Name: visualizing the conformational dynamics of membrane receptors using single-molecule fret
Uploaded: 2022-08-17T14:15:00
Description: Watch this Scientific Journal Video about Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET at JoVE.com

We thank members of the Reza Vafabakhsh lab for discussions. This work was supported by the National Institutes of Health grant R01GM140272 (to R.V.), by The Searle Leadership Fund for the Life Sciences at Northwestern University, and by the Chicago Biomedical Consortium with support from the Searle Funds at The Chicago Community Trust (to R.V.). B.W.L. was supported by the National Institute of General Medical Sciences (NIGMS) Training Grant T32GM-008061.

The overall workflow of the protocol is described in Figure 1.
1. Preparation of the sample chamber
<ol>
	<li>Slide and coverslip cleaning 
	NOTE: These steps aim to clean the surfaces of the slides as well as the coverslips and prepare them for aminosilanization. One critical requirement for conducting single-molecule fluorescence experiments on surface-tethered molecules is a passivated surface. The most reliable and reproducible passivation technique...

<ol>
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GPCRs are proteins that operate on the cell membrane to initiate signal transduction. Many GPCRs consist of multiple domains, with signaling being dependent on the cooperative interaction between the domains. To modulate the properties of these membrane receptors, it is essential to understand the dynamic behavior of the multiple domains. Single-molecule fluorescence resonance energy transfer (smFRET) is a fluorescence technique that enables the measurement of protein conformation and dynamics in real time<sup class="xre...

