This work was supported by the Burroughs Wellcome Career Award at the Scientific Interface (A.R.D.), the National Institutes of Health through the NIH Director's New Innovator Award Program 1-DP2-OD007078 (A.R.D.), the William Bowes Jr. Stanford Graduate Fellowship (A.S.A.), and the Stanford Cardiovascular Institute Younger Predoctoral Fellowship (J.C.). The authors thank James Spudich for loaning microscopy equipment.

1. Flow Cell Preparation
<ol>
	<li>Coverslips (#1.5, 22x22 mm and 22x40 mm, VWR) are cleaned using sonication.
	<ol>
		<li>Add the coverslips to a small glass container capable of holding coverslips and fitting in the sonicator (see step 2).</li>
		<li>Fill the container with isopropanol and sonicate in a bath sonicator for 20 minutes.</li>
		<li>Discard the isopropanol and rinse the coverslips with copious quantities of deionized water produced by a Barnsted MilliQ apparatus or similar device. Fill the container with water and sonicate for 20 minutes.</li>
		<li>After sonication, dry the coverslips in a stream of filtered, dust-free air. Use only coverslips that appear clean (i.e. no smudges).</li>
		<li>Gently flame the coverslips for a few seconds to clean them of any remaining dust and moisture: pass the coverslips through a gas flame produced with a Bunsen burner, taking care not to warp the coverslip due to excess heat. This step removes residual surface contaminants.</li>
	</ol>
	</li>
	<li>Using double-sided scotch tape, attach the two coverslips together. Cut the tape ~3 cm in length, and 0.3 cm in width. Attach the tape on 22x40 mm coverslip leaving a 1.6 cm channel in the middle. Use a pipette tip to gently press down on the coverslip to ensure that the tape has properly adhered.</li>
</ol>
2. Magnetic Tweezers Setup and Calibration
<ol>
	<li>Magnetic tweezers were setup using permanent rare-earth magnets as described previously10,12. An aluminum L-bracket was constructed with a 1 mm diameter pinhole and mounted on a vertical z-translator (Thorlabs) to modulate the height of the tweezers. Two permanent rare-earth magnets were attached to the bracket on either side of the pinhole to create the magnetic trap (Figure 1).</li>
	<li>DNA preparation: DNA from lambda phage labeled at its 5' and 3' ends with biotin and digoxigenin (IDT DNA, San Diego, CA) was used to calibrate the magnetic tweezers.
	<ol>
		<li>Mix 50 &mu;L of 4 nM &lambda;-DNA with 2 &mu;L of 40 nM biotin conjugated oligonucleotide and heat at 60 &deg;C for 10 minutes. Then slowly cool at room temperature over 1 hour.</li>
		<li>Add DNA ligase according to manufacturer's specifications and ligate at room temperature for 3 hours.</li>
		<li>Add 2 &mu;L of 400 nM digoxigenin conjugated oligonucleotide and incubate at room temperature for 45 minutes.</li>
		<li>Add 1.5 &mu;L of 10 mM ATP and continue ligation of the digoxigenin conjugated oligonucleotide to the &lambda;-DNA for 3 hours.</li>
		<li>Remove the excess oligonucleotides and the DNA ligase using 100 kDa spin filters (Microcon Ultracell YM-100). A total of 1000-fold buffer exchange in TE buffer is adequate to remove the excess oligos. Note: It is important not to use spin speeds &gt;500xG because it will lead to the shearing of the DNA.</li>
		<li>Examine a sample of the DNA via electrophoresis on a 0.7% agarose gel to ensure that the DNA is not denatured or sheared.</li>
		<li>Measure concentration of DNA. The use of a Nanodrop UV-Vis spectrometer (Thermo Scientific) is recommended because the available sample volume may be very small.</li>
	</ol>
	</li>
	<li>Prepare flow cells as described in section 1.</li>
	<li>For the attachment of the DNA to the flow cell, add anti-digoxigenin antibody (sheep IgG; 1 &mu;g mL-1; Roche Bioscience, Manheim, Germany) and allow it to adhere to the glass surface for 15 minutes.</li>
	<li>Passivate the surface of the flow cell to prevent non-specific sticking of DNA by adding 2 mg mL-1 BSA, 0.1% Tween-20 in 1x PBS. Incubate at room temperature for 45 minutes.</li>
	<li>Repeat step 2.5.</li>
	<li>Add 50 pM of fuctionalized &lambda;-DNA and allow to attach to the coverslip for 15 minutes.</li>
	<li>Wash away excess DNA with 1x PBS.</li>
	<li>Add streptavidin-coated superparamagnetic beads (1 &mu;m beads = 2 &mu;g mL-1; 2.8 &mu;m beads = 20 &mu;g mL-1) and allow them to attach to the immobilized DNA for 15 minutes.</li>
	<li>Wash away excess beads with 1x PBS.</li>
	<li>Magnetic Tweezers Calibration
	<ol>
		<li>Move the permanent magnets to a position that is far away from the sample surface (&gt;15 mm).</li>
		<li>Place the prepared flow cell in the microscope.</li>
