the function and biophysics of macromolecules like protein and DNA. Fluorescence microscopy allows real-time in-situ visualization of single molecules in physiological flow environments with high temporal and spatial resolution. This is unmatched by other techniques like AFM.By characterizing the shear force under which proteins our DNA have global conformational changes, one can better design drugs that mimic these biophysical properties. This protocol can be widely applied to study the behavior or other polymers, especially bio polymers, in varying flow conditions and to investigate the rheology of complex fluids. The timing of steps is critical to success.We suggest practicing the microfluidic device surface treatment and fluorescence microscopy steps many times to execute them in an efficient fashion. Capturing conformational changes in biomolecules is a visual and dynamic phenomenon best viewed on film. Scientists will better understand this method after seeing the real-time protein and DNA microscopy presented in this video.To prepare the microfluidic device, add five parts of silicone elastomer base to one part curing agent in a weigh boat. and stir the contents thoroughly for one minute to create a pre-cured PDMS solution. Place the master silicon wafer in a plastic Petri dish and pour the PDMS solution over the wafer to create a five-millimeter layer.Cover the dish and leave it in a desiccator under vacuum for one hour to remove air bubbles. Then incubate the covered Petri dish at 60 degrees Celsius overnight to cure the PDMS into a flexible solid, which will result in microfluidic channels being molded into the PDMS at the PDMS wafer interface. On the next day, use a razor to cut 20 by 10 millimeter rectangles around each microfluidic channel in the PDMS and remove the rectangular blocks with tweezers.Use a 25-gauge blunt end needle with sharpened edges to punch a 0.5 millimeter diameter hole at one end of the channel, making sure that the hole goes completely through the PDMS block. Use a thin needle to punch the PDMS out of the hole and repeat the process at the other end of the channel. Place the PDMS block with the channel side up and the coverslip into the chamber of a plasma bonding machine and start the treatment.When treatment is complete, quickly place the PDMS block on a coverslip so that the channel is in contact with the slip. Apply pressure to the edges of the block, then place the coverslip PDMS assembly on a hot plate at 115 degrees Celsius for 15 minutes to reinforce the permanent bond. Finally, insert a 10-centimeter long.0.25 millimeter inner diameter tube into the outlet hole at the top of the PDMS block to allow fluid to easily flow out of the channel. For von Willebrand factor or VWF experiments inject up to 10 microliters of 10 micrograms per milliliter BSA-Biotin dissolved in sterile 1X PBS into the inlet of the microfluidic device. Withhold a few microliters of BSA-Biotin in the pipette tip after injection and allow the tip to remain embedded in the inlet.Allow the BSA-Biotin to incubate in the device for two hours which will result in BSA non-specifically binding to the coverslip surface. Then remove the pipette tip and inject up to 10 microliters of casein blocking solution into the channel. Allow the casein solution to incubate for 30 minutes so it can block any free sites and reduce nonspecific binding of the biomolecules to the surface.Remove the tip and inject up to 10 microliters of streptavidin in sterile PBS into the channel, which will bind to the biotin groups of the BSA-Biotin. Next, inject up to 10 microliters of detergent solution to wash away excess streptavidin. Remove the tip and inject either VWF in casein solution or lambda DNA.Incubate VWF for three minutes or lambda DNA for 45 minutes. Then inject five millimolar free biotin to block the excess streptavidin binding sites. Load one milliliter of casein blocking solution into a syringe and secure it in a syringe pump.Then attach one end of the tubing to the syringe needle and flow in the solution to remove air bubbles. Approximately three minutes after the free biotin injection attach the other end of the tube to the inlet of the microfluidic device. Select the highest magnification objective of the TIRF or confocal fluorescence microscope.If needed, add a drop of immersion oil on the objective. Place the microfluidic device on the microscope stage so that the coverslip is flush with the objective. Start bright-field microscopy and adjust the focus so that features such as debris and bubbles are visible.Then adjust the stage in the X and Y directions until the edge of the microfluidic channel is visible and bisects the frame. Switch to the FITC channel and adjust the Z level and TIRF angle as needed until the individual green globular molecules can be distinguished. Adjust laser intensity and exposure time to visualize fluorescent molecules without photobleaching them too quickly.Then adjust the contrast to better visualize the molecules. Exactly five minutes after free biotin injection start and stop flow from the syringe pump to observe the changes in molecular conformation using flow rates between 5000 and 30, 000 microliters per hour. With VWF it is critical to apply at least a small amount of flow, such as 30 seconds of 10, 000 microliter per hour flow exactly five minutes after free biotin injection.Repeat this throughout various areas of the microfluidic device and locate molecules that can extend and relax upon multiple cycles of starting and stopping flow. Note how long it takes for molecules to reach maximum extension and complete relaxation and record videos of the continuous behavior of molecules under shear flow. The flexibility of a molecule to change confirmation is often demonstrated by its ability to lengthen as higher shear rates are applied due to increasing flow.The extension versus shear rate curve of a von Willebrand factor molecule is useful for characterizing the biomechanical properties of the protein. Fluorescence microscopy images of lambda DNA similarly show increased extension upon higher shear rates at 30 seconds and gradual relaxation over two minutes after flow is stopped. When working with VWF, remember to incubate it only for three minutes and free biotin for five minutes before starting flow.This is critical for forming the optimal number or biotin-streptavidin linkages needed for reversible unraveling. This method can be adapted to visualize real time interactions between multiple biomolecules and characterize their functions. For example, one could better understand platelet plug formation by visualizing VWF unraveling under high shear and its binding with the platelet adhesion molecule GPIb-alpha with this method.

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Research

JoVE Journal

Bioengineering

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 ...