The transfer of information across the plasma membrane is heavily dependent on the function of membrane receptors1. Ligand binding to a receptor leads to a conformational change and receptor activation. This process is often allosteric in nature2. With over 800 members, G protein-coupled receptors (GPCRs) are the largest family of membrane receptors in humans3. Due to their role in nearly all cellular processes, GPCRs have become important targets for therapeutic development. In the canonical model of GPCR signaling, agonist activation results in conformational changes of the receptor that subsequently activate the heterotrimeric G protein complex via exchange of GDP for GTP at the nucleotide binding pocket of G&#945;. The activated G&#945;-GTP and G&#946;&#947; subunits then control the activity of downstream effector proteins and propagate the signaling cascade4,5. This signaling process essentially depends on the ability of ligands to change the three-dimensional shape of the receptor. A mechanistic understanding of how ligands achieve this is critical for developing new therapeutics and designing synthetic receptors and sensors.
Metabotropic glutamate receptors (mGluRs) are members of the class C GPCR family and are important for the slow neuromodulatory effects of glutamate and tuning neuronal excitability6,7. Among all GPCRs, class C GPCRs are structurally unique in that they function as obligate dimers. mGluRs contain three structural domains: the Venus flytrap (VFT) domain, cysteine-rich domain (CRD), and transmembrane domain (TMD)8. The conformational changes during the activation process are complex and involve local and global conformational coupling that propagate over a 12 nm distance, as well as dimer cooperativity. The intermediate conformations, temporal ordering of states, and rate of transition between states are unknown. By following the conformation of individual receptors in real time, it is possible to identify the transient intermediate states and the sequence of conformational changes during activation. This can be achieved by applying single-molecule fluorescence resonance energy transfer9,10 (smFRET), as was recently applied to visualize the propagation of conformational changes during the activation of mGluR211. A key step in FRET experiments is the generation of FRET sensors by site-specific insertion of the donor and acceptor fluorophores into the protein of interest. An unnatural amino acid (UAA) incorporation strategy was adopted12,13,14,15 to overcome the limitations of typical site-specific fluorescent labeling technologies that require the creation of cysteine-less mutants or the insertion of a large genetically encoded tag. This allowed the conformational rearrangement of the essential compact allosteric linker, which joined the ligand-binding and signaling domains of mGluR2, to be observed. In this protocol, a step-by-step guide to performing smFRET experiments on mGluR2 is presented, including the approach for site-specific labeling of mGluR2 with UAA to attach fluorophores using the copper-catalyzed azide cyclization reaction. Moreover, this protocol describes the methodology for the direct capture of membrane proteins and data analysis. The protocol outlined here is also applicable to studying the conformational dynamics of other membrane proteins.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>(+)-Sodium L-Ascorbate</td>
			<td>Sigma Aldrich</td>
			<td>Cat # 11140-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>4-azido-L-phenylalanine</td>
			<td>Chem-Impex International</td>
			<td>Cat # 06162</td>
			<td></td>
		</tr>
		<tr>
			<td>548UAA</td>
			<td>Liauw et al. 2021</td>
			<td></td>
			<td>Transfected construct</td>
		</tr>
		<tr>
			<td>Acetic Acid</td>
			<td>Fisher Chemical</td>
			<td>64-19-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Fisher Chemical</td>
			<td>67-64-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Adobe Illustrator (2022)</td>
			<td>https://www.adobe.com/</td>
			<td>RRID:SCR_010279</td>
			<td>Software, algorithm</td>
		</tr>
		<tr>
			<td>Aminoguanidine (hydrochloride)</td>
			<td>Cayman Chemical</td>
			<td>81530</td>
			<td></td>
		</tr>
		<tr>
			<td>Aminosilane</td>
			<td>Aldrich</td>
			<td>919-30-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Bath Sonicator 2.8 L</td>
			<td>Fisher Scientific</td>
			<td></td>
			<td>Ultrasonic Bath 2.8 L</td>
		</tr>
		<tr>
			<td>Biotin-PEG</td>
			<td>Laysan Bio Inc</td>
			<td>Item# Biotin-PEG-SVA-5000-100mg</td>
			<td></td>
		</tr>
		<tr>
			<td>BTTES</td>
			<td>Click Chemistry Tools</td>
			<td>1237-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper (II) sulfate</td>
			<td>Sigma Aldrich</td>
			<td>Cat # 451657-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover slip</td>
			<td>VWR</td>
			<td>16004-306</td>
			<td>Sample chamber</td>
		</tr>
		<tr>
			<td>Cy3 Alkyne</td>
			<td>Click Chemistry Tools</td>
			<td>TA117-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Cy5 Alkyne</td>
			<td>Click Chemistry Tools</td>
			<td>TA116-5</td>
			<td></td>
		</tr>
		<tr>
			<td>DDM</td>
			<td>Anatrace</td>
			<td>Part# D310 1 GM</td>
			<td>Detergent</td>
		</tr>
		<tr>
			<td>DDM-CHS (10:1)</td>
			<td>Anatrace</td>
			<td>Part# D310-CH210 1 ML</td>
			<td>Detergent with cholecterol</td>
		</tr>
		<tr>
			<td>Defined Fetal Bovine Serum</td>
			<td>Thermo Fisher Scientific</td>
			<td>SH30070.03</td>
			<td></td>
		</tr>
		<tr>
			<td>Di01-R405/488/561/635</td>
			<td>Semrock</td>
			<td></td>
			<td>Notch filter</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Corning</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>EMCCD</td>
			<td>Andor</td>
			<td>DU-897U</td>
			<td>Camera</td>
		</tr>
		<tr>
			<td>ET542lp</td>
			<td>Chroma</td>
			<td></td>
			<td>Long pass emission filter</td>
		</tr>
		<tr>
			<td>FF640-FDi01</td>
			<td>Semrock</td>
			<td></td>
			<td>Emission dichroic filter</td>
		</tr>
		<tr>
			<td>FLAG-tag antibody</td>
			<td>Genscript</td>
			<td>A01429</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent bead</td>
			<td>Invitrogen T7279</td>
			<td>TetraSpeck microspheres</td>
			<td>Spherical bead</td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>Fisherfinest</td>
			<td>12-544-4</td>
			<td>sample chamber</td>
		</tr>
		<tr>
			<td>Glutamate</td>
			<td>Sigma Aldrich</td>
			<td>Cat # 6106-04-3</td>
			<td></td>
		</tr>
		<tr>
			<td>HEK 293T</td>
			<td>Sigma Aldrich</td>
			<td>Cat # 12022001</td>
			<td>Cell line</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>FisherBioReagents</td>
			<td>7365-45-9</td>
			<td></td>
		</tr>
		<tr>
			<td>Image splitter</td>
			<td>OptoSplit II</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>Fluka</td>
			<td>1310-58-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser</td>
			<td>Oxxius</td>
			<td></td>
			<td>4-line laser combiner</td>
		</tr>
		<tr>
			<td>Lipofectamine 3000 Transfection Reagent</td>
			<td>Thermo Fisher Scientific</td>
			<td>L3000015</td>
			<td>Transfection Reagent</td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Fisher Chemical</td>
			<td>67-56-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td>Olympus IX83</td>
			<td></td>
		</tr>
		<tr>
			<td>Milli-Q water</td>
			<td>Barnstead</td>
			<td></td>
			<td>Water Deionizer</td>
		</tr>
		<tr>
			<td>m-PEG</td>
			<td>Laysan Bio Inc</td>
			<td>Item# MPEG-SIL-5000-1g</td>
			<td></td>
		</tr>
		<tr>
			<td>NF03-405/488/532/635</td>
			<td>Semrock</td>
			<td></td>
			<td>Dichroic mirror</td>
		</tr>
		<tr>
			<td>OptiMEM</td>
			<td>Thermo Fisher Scientific</td>
			<td>51985091</td>
			<td>Reduced Serum Medium</td>
		</tr>
		<tr>
			<td>OptiMEM/Reduced serum medium</td>
			<td>Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>OriginPro (2020b)</td>
			<td>https://www.originlab.com/</td>
			<td>RRID:SCR_014212</td>
			<td>Data analysis and graphing software</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>pIRE4-Azi</td>
			<td>Addgene</td>
			<td>Plasmid # 105829</td>
			<td>Transfected construct</td>
		</tr>
		<tr>
			<td>Poly-L-lysine hydrobromide</td>
			<td>Sigma Aldrich</td>
			<td>Cat # P2636</td>
			<td></td>
		</tr>
		<tr>
			<td>Protocatechuic acid (PCA)</td>
			<td>HWI group</td>
			<td>99-50-3</td>
			<td></td>
		</tr>
		<tr>
			<td>smCamera (Version 1.0)</td>
			<td>http://ha.med.jhmi.edu/resources/</td>
			<td></td>
			<td>Camera software</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>FisherBioReagents</td>
			<td>144-55-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sigma</td>
			<td>1310-73-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter</td>
			<td>Whatman UNIFLO</td>
			<td>Cat#9914-2502</td>
			<td>Liquid filtration</td>
		</tr>
		<tr>
			<td>Trolox</td>
			<td>Sigma</td>
			<td>53188-07</td>
			<td></td>
		</tr>
	</tbody>
</table>