		<li>Move the permanent magnets into position above the flow cell.</li>
		<li>Choose the focal plane of &lambda;-DNA functionalized beads by focusing on the beads away from both glass surfaces. The correct focal plane is one in which the desired beads have a bright center.</li>
		<li>Take a video of bead motion at 80 Hz or greater for 500 frames.</li>
		<li>Move the magnet closer to the sample surface. For distances beyond 5 mm, move in 1 mm increments. For distances between 1 and 5 mm, move in 0.5 mm increments. For distances less than 1 mm, move in 0.25 mm increments.</li>
		<li>Repeat steps 2.11.5 and 2.11.6 at each new magnet position.</li>
		<li>For each data set, track the center of the beads using a 2D Gaussian fit.</li>
		<li>Calculate the force via Brownian fluctuations using:</li>
	</ol>
	 <img alt="Equation 1" src="/files/ftp_upload/3520/3520eq1.jpg" /> 
	where kBT is the Boltzmann thermal energy, L is the length of &lambda;-DNA, and &lt;x2&gt; is the variance in centroid position. 
	<ol start="10">
		<li>Confirm the accuracy of the forces by checking the force calculation via a power spectrum Lorentzian fit in order to find the rolloff frequency. Applied force is related to the rolloff frequency by the relation:</li>
	</ol>
	 <img alt="Equation 2" src="/files/ftp_upload/3520/3520eq2.jpg" /> 
	Where &fnof;0 is the rolloff frequency, a is the bead radius, and &mu; is the fluid viscosity12. </li>
</ol>
3. Collagen Peptide Attachment to Flow Cells
In order to provide a concrete example for the application of our methodology, we describe recent work in our laboratory that characterizes the proteolysis of a trimeric collagen model peptide. We anticipate that this general approach may be broadly applied to other proteins and polynucleotides.
<ol>
	<li>The collagen model peptide consists of a N-terminal 6x His-tag for purification, followed by a 5x myc tag, (GPP)10 to enforce triple helix formation, the collagen &alpha;1 residues 772-786 (GPQGIAGQRGVVGL), which form the MMP-1 recognition site, the foldon sequence (GSGYIPEAPRDGQAYVRKDGEWVLLSTFL), and a C-terminal KKCK to facilitate labeling with biotin-maleimide. Foldon derives from the T4-phage protein fibritin, and stabilizes collagen model trimers when fused at either the N- or C-terminus13,14.</li>
	<li>The collagen peptide and MMP-1 proteins were expressed and purified as described previously4,15.</li>
	<li>Add anti-myc (15 &mu;g mL-1) to the flow cell and incubate at room temperature for 20 minutes to allow the anti-myc to attach to the surface of the flow cell.</li>
	<li>Passivate the flow cell to prevent non-specific attachment of proteins using 5 mg mL-1 BSA in the same way as described in step 2.5.</li>
	<li>Add 150 pM collagen peptide and allow it to attach to the antibody via the myc tag for 45 minutes.</li>
	<li>Wash away excess collagen using PBS buffer.</li>
	<li>Add streptavidin coated superparamagnetic beads (1 &mu;m beads = 2 &mu;g mL-1and 2.8 &mu;m beads = 20 &mu;g mL-1) and allow them to attach to the collagen molecules for 45 minutes. The 1 &mu;m beads should be diluted to the desired concentration in PBS. The 3 &mu;m beads should be separated from the solution using strong magnets, then resolubilized in PBS. This process should be repeated 3 times. We noticed that this step is essential to get the 3 &mu;m beads to bind to surface-immobilized collagen peptide.</li>
</ol>
4. Force Proteolysis Assay
<ol>
	<li>Once the flow cell is assembled with collagen peptide and magnetic beads, image the flow cell in the magnetic trap under low, ~1 pN force. In our experience, a subpopulation of the beads is sometimes weakly adhered to the surface in a non-specific fashion. The application of modest force ensures that these beads are liberated.</li>
	<li>Add activated enzyme to the flow cell. In this case MMP-1 was pre-activated by adding 3.5 mM APMA (4-aminophenylmercuric acetate) and incubated at 37 &deg;C for 3 hours. The activation was verified using SDS-PAGE.</li>
	<li>As soon as the flow cell is re-introduced into the magnetic tweezers apparatus, record a video spanning a few fields of view, typically yielding several hundred attached beads. This will be t = 0.</li>
	<li>Repeat the same process of recording several fields of view (while returning to the same field of view) at regular time points, till all the beads detach or no further proteolysis is discernable.</li>
	<li>Kinetic data analysis
	<ol>
		<li>Count the beads in the various fields of view at different time points and average them. Normalize the number of beads by dividing the average number of beads at each time point by the number of beads at t = 0. This ensures that we can compare the rates of proteolysis from different experiments, because the absolute number of beads will differ from experiment to experiment.</li>
