We thank Beijing Wu for use of her data. This work was supported by an NSF CAREER award to J.E.K.

1. Preparation of ~50 nm Diameter Small Unilamellar Vesicles (SUVs) for Membrane Protein Folding
<ol>
 <li>A solution of 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) lipids in chloroform is purchased and aliquotted into clean glass vials in 20 mg per vial quantities for storage. A layer of nitrogen gas is added to each vial to prevent lipid oxidation, and the vials are sealed with caps and parafilm. The vials are stored in a -20&deg; C freezer until use.</li>
 <li>A single vial that contains an aliquot of 20 mg lipid in chloroform is used for each experiment. The contents of the vial are dried using a stream of nitrogen gas for 1 hour until no solvent remains.</li>
 <li>Dried lipids are resuspended in 1 mL of 20 mM potassium phosphate buffer (pH = 7.3) using a water bath sonicator for ~ 30 seconds. This solution of suspended lipid will appear cloudy, and is transferred to a plastic 15-mL tube with a conical bottom. Another 1 mL aliquot of phosphate buffer is added to the empty glass vial that originally contained lipid, and sonication is repeated; the solution is then added to the same 15-mL tube. This process is repeated a total of 4 times until the final volume in the tube is 4 mL, resulting in a lipid concentration of 5 mg/mL in this stock vesicle solution.</li>
 <li>The 4 mL volume of lipid vesicle solution is placed in a warm (~ 30&deg; C) water bath, and sonicated using an ultrasonicator microtip for 1 hour at 50% duty cycle. The purpose of the water bath is twofold. First, it prevents the lipid solution from becoming too hot because of the sonication process. Second, the warm bath ensures that the temperature of the aqueous lipid solution remains above the bilayer phase transition temperature during sonication; for DMPC, this transition temperature is ~23&deg; C. </li>
 <li>The sonicated solution is passed through a 0.22 &mu;m syringe filter to remove debris from the sonicator tip, and this filtered solution is equilibrated overnight at 37&deg; C.</li>
</ol>
2. Sample Preparation for Initial Fluorescence Unfolding Curve
<ol>
 <li>A stock solution of 10 M urea in 20 mM phosphate buffer (pH 7.3) is made. It is essential that water be added to the solid (and not vice versa) because the large quantity of urea occupies a significant volume in the solution. This urea solution can be made by adding solutes to a clean, empty bottle and adding water up to the intended final volume. This solution may require warming in a water bath or on a hot plate with stirring to dissolve the solute. Urea is hygroscopic and therefore, the actual concentration of the urea stock solution should be determined using refractive index.1</li>
 <li>A stock buffer solution of 20 mM phosphate (pH 7.3) is made.</li>
 <li>A stock solution of unfolded protein is prepared. This stock protein solution should have a concentration of ~200 &mu;M protein in 8 M urea, 20 mM phosphate buffer.</li>
 <li>Samples for fluorescence studies are prepared in the following manner. Appropriate volumes of stock protein (section 2.3), stock lipid solution (5 mg/mL lipid, section 1), stock 10 M urea solution (section 2.1), and stock phosphate buffer (section 2.2) are combined to make samples containing ~4 &mu;M protein, 1 mg/mL lipid, and 0 to 8 M urea in 1 M increments. The total volume of each sample is 200 &mu;L. An example spreadsheet (Table 1) is included that lists volumes required using a 200 &mu;M stock protein solution.</li>
 <li>Blank samples are made as described in section 2.4, however, protein is not added. In place of protein, 4 &mu;L of an 8 M urea solution may be added so concentrations of other components are identical to those in section 2.4. These blanks are used to subtract scattering and other background signal that appears in the fluorescence spectra of protein. </li>
 <li>Samples from sections 2.4 and 2.5 are allowed to incubate at 37&deg; C for at least 2 hours before measurement of fluorescence spectra to ensure complete folding and equilibration.</li>
</ol>
3. Measurement of Fluorescence Spectra
Tryptophan fluorescence of each sample and blank is measured using a steady-state fluorometer. Fluorescence spectra should be recorded with excitation wavelength of 290 nm to avoid excitation of tyrosine residues, and scanned from 305 to 500 nm. Typical entrance and exit bandpass is 3 nm. The wavelength increment and integration time can be optimized for signal-to-noise ratio. Typical values for wavelength increment and integration time are 1 nm/step and 0.5 sec/step, respectively. Membrane proteins generally fold into synthetic lipids only when the temperature is above the bilayer phase transition temperature. Therefore, in this case of DMPC, the temperature of the sample is held constant at 30 &deg;C. A microvolume fused silica cuvette with 160 &mu;L capacity is utilized in these experiments.
4. Generation of Initial Unfolding Curve and Estimation of Gibbs Free Energy of Membrane Protein Unfolding
<ol>
 <li>Fluorescence spectra in the form of an x-y dataset are loaded into software such as Igor Pro, Matlab, Origin, or Excel.</li>