visualizing the conformational dynamics of membrane receptors using single-molecule fret

The ability of cells to respond to external signals is essential for cellular development, growth, and survival. To respond to a signal from the environment, a cell must be able to recognize and process it. This task mainly relies on the function of membrane receptors, whose role is to convert signals into the biochemical language of the cell. G protein-coupled receptors (GPCRs) constitute the largest family of membrane receptor proteins in humans. Among GPCRs, metabotropic glutamate receptors (mGluRs) are a unique subclass that function as obligate dimers and possess a large extracellular domain that contains the ligand-binding site. Recent advances in structural studies of mGluRs have improved the understanding of their activation process. However, the propagation of large-scale conformational changes through mGluRs during activation and modulation is poorly understood. Single-molecule fluorescence resonance energy transfer (smFRET) is a powerful technique to visualize and quantify the structural dynamics of biomolecules at the single-protein level. To visualize the dynamic process of mGluR2 activation, fluorescent conformational sensors based on unnatural amino acid (UAA) incorporation were developed that allowed site-specific protein labeling without perturbation of the native structure of receptors. The protocol described here explains how to perform these experiments, including the novel UAA labeling approach, sample preparation, and smFRET data acquisition and analysis. These strategies are generalizable and can be extended to investigate the conformational dynamics of a variety of membrane proteins.

Expression and fluorescent labeling of UAA-based FRET sensor 
Herein, exemplary results of the insertion and fluorescent labeling of a UAA (AZP) within the CRD of mGluR2 (548UAA) are discussed11. As mentioned previously, to insert AZP into mGluR2, co-expression of the engineered translational machinery, which includes a modified tRNA synthetase and complementary tRNA (pIRE4-Azi), and mGluR2 containing an amber codon at position 548, created using mutagenesis, is necessary (<s...