		<li>Plot the ratio as a function of time. In this case, we fitted it to an exponential decay plus a constant.</li>
	</ol>
	 <img alt="Equation 3" src="/files/ftp_upload/3520/3520eq3.jpg" /> 
	where &fnof;(t) is the ratio of the beads attached (or collagen molecules unproteolyzed), t is time, a is the fraction of beads attached by a collagen tether, b is the decay rate constant, and c is the fraction of beads that are attached non-specifically. 
	<ol start="3">
		<li>The same experiment is repeated at varying force and MMP-1 concentrations to elucidate the effect of force on collagen proteolysis by MMP-1.</li>
	</ol>
	</li>
</ol>
5. Representative Results
The above protocol describes a novel use of magnetic tweezers (Figure 1) for studying the effect of force on enzymatic proteolysis. We calibrated the tweezers for 1 &mu;m and 3 &mu;m beads using both the magnitude of the observed Brownian fluctuations and calculation of the roll-off frequency at varying magnet positions (Figure 2). In the force proteolysis experiments, the setup is similar except that the DNA is replaced with collagen (Figures 3, 4). The normalized number of beads remaining can be plotted as function of time to find the proteolysis rates (Figure 5), and this process can be repeated for varying enzyme concentrations and forces.
<img alt="Figure 1" src="/files/ftp_upload/3520/3520fig1.jpg" /> 
Figure 1. Schematic of the magnetic trap calibration process (not to scale). Two permanent rare earth magnets create a magnetic field that pulls on the superparamagnetic bead. Translating the magnet up and down adjusts the applied force. The beads are imaged using a conventional bright-field microscope with the light passing through a pinhole between the two magnets. Inset: Image taken with a 40x air objective. The sharp, round spots correspond to the beads attached to the coverslip surface. The out-of-focus objects are detached beads.
<img alt="Figure 2" src="/files/ftp_upload/3520/3520fig2.jpg" /> 
Figure 2. Plots of the calibrated force as a function of magnet distance from the sample surface for 1 &mu;m (left) and 2.8 &mu;m (right) beads. Data were fit to the empirical function <img alt="Equation4" src="/files/ftp_upload/3520/3520eq4.jpg" />, where x is the distance from the magnet. a = 31.8, b= 5.61, and c = 4.39 for the 1 &mu;m beads and a = 140, b = 3.10 and c = 1.86 for the 3 &mu;m beads. These values are specific to our specific instrument and experimental geometry, and each instrument should be calibrated individually. The error bars at each point represent a ~10% variability in applied force on a bead to bead basis, due to the variability in bead size. The force applied is proportional to the volume of the beads, and the bead volume varies ~9% over the mean (according to manufacturer specifications).
<img alt="Figure 3" src="/files/ftp_upload/3520/3520fig3.jpg" /> 
Figure 3. Force proteolysis assay setup (Not to scale). The collagen model trimer is attached to the surface of the coverslip via myc/anti-myc conjugation. The streptavidin-coated superparamagnetic beads are attached to the collagen trimers via a biotin-streptavidin linkage. Activated MMP-1 cuts the collagen over time, causing the beads to detach from the surface and move away from the focal plane.
<img alt="Figure 4" src="/files/ftp_upload/3520/3520fig4.jpg" /> 
Figure 4. Schematic cartoon of proteolysis as a function of time. The cartoon shows a sample field of view over time as proteolysis occurs. Over time, MMP-1 cuts the collagen and the beads detach and move away from the focal plane under the influence of the magnetic field.
<img alt="Figure 5" src="/files/ftp_upload/3520/3520fig5.jpg" /> 
Figure 5. Proteolysis rates depend on applied force16. Shown are data collected at 1.0 pN (3 &mu;M MMP-1; black), 6.2 pN (3 &mu;M MMP-1; red) and 13 pN (0.2 &mu;M MMP-1; magenta). The rates of proteolysis (fit parameters) are: 0.22 &plusmn; 0.02 min-1 (1 pN), 0.46 &plusmn; 0.09 min-1 (6.2 pN), and 2.08 &plusmn; 0.18 min-1 (13 pN). The fraction of beads unproteolyzed at long time points (&gt; 15 minutes) remains approximately constant at ~0.25 across different experiments. Error bars correspond to the Poisson statistics reflecting the number of observations at each time point. The error at each time point for n beads is n1/2. The error in fraction of beads attached was calculated by error propagation.1 &mu;m beads were used for 1.0 pN and 6.2 pN experiments and 2.8 &mu;m beads were used for 13 pN experiments.