 <li>Signal from the urea and lipid vesicle background must be subtracted using the following procedure: 
 Spectrum A = raw fluorescence spectrum of protein in the presence of lipid vesicles and specific concentration of urea. 
 Spectrum B = raw fluorescence spectrum of corresponding blank sample that contains lipid vesicles and same urea concentration as that in spectrum A; no protein is in the blank samples. 
 Corrected Spectrum = Spectrum A - C*spectrum B. C is a scalar typically equal to 1.0. In some cases, C is less than or greater than 1.0 because of scattering issues that cause over- or under-subtraction near the ~305-320 nm region.</li>
 <li>The wavelength of maximum fluorescence, &lambda;max, is tabulated from corrected spectra for each urea concentration. A typical value for fully unfolded membrane protein is 350 nm while that of folded protein is ~330 nm. Tabulated wavelengths are converted to fraction unfolded by converting the range of &lambda;max values (~330 to ~350 nm) to the range 0.0 to 1.0 to correlate the &lambda;max value to fraction of unfolded population. For example, a &lambda;max value of 330 nm corresponds to 0.0, 350 nm corresponds to 1.0, and 340 nm corresponds to 0.5 in terms of fraction unfolded. This fraction of unfolded protein is then plotted against urea concentration. As stated above, the urea concentration should be determined experimentally by measuring refractive index.</li>
 <li>We assume that the protein can exist in one of two states: folded or unfolded. The plot of fraction unfolded, f, versus urea concentration, C, can be fit to the equation:2 
 <img src="/files/ftp_upload/2669/2669eq2.jpg" alt="Equation 2" /> 
 The coefficient m corresponds to the rate of change of the free energy with respect to denaturant concentration, and Cm corresponds to the midpoint urea concentration at which the folded population equals the unfolded population. These values are the fitting variables obtained from least-squares fitting by data analysis software. R (0.001987 kcal/mol &bull; K) is the gas law constant, and T is temperature (303 K).</li>
 <li>The coefficients obtained from the fit are utilized to determine the Gibbs free energy (kcal/mol) of unfolding in the absence of urea: <img src="/files/ftp_upload/2669/2669eq3.jpg" alt="Equation 3"/>.</li>
</ol>
5. Optimized Unfolding Curve for More Precise Measurement of Gibbs Free Energy of Unfolding.
<ol>
 <li>The unfolding curve described above reveals the range of urea concentrations that causes the greatest change in the &lambda;max. This range is typically small, and the majority of unfolding occurs within this small range. In order to precisely determine the free energy of unfolding, additional samples are analyzed in this region to obtain a plot that contains many data points necessary for optimized least-squares fitting. For example, if the protein unfolds between 2 and 4 M urea, samples can be made that contain 2 - 4 M urea in 0.2 M increments to more precisely reveal the shape of this important region in the unfolding curve. It is not necessary to measure blank spectra at each urea concentration; it is sufficient to measure a blank spectrum every ~0.5 M step.</li>
 <li>The free energy of unfolding obtained from this plot may differ slightly from that obtained in the original unfolding curve, but reflects a more precise value.</li>
</ol>
6. Representative Results:
<table border="1">
 <tbody>
 <tr>
 <td>Sample</td>
 <td>final [Urea], M</td>
 <td>stock protein (&mu;L)</td>
 <td>stock lipid (&mu;L)</td>
 <td>stock 10 M urea (&mu;L)</td>
 <td>stock buffer (&mu;L)</td>
 </tr>
 <tr>
 <td>1</td>
 <td>~ 0.16</td>
 <td>4</td>
 <td>40</td>
 <td>0</td>
 <td>156</td>
 </tr>
 <tr>
 <td>2</td>
 <td>1</td>
 <td>4</td>
 <td>40</td>
 <td>20</td>
 <td>136</td>
 </tr>
 <tr>
 <td>3</td>
 <td>2</td>
 <td>4</td>
 <td>40</td>
 <td>40</td>
 <td>116</td>
 </tr>
 <tr>
 <td>4</td>
 <td>3</td>
 <td>4</td>
 <td>40</td>
 <td>60</td>
 <td>96</td>
 </tr>
 <tr>
 <td>5</td>
 <td>4</td>
 <td>4</td>
 <td>40</td>
 <td>80</td>
 <td>76</td>
 </tr>
 <tr>
 <td>6</td>
 <td>5</td>
 <td>4</td>
 <td>40</td>
 <td>100</td>
 <td>56</td>
 </tr>
 <tr>
 <td>7</td>
 <td>6</td>
 <td>4</td>
 <td>40</td>
 <td>120</td>
 <td>36</td>
 </tr>
 <tr>
 <td>8</td>
 <td>7</td>
 <td>4</td>
 <td>40</td>
 <td>140</td>
 <td>16</td>
 </tr>
 </tbody>
 </table>
Table 1. Volume of stock solutions required to make fluorescence samples
<img src="/files/ftp_upload/2669/2669fig1.jpg" alt="Figure 1" /> 
Figure 1. Tryptophan fluorescence spectra of ~ 5 &mu;M representative membrane protein that contains a single tryptophan residue. (A) Raw fluorescence spectra of protein (spectrum A from 4.2); (B) Raw fluorescence spectra of blank (spectrum B from 4.2); (C) Corrected spectrum from 4.2.
<img src="/files/ftp_upload/2669/2669fig2.jpg" alt="Figure 2" /> 
Figure 2. Corrected tryptophan fluorescence spectra of membrane protein from Figure 1 for numerous urea concentrations.
<img src="/files/ftp_upload/2669/2669fig3.jpg" alt="Figure 3" /> 
Figure 3. Unfolding curve obtained from data in Figure 2, with fit to equation in 4.4. The Gibbs free energy is calculated according to section 4.5.