Watch this Scientific Journal Video about Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET at JoVE.com

Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET

This study presents a detailed procedure to perform single-molecule fluorescence resonance energy transfer (smFRET) experiments on G protein-coupled receptors (GPCRs) using site-specific labeling via unnatural amino acid (UAA) incorporation. The protocol provides a step-by-step guide for smFRET sample preparation, experiments, and data analysis.

The ability of cells to respond to external signals is essential for cellular development, growth, and survival. To respond to a signal from the ...

Here, we used single-molecule FRET, and site-specific protein labeling via a natural amino acid incorporation to study the conformational dynamics of mGluR2. This allows to study the protein dynamics without perturbing protein structure. Atomic structure provide a static image of a protein.However, single-molecule FRET enables us to visualize the full range of protein motion in real-time and solution. This method can be applied to study the dynamics of any protein. Moreover, a natural amino acid incorporation can be used to study protein-protein interactions, and the engineering of synthetic receptors.Sample loading and optimization of imaging requires patience and practice. To begin, mark holes to be drilled on the glass slide with a marker. Use a Dremel to drill small holes on the slide while the slide is dipped in water.Sonicate the slides and cover slips in acetone for 30 minutes in a bath sonicator at 23 degrees Celsius. Dispose of acetone from the slide and cover slip jars. Rinse thoroughly with water, and sonicate in five-molar potassium hydroxide for 30 minutes.Next, rinse the slides and cover slip with water, and sonicate in methanol for two minutes. Leave the jars filled with methanol until the amino salinization step. Pour freshly prepared amino salinization mixture into the slide and cover slip jars.Incubate, sonicate, and again incubate the slides and cover slips in an amino salinization mixture before discarding the solution. Next, wash the jars with methanol before filling them with water. Then rinse the slides with water.Dry them with air blow, and place them in the assembly boxes. Apply 60 milliliters of PEGylation solution to the slide, and place a cover slip on top of it. Mark the non-PEGylated side before storage.After incubation, gently disassemble, and rinse the slides and cover slips with water. Then dry them by blowing air, and place them in a sterile 50-milliliter tube with the PEGylated surface facing away from each other. After passaging HEK 293T cells with 0.05%trypsin-EDTA, seed the cells on poly-L/D-lysine cover slips.Place the standard media with AZP supplemented media for growing cells and metabotropic glutamate receptor 2, or mGluR2 expression. Then transfect the cells with a transfection reagent, and incubate for 24 hours before changing the medium with fresh AZP supplemented media. 20 minutes before labeling with alkyne cyanine dyes, wash the cover slips twice with warm Recording Buffer, or RB, and place them in a warm standard media with no AZP.Then remove the standard media and wash the cells with RB before adding the freshly prepared labeling solution. Incubate for 15 minutes at 37 degrees Celsius in the dark. After incubation, remove the labeling solution.Wash the cover slip containing transfected cells twice with the RB, and re-suspend in the same medium. Next, pellet the cells by spinning at 1, 000 g at four degrees Celsius for five minutes, and remove the supernatant before re-suspending the pellet in 80 to 130 microliters of the lysis solution. Break the pellet by pipetting.Then wrap it in foil, and place it on the rocker at four degrees Celsius for 30 minutes to one hour to lyse the cells. Pellet the insoluble fraction by centrifugation at 20, 000 g and four degree Celsius for 20 minutes. Then transfer the supernatant to a fresh cold tube, and store it on ice.Remove the slide and cover slip from the freezer, and allow them to warm at room temperature in the dark. Assemble the chamber by sandwiching the strips of double-sided tape between the slide and cover slip, ensuring the PEGylated surfaces form the interior of the flow chamber. Using a pipette tip, press the cover slip to ensure the tape is contacting the cover slip and slide.Apply epoxy to the edges of the slide. Apply 40 microliters of 500-nanomolar NeutrAvidin to each lane, and incubate inside a humidified dark box. After two minutes, wash each chamber lane with 100 microliters of T50 buffer.Then add 20 nanomolar biotinylated antibody solution, and incubate for 30 minutes before washing with 200 microliters of T50 buffer per lane. Turn on the computer and microscope. Then turn on the lasers to warm up.Next, turn on the EMCCD camera, and open the software. Mount the sample chamber on the microscope stage, and add the protein sample gradually to achieve approximately 400 molecules per field of view. Wash the chamber with 100 microliters of imaging buffer supplemented with 0.3 units of protocatechuate 3, 4-dioxygenase.Adjust the gain, acquisition rate, and laser powers to detect single-molecule fluorescence signals in both the donor and accepter channels. Then excite the donor, and acquire time traces until 80%of the donor molecules in the field of view are photobleached. At the end of the movie, turn on the 640-nanometer laser to directly excite the accepter until some of the molecules are photobleached.Change the field of view, and repeat the steps for excitation of donor and accepter to collect three movies per condition. For selecting single-molecule FRET traces of particle picking, select the traces where the total intensity of donor and accepter traces is stable over time. Then select the traces with anti-correlated changes in donor and accepter intensities.Select the donor and accepter molecules showing single-step photobleaching and traces more than five seconds long. Calculate the FRET deficiency. Finally, identify the conformational state by Hidden Markov Modeling, or HMM, by following the steps described in the manuscript.The labeling of AZP by cyanine dyes was achieved by a copper-catalyzed cycloaddition reaction, and resulted in effective plasma membrane labeling of 548UAA with the cell surface population labeled with donor and accepter fluorophores. In SDS-PAGE electrophoresis, a single band at 250 kilodalton was observed in HEK 293T cells expressing 548UAA, which coincided with the full-length dimeric mGluR2. Representative selective traces for multiple short-lived and long-lived for donor and accepter bleaching events are shown here.Conformational states were identified using HMM analysis. Transitions between discrete FRET states were extracted from the idealized fits, and plotted as a transition density heat map. The idealized FRET traces also yield information on dwell time for each identified confirmation.The single-molecule's traces with donor and accepter channel is shown here. Further, single molecule FRET reveals four conformational states of the mGluR2 cysteine-rich domain. Efficient peak preservation is essential to single-molecule pull-down, and care should be taken during this process.In addition, the oxygen-scavenging agent should be added immediately prior to imaging. This labeling method and single-molecule FRET experiments have been applied to multiple receptors, and have provided new insights into the mechanism of receptor activation and modulation of receptor function.