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No conflicts of interest declared.

This protocol describes a new use for a classical single molecule technique. Magnetic tweezers allow medium to high-throughput single molecule assays in a cost-efficient manner. However, like all experimental techniques there are challenges and potential pitfalls.
Limitations of magnetic tweezers
Compared to an optical trap the spatial and temporal resolution of a MT apparatus is low. Moreover, the forces generated by the simple MT described here are 30 pN or less, sig...

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 </tr>
 <tr>
 <td>Micro Cover Glass #1.5 (22x22)</td>
 <td>VWR</td>
 <td>48366-067</td>
 </tr>
 <tr>
 <td>Micro Cover Glass #1.5 (22x40)</td>
 <td>VWR</td>
 <td>48393-048</td>
 </tr>
 <tr>
 <td>Lambda DNA</td>
 <td>Invitrogen</td>
 <td>25250-010</td>
 </tr>
 <tr>
 <td>T4 DNA Ligase</td>
 <td>Invitrogen</td>
 <td>15224-041</td>
 </tr>
 <tr>
 <td>Microcon Ultracel YM-100</td>
 <td>Millipore</td>
 <td>42413</td>
 </tr>
 <tr>
 <td>Anti-Digoxigenin</td>
 <td>Roche Diagnostics</td>
 <td>11-333-089-001</td>
 </tr>
 <tr>
 <td>Tween 20</td>
 <td>Sigma</td>
 <td>P9416-100ML</td>
 </tr>
 <tr>
 <td>Anti-myc Antibody</td>
 <td>Invitrogen</td>
 <td>46-0603</td>
 </tr>
 <tr>
 <td>Bovine Serum Albumin</td>
 <td>Sigma</td>
 <td>B4287-5G</td>
 </tr>
 <tr>
 <td>Dynabeads M-280 Streptavidin</td>
 <td>Invitrogen</td>
 <td>658.01D</td>
 </tr>
 <tr>
 <td>Dynabeads MyOne T1 Streptavidin</td>
 <td>Invitrogen</td>
 <td>658.01D</td>
 </tr>
 <tr>
 <td>p-Aminophenylmercuric Acetate</td>
 <td>Calbiochem</td>
 <td>164610</td>
 </tr>
 <tr>
 <td>Biotin-Maleimide</td>
 <td>Sigma Aldrich</td>
 <td>B1267</td>
 </tr>
 <tr>
 <td>Biotin labeled oligo</td>
 <td>IDT DNA</td>
 <td>Custom synthesis</td>
 </tr>
 <tr>
 <td>Digoxigenin labeled oligo</td>
 <td>IDT DNA</td>
 <td>Custom synthesis</td>
 </tr>
 <tr>
 <td>Collagen peptide gene</td>
 <td>DNA 2.0</td>
 <td>Custom synthesis</td>
 </tr>
 <tr>
 <td>MMP-1 cDNA</td>
 <td>Harvard Plasmid Database</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>z-translator</td>
 <td>Thorlabs</td>
 <td>MTS50</td>
 </tr>
 <tr>
 <td>Servo controller for translator</td>
 <td>Thorlabs</td>
 <td>TDC001</td>
 </tr>
 </tbody>
 </table>

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.