<ol>
	<li>Shirley, B. A., Shirley, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Protein Folding and Stability. , 177-190 (1995).</li><li>Pace, C. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+and+Analysis+of+Urea+and+Guanidine+Hydrochloride+Denaturation+Curves.">Determination and Analysis of Urea and Guanidine Hydrochloride Denaturation Curves.</a> Methods Enzymol. 131, 266-279 (1986).</li><li>Lau, F. W., Bowie, J. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Method+for+Assessing+the+Stability+of+a+Membrane+Protein.">A Method for Assessing the Stability of a Membrane Protein.</a> Biochemistry. 36, 5884-5892 (1997).</li><li>Burgess, N. K., Dao, T. P., Stanley, A. M., Fleming, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=&#946;-Barrel+Proteins+that+Reside+in+the+Escherichia+coli+Outer+Membrane+in+Vivo+Demonstrate+Varied+Folding+Behavior+in+Vitro.">&#946;-Barrel Proteins that Reside in the Escherichia coli Outer Membrane in Vivo Demonstrate Varied Folding Behavior in Vitro.</a> J. Biol. Chem. 283, 26748-26758 (2008).</li><li>Booth, P. J., Curnow, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Folding+scene+investigation:+membrane+proteins.">Folding scene investigation: membrane proteins.</a> Curr. Opin. Struct. Biol. 19, 8-13 (2009).</li><li>Hong, H., Tamm, L. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elastic+coupling+of+integral+membrane+protein+stability+to+lipid+bilayer+forces.">Elastic coupling of integral membrane protein stability to lipid bilayer forces.</a> Proc. Natl. Acad. Sci. U.S.A. 101, 4065-4070 (2004).</li><li>Sanchez, K. M., Gable, J. E., Schlamadinger, D. E., Kim, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+Tryptophan+Microenvironment,+Soluble+Domain,+and+Vesicle+Size+on+the+Thermodynamics+of+Membrane+Protein+Folding:+Lessons+from+the+Transmembrane+Protein+OmpA.">Effects of Tryptophan Microenvironment, Soluble Domain, and Vesicle Size on the Thermodynamics of Membrane Protein Folding: Lessons from the Transmembrane Protein OmpA.</a> Biochemistry. 47, 12844-12852 (2008).</li></ol>

No conflicts of interest declared.