visualizing-conformational-dynamics-membrane-receptors-using-single

Research

JoVE Journal

Biochemistry

Single-molecule Super-resolution Imaging of Phosphatidylinositol 4,5-bisphosphate in the Plasma Membrane with Novel Fluorescent Probes

Phosphoinositides in the cell membrane are signaling lipids with multiple cellular functions. Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is a determinant phosphoinositide of the plasma membrane (PM), and it is required to modulate ion channels, actin dynamics, exocytosis, endocytosis, intracellular signaling, and many other cellular processes. However, the spatial organization of PI(4,5)P2 in the PM is controversial, and its nanoscale distribution is poorly understood due to the technical limitations of research approaches. Here by utilizing single molecule localization microscopy and the Pleckstrin Homology (PH) domain based dual color fluorescent probes, we describe a novel method to visualize the nanoscale distribution of PI(4,5)P2 in the PM in fixed membrane sheets as well as live cells.

PI(4,5)P2 regulates various cellular functions, but its nanoscale organization in the cell plasma membrane is poorly understood. By labeling PI(4,5)P2 with a dual-color fluorescent probe fused to the Pleckstrin Homology domain, we describe a novel approach to study the PI(4,5)P2 spatial distribution in the plasma membrane at the nanometer scale.

Phosphoinositides in the cell membrane are signaling lipids with multiple cellular functions. Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is a ...

Structural Information from Single-molecule FRET Experiments Using the Fast Nano-positioning System

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. Thereby, multiple smFRET measurements are used to localize an unknown dye position inside a protein complex by means of trilateration. In order to obtain quantitative information, the Nano-Positioning System (NPS) uses probabilistic data analysis to combine structural information from X-ray crystallography with single-molecule fluorescence data to calculate not only the most probable position but the complete three-dimensional probability distribution, termed posterior, which indicates the experimental uncertainty. The concept was generalized for the analysis of smFRET networks containing numerous dye molecules. The latest version of NPS, Fast-NPS, features a new algorithm using Bayesian parameter estimation based on Markov Chain Monte Carlo sampling and parallel tempering that allows for the analysis of large smFRET networks in a comparably short time. Moreover, Fast-NPS allows the calculation of the posterior by choosing one of five different models for each dye, that account for the different spatial and orientational behavior exhibited by the dye molecules due to their local environment.
Here we present a detailed protocol for obtaining smFRET data and applying the Fast-NPS. We provide detailed instructions for the acquisition of the three input parameters of Fast-NPS: the smFRET values, as well as the quantum yield and anisotropy of the dye molecules. Recently, the NPS has been used to elucidate the architecture of an archaeal open promotor complex. This data is used to demonstrate the influence of the five different dye models on the posterior distribution.

We present the setup and experimental procedure to obtain smFRET data from large donor-acceptor networks with a TIRF microscope. The step-by-step analysis of these measurements with the Bayesian inference software Fast-NPS yields high-resolved structural information via the application of adapted dye models.

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. ...

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of reactive oxygen species, and more. These fluctuations can lead to a widespread protein unfolding, aggregation, and cell death. Therefore, cells have evolved a dynamic and stress-specific network of molecular chaperones, which maintain a "healthy" proteome during stress conditions. ATP-independent chaperones constitute one major class of molecular chaperones, which serve as first-line defense molecules, protecting against protein aggregation in a stress-dependent manner. One feature these chaperones have in common is their ability to utilize structural plasticity for their stress-specific activation, recognition, and release of the misfolded client. 
In this paper, we focus on the functional and structural analysis of one such intrinsically disordered chaperone, the bacterial redox-regulated Hsp33, which protects proteins against aggregation during oxidative stress. Here, we present a toolbox of diverse techniques for studying redox-regulated chaperone activity, as well as for mapping conformational changes of the chaperone, underlying its activity. Specifically, we describe a workflow which includes the preparation of fully reduced and fully oxidized proteins, followed by an analysis of the chaperone anti-aggregation activity in vitro using light-scattering, focusing on the degree of the anti-aggregation activity and its kinetics. To overcome frequent outliers accumulated during aggregation assays, we describe the usage of Kfits, a novel graphical tool which allows easy processing of kinetic measurements. This tool can be easily applied to other types of kinetic measurements for removing outliers and fitting kinetic parameters. To correlate the function with the protein structure, we describe the setup and workflow of a structural mass spectrometry technique, hydrogen-deuterium exchange mass spectrometry, that allows the mapping of conformational changes on the chaperone and substrate during different stages of Hsp33 activity. The same methodology can be applied to other protein-protein and protein-ligand interactions.

One of the most challenging stress conditions that organisms encounter during their lifetime involves the accumulation of oxidants. During oxidative stress, cells heavily rely on molecular chaperones. Here, we present methods used to investigate the redox-regulated anti-aggregation activity, as well as to monitor structural changes governing the chaperone function using HDX-MS.

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of ...

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

Visualizing Surface T-Cell Receptor Dynamics Four-Dimensionally Using Lattice Light-Sheet Microscopy

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the structure-function relationship of these receptors in situ, we need to visualize and track them on the live cell surface with enough spatiotemporal resolution. Here we show how to use recently developed Lattice Light-Sheet Microscopy (LLSM) to image T-cell receptors (TCRs) four-dimensionally (4D, space and time) at the live cell membrane. T cells are one of the main effector cells of the adaptive immune system, and here we used T cells as an example to show that the signaling and function of these cells are driven by the dynamics and interactions of the TCRs. LLSM allows for 4D imaging with unprecedented spatiotemporal resolution. This microscopy technique therefore can be generally applied to a wide array of surface or intracellular molecules of different cells in biology.

The goal of this protocol is to show how to use Lattice Light-Sheet Microscopy to four-dimensionally visualize surface receptor dynamics in live cells. Here T cell receptors on CD4+ primary T cells are shown.