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Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

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 ...

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Research

JoVE Journal

Bioengineering

11.5K Views.  Stanford University. В этой статье мы рассмотрим использование магнитного пинцета, чтобы изучить влияние силы на ферментативную протеолиза на одном уровне молекулы хорошо подходит для распараллеливания образом.

Video: Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

Stretching Short Sequences of DNA with Constant Force Axial Optical Tweezers

Single-molecule techniques for stretching DNA of contour lengths less than a kilobase are fraught with experimental difficulties. However, many interesting biological events such as histone binding and protein-mediated looping of DNA1,2, occur on this length scale. In recent years, the mechanical properties of DNA have been shown to play a significant role in fundamental cellular processes like the packaging of DNA into compact nucleosomes and chromatin fibers3,4. Clearly, it is then important to understand the mechanical properties of short stretches of DNA. In this paper, we provide a practical guide to a single-molecule optical tweezing technique that we have developed to study the mechanical behavior of DNA with contour lengths as short as a few hundred basepairs.
The major hurdle in stretching short segments of DNA is that conventional optical tweezers are generally designed to apply force in a direction lateral to the stage5,6, (see Fig. 1). In this geometry, the angle between the bead and the coverslip, to which the DNA is tethered, becomes very steep for submicron length DNA. The axial position must now be accounted for, which can be a challenge, and, since the extension drags the microsphere closer to the coverslip, steric effects are enhanced. Furthermore, as a result of the asymmetry of the microspheres, lateral extensions will generate varying levels of torque due to rotation of the microsphere within the optical trap since the direction of the reactive force changes during the extension.
Alternate methods for stretching submicron DNA run up against their own unique hurdles. For instance, a dual-beam optical trap is limited to stretching DNA of around a wavelength, at which point interference effects between the two traps and from light scattering between the microspheres begin to pose a significant problem. Replacing one of the traps with a micropipette would most likely suffer from similar challenges. While one could directly use the axial potential to stretch the DNA, an active feedback scheme would be needed to apply a constant force and the bandwidth of this will be quite limited, especially at low forces.
We circumvent these fundamental problems by directly pulling the DNA away from the coverslip by using a constant force axial optical tweezers7,8. This is achieved by trapping the bead in a linear region of the optical potential, where the optical force is constant-the strength of which can be tuned by adjusting the laser power. Trapping within the linear region also serves as an all optical force-clamp on the DNA that extends for nearly 350 nm in the axial direction. We simultaneously compensate for thermal and mechanical drift by finely adjusting the position of the stage so that a reference microsphere stuck to the coverslip remains at the same position and focus, allowing for a virtually limitless observation period.

We illustrate the use of a constant force axial optical tweezers to explore the mechanical properties of short DNA molecules. By stretching DNA axially, we minimize steric hindrances and artifacts arising in conventional lateral manipulation, allowing us to study DNA molecules as short as ~100 nm.

Single-molecule techniques for stretching DNA of contour lengths  less than a kilobase are fraught with experimental difficulties. However, many ...

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 ...

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 ...