The current protocol describes the generation of unfolding curves of membrane-associated proteins and peptides that contain tryptophan residues. Here, it is assumed that the tryptophan fluorescence reflects whether the protein is folded and inserted into synthetic lipid vesicles, or unfolded in solution. Additional assumptions, such as two-state folding and linear dependence of free energy with denaturant concentration, are made in the current report; modification of these assumptions result in different equations.<sup...

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		<th>Material Name</th>
		<th>Type</th>
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		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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<tr>
<td>DMPC</td>
<td>&nbsp;</td>
<td>Avanti Polar Lipids</td>
<td>850345C</td>
<td>&nbsp;</td>
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<td>Urea</td>
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<td>MP Biochemicals</td>
<td>04821527</td>
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<td>&nbsp;</td>
<td>Fisher</td>
<td>P288</td>
<td>&nbsp;</td>
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thermodynamics of membrane protein folding measured by fluorescence spectroscopy

Membrane protein folding is an emerging topic with both fundamental and health-related significance. The abundance of membrane proteins in cells underlies the need for comprehensive study of the folding of this ubiquitous family of proteins. Additionally, advances in our ability to characterize diseases associated with misfolded proteins have motivated significant experimental and theoretical efforts in the field of protein folding. Rapid progress in this important field is unfortunately hindered by the inherent challenges associated with membrane proteins and the complexity of the folding mechanism. Here, we outline an experimental procedure for measuring the thermodynamic property of the Gibbs free energy of unfolding in the absence of denaturant, &Delta;G&deg;H2O, for a representative integral membrane protein from E. coli. This protocol focuses on the application of fluorescence spectroscopy to determine equilibrium populations of folded and unfolded states as a function of denaturant concentration. Experimental considerations for the preparation of synthetic lipid vesicles as well as key steps in the data analysis procedure are highlighted. This technique is versatile and may be pursued with different types of denaturant, including temperature and pH, as well as in various folding environments of lipids and micelles. The current protocol is one that can be generalized to any membrane or soluble protein that meets the set of criteria discussed below.

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Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

This video article details the experimental procedure for obtaining the Gibbs free energy of membrane protein folding by tryptophan fluorescence.

Membrane protein folding is an emerging topic with both fundamental and health-related significance.  The abundance of membrane proteins in cells ...

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18.2K Views.  University of California San Diego - UCSD. This video article details the experimental procedure for obtaining the Gibbs free energy of membrane protein folding by tryptophan fluorescence.

Video: Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

Fluorescence detection methods for microfluidic droplet platforms

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4 Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that ...

Microfluidic Mixers for Studying Protein Folding

The process by which a protein folds into its native conformation is highly relevant to biology and human health yet still poorly understood. One reason for this is that folding takes place over a wide range of timescales, from nanoseconds to seconds or longer, depending on the protein1. Conventional stopped-flow mixers have allowed measurement of folding kinetics starting at about 1 ms. We have recently developed a microfluidic mixer that dilutes denaturant ~100-fold in ~8 &mu;s2. Unlike a stopped-flow mixer, this mixer operates in the laminar flow regime in which turbulence does not occur. The absence of turbulence allows precise numeric simulation of all flows within the mixer with excellent agreement to experiment3-4.
Laminar flow is achieved for Reynolds numbers Re &le;100. For aqueous solutions, this requires micron scale geometries. We use a hard substrate, such as silicon or fused silica, to make channels 5-10 &mu;m wide and 10 &mu;m deep (See Figure 1). The smallest dimensions, at the entrance to the mixing region, are on the order of 1 &mu;m in size. The chip is sealed with a thin glass or fused silica coverslip for optical access. Typical total linear flow rates are ~1 m/s, yielding Re~10, but the protein consumption is only ~0.5 nL/s or 1.8 &mu;L/hr. Protein concentration depends on the detection method: For tryptophan fluorescence the typical concentration is 100 &mu;M (for 1 Trp/protein) and for FRET the typical concentration is ~100 nM.
The folding process is initiated by rapid dilution of denaturant from 6 M to 0.06 M guanidine hydrochloride. The protein in high denaturant flows down a central channel and is met on either side at the mixing region by buffer without denaturant moving ~100 times faster (see Figure 2). This geometry causes rapid constriction of the protein flow into a narrow jet ~100 nm wide. Diffusion of the light denaturant molecules is very rapid, while diffusion of the heavy protein molecules is much slower, diffusing less than 1 &mu;m in 1 ms. The difference in diffusion constant of the denaturant and the protein results in rapid dilution of the denaturant from the protein stream, reducing the effective concentration of the denaturant around the protein. The protein jet flows at a constant rate down the observation channel and fluorescence of the protein during folding can be observed using a scanning confocal microscope5.

In this work we explain the fabrication and use of a microfluidic mixer capable of mixing two solutions in ~8 &mu;s. We also demonstrate the use of these mixers with spectroscopic detection using UV fluorescence and fluorescence resonance energy transfer (FRET).

The process by which a protein folds into its native conformation is highly relevant to biology and human health yet still poorly understood. One reason ...

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to identify and characterize their interactions are necessary5. To this end, we developed the Membrane Strep-protein interaction experiment, called Membrane-SPINE6. This technique combines in vivo cross-linking using the reversible cross-linker formaldehyde with affinity purification of a Strep-tagged membrane bait protein. During the procedure, cross-linked prey proteins are co-purified with the membrane bait protein and subsequently separated by boiling. Hence, two major tasks can be executed when analyzing protein-protein interactions (PPIs) of membrane proteins using Membrane-SPINE: first, the confirmation of a proposed interaction partner by immunoblotting, and second, the identification of new interaction partners by mass spectrometry analysis. Moreover, even low affinity, transient PPIs are detectable by this technique. Finally, Membrane-SPINE is adaptable to almost any cell type, making it applicable as a powerful screening tool to identify PPIs of membrane proteins.

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

A biochemical approach is described to identify in vivo protein-protein interactions (PPI) of membrane proteins. The method combines protein cross-linking, affinity purification and mass spectrometry, and is adaptable to almost any cell type or organism. With this approach, even the identification of transient PPIs becomes possible.

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to ...

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., ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

Preparation of Extracellular Matrix Protein Fibers for Brillouin Spectroscopy

Brillouin spectroscopy is an emerging technique in the biomedical field. It probes the mechanical properties of a sample through the interaction of visible light with thermally induced acoustic waves or phonons propagating at a speed of a few km/sec. Information on the elasticity and structure of the material is obtained in a nondestructive contactless manner, hence opening the way to in vivo applications and potential diagnosis of pathology. This work describes the application of Brillouin spectroscopy to the study of biomechanics in elastin and trypsin-digested type I collagen fibers of the extracellular matrix. Fibrous proteins of the extracellular matrix are the building blocks of biological tissues and investigating their mechanical and physical behavior is key to establishing structure-function relationships in normal tissues and the changes which occur in disease. The procedures of sample preparation followed by measurement of Brillouin spectra using a reflective substrate are presented together with details of the optical system and methods of spectral data analysis.

We present a protocol for the application of Brillouin light scattering spectroscopy to elastin and trypsin-purified type I collagen fibers of the extracellular matrix to extract their full elastic properties.

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Engineering 'Golden' Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and engineered variants of the green fluorescent protein from Aequorea victoria (avGFP) as well as homologs from other species already cover large parts of the optical spectrum, a spectral gap remains in the near-infrared region, for which avGFP-based fluorophores are not available. Red-shifted fluorescent protein (FP) variants would substantially expand the toolkit for spectral unmixing of multiple molecular species, but the naturally occurring red-shifted FPs derived from corals or sea anemones have lower fluorescence quantum yield and inferior photo-stability compared to the avGFP variants. Further manipulation and possible expansion of the chromophore&#39;s conjugated system towards the far-red spectral region is also limited by the repertoire of 20 canonical amino acids prescribed by the genetic code. To overcome these limitations, synthetic biology can achieve further spectral red-shifting via insertion of non-canonical amino acids into the chromophore triad. We describe the application of SPI to engineer avGFP variants with novel spectral properties. Protein expression is performed in a tryptophan-auxotrophic E. coli strain and by supplementing growth media with suitable indole precursors. Inside the cells, these precursors are converted to the corresponding tryptophan analogs and incorporated into proteins by the ribosomal machinery in response to UGG codons. The replacement of Trp-66 in the enhanced &#34;cyan&#34; variant of avGFP (ECFP) by an electron-donating 4-aminotryptophan results in GdFP featuring a 108 nm Stokes shift and a strongly red-shifted emission maximum (574 nm), while being thermodynamically more stable than its predecessor ECFP. Residue-specific incorporation of the non-canonical amino acid is analyzed by mass spectrometry. The spectroscopic properties of GdFP are characterized by time-resolved fluorescence spectroscopy as one of the valuable applications of genetically encoded FPs in life sciences.