Magnetic Tweezers for the Measurement of Twist and Torque

Single-molecule techniques make it possible to investigate the behavior of individual biological molecules in solution in real time. These techniques include so-called force spectroscopy approaches such as atomic force microscopy, optical tweezers, flow stretching, and magnetic tweezers. Amongst these approaches, magnetic tweezers have distinguished themselves by their ability to apply torque while maintaining a constant stretching force. Here, it is illustrated how such a &#8220;conventional&#8221; magnetic tweezers experimental configuration can, through a straightforward modification of its field configuration to minimize the magnitude of the transverse field, be adapted to measure the degree of twist in a biological molecule. The resulting configuration is termed the freely-orbiting magnetic tweezers. Additionally, it is shown how further modification of the field configuration can yield a transverse field with a magnitude intermediate between that of the &#8220;conventional&#8221; magnetic tweezers and the freely-orbiting magnetic tweezers, which makes it possible to directly measure the torque stored in a biological molecule. This configuration is termed the magnetic torque tweezers. The accompanying video explains in detail how the conversion of conventional magnetic tweezers into freely-orbiting magnetic tweezers and magnetic torque tweezers can be accomplished, and demonstrates the use of these techniques. These adaptations maintain all the strengths of conventional magnetic tweezers while greatly expanding the versatility of this powerful instrument.

Magnetic tweezers, a powerful single-molecule manipulation technique, can be adapted for the direct measurements of the twist (using a configuration called freely-orbiting magnetic tweezers) and torque (using a configuration termed magnetic torque tweezers) in biological macromolecules. Guidelines for performing such measurements are given, including applications to the study of DNA and associated nucleo-protein filaments.

Single-molecule techniques make it possible to investigate the behavior of individual biological molecules in solution in real time. These techniques ...

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., ...

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 ...

Rapid Scan Electron Paramagnetic Resonance Opens New Avenues for Imaging Physiologically Important Parameters In Vivo

We demonstrate a superior method of 2D spectral-spatial imaging of stable radical reporter molecules at 250 MHz using rapid-scan electron-paramagnetic-resonance (RS-EPR), which can provide quantitative information under in vivo conditions on oxygen concentration, pH, redox status and concentration of signaling molecules (i.e., OH&#8226;, NO&#8226;). The RS-EPR technique has a higher sensitivity, improved spatial resolution (1 mm), and shorter acquisition time in comparison to the standard continuous wave (CW) technique. A variety of phantom configurations have been tested, with spatial resolution varying from 1 to 6 mm, and spectral width of the reporter molecules ranging from 16 &#181;T (160 mG) to 5 mT (50 G). A cross-loop bimodal resonator decouples excitation and detection, reducing the noise, while the rapid scan effect allows more power to be input to the spin system before saturation, increasing the EPR signal. This leads to a substantially higher signal-to-noise ratio than in conventional CW EPR experiments.

Rapid Scan Electron Paramagnetic Resonance Opens New Avenues for Imaging Physiologically Important Parameters In Vivo

A new electron paramagnetic resonance (EPR) method, rapid scan EPR (RS-EPR), is demonstrated for 2D spectral spatial imaging which is superior to the traditional continuous wave (CW) technique and opens new venues for in vivo imaging. Results are demonstrated at 250 MHz, but the technique is applicable at any frequency.

We demonstrate a superior method of 2D spectral-spatial imaging of stable radical reporter molecules at 250 MHz using rapid-scan ...

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.

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 ...

Fabrication of Zero Mode Waveguides for High Concentration Single Molecule Microscopy

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to nanomolar ranges, several orders of magnitude less than many physiological ligand concentrations. Optical nanostructures called zero mode waveguides (ZMWs), which are 100&#8722;200 nm in diameter apertures fabricated in a thin conducting metal such as aluminum or gold, allow imaging of individual molecules at micromolar concentrations of fluorophores by confining visible light excitation to zeptoliter effective volumes. However, the need for expensive and specialized nanofabrication equipment has precluded the widespread use of ZMWs. Typically, nanostructures such as ZMWs are obtained by direct writing using electron beam lithography, which is sequential and slow. Here, colloidal, or nanosphere, lithography is used as an alternative strategy to create nanometer-scale masks for waveguide fabrication. This report describes the approach in detail, with practical considerations for each phase. The method allows thousands of aluminum or gold ZMWs to be made in parallel, with final waveguide diameters and depths of 100&#8722;200 nm. Only common lab equipment and a thermal evaporator for metal deposition are required. By making ZMWs more accessible to the biochemical community, this method can facilitate the study of molecular processes at cellular concentrations and rates.

Described here is a nanosphere lithography method for parallel fabrication of zero mode waveguides, which are arrays of nanoapertures in a metal-clad glass microscopy coverslip for single molecule imaging at nano- to micromolar concentrations of fluorophores. The method takes advantage of colloidal crystal self-assembly to create a waveguide template.

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to ...