Engineering &#039;Golden&#039; Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Synthetic biology enables the engineering of proteins with unprecedented properties using the co-translational insertion of non-canonical amino acids. Here, we presented how a spectrally red-shifted variant of a GFP-type fluorophore with novel fluorescence spectroscopic properties, termed &#34;gold&#34; fluorescent protein (GdFP), is produced in E. coli via selective pressure incorporation (SPI).

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and ...

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate (LAL) Assays

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause inflammation, fever, hypo- or hypertension, and, in extreme cases, can lead to tissue and organ damage that may become fatal. The amounts of endotoxin in pharmaceutical products, therefore, are strictly regulated. Among the methods available for endotoxin detection and quantification, the Limulus Amoebocyte Lysate (LAL) assay is commonly used worldwide. While any pharmaceutical product can interfere with the LAL assay, nano-formulations represent a particular challenge due to their complexity. The purpose of this paper is to provide a practical guide to researchers inexperienced in estimating endotoxins in engineered nanomaterials and nanoparticle-formulated drugs. Herein, practical recommendations for performing three LAL formats including turbidity, chromogenic and gel-clot assays are discussed. These assays can be used to determine endotoxin contamination in nanotechnology-based drug products, vaccines, and adjuvants.

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate LAL Assays

Detection of endotoxins in engineered nanomaterials represents one of the grand challenges in the field of nanomedicine. Here, we present a case study that describes the framework composed of three different LAL formats to estimate potential endotoxin contamination in nanoparticles.

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause ...

In Vesiculo Synthesis of Peptide Membrane Precursors for Autonomous Vesicle Growth

Compartmentalization of biochemical reactions is a central aspect of synthetic cells. For this purpose, peptide-based reaction compartments serve as an attractive alternative to liposomes or fatty acid-based vesicles. Externally or within the vesicles, peptides can be easily expressed and simplify the synthesis of membrane precursors. Provided here is a protocol for the creation of vesicles with diameters of ~200 nm based on the amphiphilic elastin-like polypeptides (ELP) utilizing dehydration-rehydration from glass beads. Also presented are protocols for bacterial ELP expression and purification via inverse temperature cycling, as well as their covalent functionalization with fluorescent dyes. Furthermore, this report describes a protocol to enable the transcription of RNA aptamer dBroccoli inside ELP vesicles as a less complex example for a biochemical reaction. Finally, a protocol is provided, which allows in vesiculo expression of fluorescent proteins and the membrane peptide, whereas synthesis of the latter results in vesicle growth.

Presented here are protocols for the creation of peptide-based small unilamellar vesicles capable of growth. To facilitate in vesiculo production of the membrane peptide, these vesicles are equipped with a transcription-translation system and the peptide-encoding plasmid.

Compartmentalization of biochemical reactions is a central aspect of synthetic cells. For this purpose, peptide-based reaction compartments serve as an ...

Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure single-molecular binding kinetics, assess mechanical properties of ligand-receptor interactions, visualize protein dynamic conformational changes and more excitingly elucidate receptor mediated cell mechanosensing mechanisms. More recently, BFP has been used to measure the spring constant of molecular bonds. This protocol describes the step-by-step procedure to perform molecular spring constant DFS analysis. Specifically, two BFP operation modes are discussed, namely the Bead-Cell and Bead-Bead modes. This protocol focuses on deriving spring constants of the molecular bond and cell from DFS raw data.

A biomembrane force probe (BFP) is an in situ dynamic force spectroscopy (DFS) technique. BFP can be used to measure the spring constant of molecular interactions on living cells. This protocol presents spring constant analysis for molecular bonds detected by BFP.

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure ...