This work was supported by funds from NIGMS R35GM133488 and from the Roy J. Carver Charitable Trust to V.V.

1. Preparation of the NMR sample
<ol>
	<li>Express and purify a 2H,15N-labled sample of the protein of interest. 
	NOTE: While a 15N-labeled protein sample can be used for acquisition of the CPMG RD experiment, perdeuteration (where possible) dramatically increases the quality of the obtained data. Protocols for the production of perdeuterated proteins are available in the literature13.</li>
	<li>Buffer exchange the purified protein sample into a degassed NMR buffer.</li>
	<li>Transfer the NMR sample into the NMR tube. 
	NOTE: The concentration of the NMR sample needs to be carefully optimized in order to maximize the signal-to-noise ratio and minimize the occurrence of protein-protein interactions. In general, the typical concentration range for CPMG experiments is 0.1-1.0 mM.</li>
</ol>
2. First time set-up of the NMR experiment
<ol>
	<li>Download and unzip the Supplemental Files.</li>
	<li>Copy the files bits.vv and trosy_15N_cpmg.vv (located the in the folder named pulseprogram) into the pulse program directory (&#60;path&#62;/exp/stan/nmr/lists/pp/user). 
	NOTE: Make sure the path to bits.vv listed on the first line of the trosy_15N_cpmg.vv file is correct.</li>
	<li>Open acquisition software and copy a previously run 1H-15N HSQC experiment into a new experiment using the command edc.</li>
	<li>Using the command pulprog, load the pulse program trosy_15N_cpmg.vv into the newly created experiment.</li>
	<li>Set up the CPMG experiment using the instructions provided at the end of the pulse program file (trosy_15N_cpmg.vv).</li>
</ol>
3. Routine set-up of the NMR experiment
<ol>
	<li>Introduce the sample in the magnet and perform all the basic steps for acquisition of any NMR experiment: lock and shim the sample; match and tune the 1H and 15N channels.</li>
	<li>Set p1 and p7 to the duration of the 1H and 15N hard 90&#176; pulses, respectively. 
	NOTE: For optimal results, it is important to calibrate the 15N 90&#176; pulse with great care. Calibration is usually accomplished using a 100 mM sample of 15N-labeled urea in DMSO as described in the spectrometer manual. In addition, it is possible to double-check the calibration directly on the working NMR sample by acquiring the first increment of a 1H-15N HSQC spectrum in which the 15N 90&#176; pulse of the first INEPT block is switched to a 180&#176; pulse. If the calibration is correct, a null should be obtained.</li>
	<li>In the acquisition window, set the center and spectral width for the 1H and 15N dimensions. 
	NOTE: Center the 1H spectrum at the frequency of the water signal for optimal water suppression.</li>
	<li>Set the relaxation delay (d30) equal to 0.7 T2, where T2&#160;is the expected 15N transverse relaxation time of your protein. 
	NOTE: The value of d30 can be optimized empirically to obtain the best results.</li>
	<li>Use the command vclist to create a list of integer numbers corresponding to the parameter n in Figure 1A and Figure 1B. Ensure that each entry in the list corresponds to a different CPMG field (&#957;cpmg) according to &#957;cpmg = 4 x n / d30. Make sure that the first number in the list is 0 (this corresponds to the reference experiment for which the CPMG block is skipped and d30 = 0 s) and not to use numbers larger than 1,000 x d30 / 4. Numbers larger than this threshold result in &#957;cpmg &#62; 1 kHz and might damage the probe.</li>
	<li>Set l8 to the number of entries in the vclist, l3 to the number of real points for the indirect dimension (usually a range of 64-200 is set for l3), and 1 td equal to l8 x l3 x 2. 
	NOTE: Perform the steps 3.7-3.11 to optimize the water suppression.</li>
	<li>Set the receiver gain (rg) to 1; open the pulse program file (edcpul), go to line 91, remove the semicolon preceding goto 999, and save the file.</li>
	<li>Using the command gs adjust the parameters spdb0 (or spdw0) in order to minimize the intensity of the FID signal.</li>
	<li>Reintroduce the semicolon at line 91 of the pulse program file and save the file.</li>
	<li>Repeat the steps 3.7-3.9 to optimize spdb11 (line 168 of the pulse program file), spdb2 (line 179 of the pulse program file), and pldb2 (line 184 of the pulse program file).</li>
	<li>Run the command rga to optimize the receiver gain.</li>
	<li>Set the number of scans (ns) to a multiple of 8.</li>
	<li>Run the command zg to start the experiment.</li>
</ol>
4. Processing and analysis of the NMR data
<ol>
	<li>Copy the folder named Process (Supplemental Files) into the directory containing the ser file.</li>
	<li>Use the files sep_fid.com and ft2D.com to process the NMR data. 
	NOTE: Instruction as to how to edit the processing scripts is provided within the sep_fid.com and ft2D.com files.</li>
	<li>Open the ucsf files contained in the directory CPMG_Sparky_files in Sparky. 
	NOTE: The ucsf files are created by the processing scripts. There is one ucsf file for each entry in the vclist. The first ucsf file (test_1.ucsf) contains the reference experiment. The subsequent ucsf files (test_2.ucsf, test_3.ucsf,&#8230; test_n.ucsf) are ordered from the lowest to the largest value of &#957;cpmg.</li>
	<li>Pick the NMR cross-peaks on the reference NMR spectrum (test_1.ucsf).</li>
	<li>Select all the picked cross-peaks using the Sparky command pa.</li>
	<li>Run the Sparky command rh. This command opens a dialogue window. Select the option heights at the same position in each spectrum. Click on Setup and check all the NMR spectra. Click on Update and save the output file in the working directory. The output file contains the matrix of the signal intensities over all opened NMR spectra. 
	NOTE: More accurate protocols that use lineshape fitting for recovering intensities from interleaved pseudo-3D experiments have been described in the literature14. Freely available software for lineshape fitting is available at https://pint-nmr.github.io/PINT/ (PINT), https://www.ucl.ac.uk/hansen-lab/fuda/&#160;(FUDA), and within NMRPipe (nLinLS module) and SPARKY (it module).</li>
	<li>For each CPMG field, convert the signal intensities to R2 rates using the formula <img alt="Equation 3" src="/files/ftp_upload/62395/62395eq03.jpg" />, where I0 and Id30 are the intensity of the reference (vc = 0) and relaxed (vc &#62; 0) NMR spectra, respectively. 
	NOTE: In the Supplemental Files, a template of the spreadsheet file (R2_calc.xls) used to convert the signal intensities to R2 rates and to visualize the RD profiles is provided.</li>
	<li>Read the noise level in the reference spectrum using the Sparky command st and propagate the error on the R2 rates. 
	NOTE: In the Supplemental Files, a template of the spreadsheet file (R2_calc.xls) used to propagate the error on the measured R2 rates is provided.</li>
</ol>
5. Fitting RD curves
<ol>
	<li>Copy the folder named RD_fitting (Supplemental Files) into the working directory.</li>
	<li>For each assigned NMR peak, estimate Rex by calculating the <img alt="Equation 4" src="/files/ftp_upload/62395/62395eq04.jpg" />. <img alt="Equation 5" src="/files/ftp_upload/62395/62395eq05.jpg" /> and <img alt="Equation 6" src="/files/ftp_upload/62395/62395eq06.jpg" /> are the R2 rates measured at the lowest and highest &#957;cpmg in the vclist.</li>
	<li>Visually inspect the RD curves with Rex larger than two times the estimated error and discard all the RD curves that are too noisy to be modeled accurately. 
	NOTE: Noise on Rex can be propagated from the errors on <img alt="Equation 5" src="/files/ftp_upload/62395/62395eq05.jpg" /> and <img alt="Equation 6" src="/files/ftp_upload/62395/62395eq06.jpg" />.</li>
	<li>Prepare an input file for the fitting script using all RD curves with Rex larger than two times the estimated error. 
	NOTE: Detailed instruction on the preparation of the input files are provided within the fitting scripts. Example input files are provided as Supplemental Files. Commonly, RD data are measured at two different static fields and fitted simultaneously. In this protocol, two different input files are required (Supplemental Files).</li>
	<li>Fit the RD data using the scripts provided in Supplemental Files. 
	NOTE: Two different scripts are provided to perform a residue-based or a global fit of the RD curves, respectively. Both scripts fit the RD curves using a two-site exchange model and the Carver-Richards equation. More detailed instructions are provided within the fitting scripts. Additional software packages such as Chemex (https://github.com/gbouvignies/ChemEx)&#160;and CATIA (https://www.ucl.ac.uk/hansen-lab/catia/&#160;) are available to carry out data fitting using the Block-McConnell equations.</li>
	<li>Test the reliability of the fitted parameters estimating the reduced &#967;2 as a function of pb and kex. 
	NOTE: The reduced &#967;2 is provided in the output file. pb and kex can be restrained to specific values using lower and upper bounds in the fitting procedures. In our scripts, the lower and upper bounds for pb are lb(2) and ub(2), respectively. The lower and upper bounds for kex are lb(3) and ub(3), respectively.</li>
	<li>Estimate the error on the fitted parameters. This can be done by setting the value of MC_fac in the script to 1 and repeating the fitting multiple times (typically &#62;20 repeats). The error on each parameter is estimated as the standard deviation of the distribution. 
	NOTE: Setting MC_fac to 1 generates a synthetic dataset in which a Gaussian distributed error (calculated based on the experimental error) is added to the experimental data.</li>
</ol>

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All authors have read and approved the manuscript. We declare no conflicts of interest.

This manuscript describes the protocol implemented in the laboratory for acquisition and analysis of 15N RD data on proteins. In particular, the crucial steps for preparation of the NMR sample, measurement of the NMR data, and analysis of the RD profiles are covered. Below some important aspects regarding the acquisition and analysis of RD experiments are discussed. However, for a more in-depth description of the experiment and data analysis, careful studying of the original literature is highly recommended<su...

Carr-Purcell Meiboom-Gill (CPMG) relaxation dispersion (RD) experiments are used on a routine base to characterize conformational equilibria occurring on the &#181;s-ms timescale by solution NMR spectroscopy1,2,3,4,5. Compared to other methods for investigation of conformational dynamics, CPMG techniques are relatively easy to implement on modern NMR spectrometers, do not require specialized sample preparation steps (i.e., crystallization, sample freezing or alignment, and/or covalent conjugation with a fluorescent or paramagnetic tag), and provide a comprehensive characterization of conformational equilibria returning structural, kinetic, and thermodynamic information on exchange processes. In order for a CPMG experiment to report on a conformational equilibrium, two conditions must apply: (i) the observed NMR spins must possess different chemical shifts in the states undergoing conformational exchange (microstates) and (ii) the exchange has to occur at a time scale ranging from ~50 &#181;s to ~10 ms. Under these conditions, the observed transverse relaxation rate (<img alt="Equation 1" src="/files/ftp_upload/62395/62395eq01v2.jpg" />) is the sum of the intrinsic R2 (the R2 measured in the absence of &#181;s-ms dynamics, <img alt="Equation 2" src="/files/ftp_upload/62395/62395eq02v3.jpg" />) and the exchange contribution to the transverse relaxation (Rex). The Rex contribution to R2obs can be progressively quenched by reducing the spacing between the 180&#176; pulses constituting the CPMG block of the pulse sequence, and the resulting RD curves can be modeled using the Bloch-McConnell theory to obtain the chemical shift difference among microstates, the fractional population of each microstate, and the rates of exchange among microstates (Figure 1)1,2,3.
Several different pulse sequences and analysis protocols have been reported in the literature for 15N CPMG experiments. Herein, the protocol implemented in the laboratory is described. In particular, the crucial steps for preparation of the NMR sample, set up and acquisition of the NMR experiments, and processing and analysis of the NMR data will be introduced (Figure 2). To facilitate transfer of the protocol to other laboratories, the pulse program, processing and analysis scripts, and one example dataset are provided as Supplemental Files and are available for download at (https://group.chem.iastate.edu/Venditti/downloads.html). The provided pulse sequence incorporates a four-step phase cycle in the CPMG block for suppression of offset-dependent artifacts6 and it is coded for acquisition of several interleaved experiments. These interleaved experiments have an identical relaxation period but different numbers of refocusing pulses in order to achieve different CPMG fields7. It is also important to notice that the described pulse program measures the 15N R2 of the TROSY component of the NMR signal8. Overall, the protocol has been successfully applied for the characterization of conformational exchange in medium and large-sized proteins4,5,9,10. For smaller systems (&#60;20 kDa), the use of an Heteronuclear Single Quantum Coherence (HSQC)-based pulse sequence11,12 is advisable.

<table><tbody><tr><td>Cryoprobe</td><td>Bruker</td><td>5mm TCI 800 H-C/N-D cryoprobe</td><td>Improve sensitivity</td></tr><tr><td>Deuterium Oxide</td><td>Sigma Aldrich</td><td>756822-1</td><td>&gt;99.8% pure, utilised in preparing NMR samples and deuterated cultures</td></tr><tr><td>Hand driven centrifuge</td><td>United Scientific supply</td><td>CENTFG1</td><td>Used to remove any air bubbles or residual liquid stuck on the walls of NMR tube.</td></tr><tr><td>High Field NMR spectrometer</td><td>Bruker</td><td>Bruker Avance II 600, Bruker Avance 800</td><td>Acquisition of the NMR data</td></tr><tr><td>MATLAB</td><td>MathWorks</td><td>https://www.mathworks.com/products/get-matlab.html</td><td>Modeling of the NMR data</td></tr><tr><td>NMR pasteur Pipette</td><td>Corning Incorporation</td><td>7095D-NMR</td><td>Pyrex glass pastuer pipette to transfer liquid sample in NMR tube</td></tr><tr><td>NMR tube</td><td>Willmad Precision</td><td>535-PP-7</td><td>5mm thin wall 7&#039;&#039; cylinderical glass tube</td></tr><tr><td>NMRPipe</td><td>Institute of Biosciences and Biotechnology research</td><td>https://www.ibbr.umd.edu/nmrpipe/install.html</td><td>NMR data processing</td></tr><tr><td>SPARKY</td><td>University of California, San Francisco</td><td>https://www.cgl.ucsf.edu/home/sparky/</td><td>Analysis of the NMR data</td></tr><tr><td>Tospin 3.2 (or newer)</td><td>Bruker</td><td>https://www.bruker.com/protected/en/services/software-downloads/nmr/pc/pc-topspin.html</td><td>Acquisition Software</td></tr></tbody></table>

15n cpmg relaxation dispersion for the investigation of protein conformational dynamics on the &#181;s-ms timescale

Protein conformational dynamics play fundamental roles in regulation of enzymatic catalysis, ligand binding, allostery, and signaling, which are important biological processes. Understanding how the balance between structure and dynamics governs biological function is a new frontier in modern structural biology and has ignited several technical and methodological developments. Among these, CPMG relaxation dispersion solution NMR methods provide unique, atomic-resolution information on the structure, kinetics, and thermodynamics of protein conformational equilibria occurring on the &#181;s-ms timescale. Here, the study presents detailed protocols for acquisition and analysis of a&#160;15N relaxation dispersion experiment. As an example, the pipeline for the analysis of the &#181;s-ms dynamics in the C-terminal domain of bacteria Enzyme I is shown.

The protocol described here results in acquisition of RD profiles for each peak in the 1H-15N TROSY spectrum (Figure 3A). From the acquired RD profiles, it is possible to estimate the exchange contribution to the 15N transverse relaxation of each backbone amide group (Figure 3A,3B). By plotting the Rex on the 3D structure of the protein under investigation, it is possible to identify the structural reg...

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15N CPMG Relaxation Dispersion for the Investigation of Protein Conformational Dynamics on the  &#181;s-ms Timescale

Here, a detailed description of the protocol implemented in the laboratory for acquisition and analysis of 15N relaxation dispersion profiles by solution NMR spectroscopy is provided.

15N CPMG Relaxation Dispersion for the Investigation of Protein Conformational Dynamics on the &#181;s-ms Timescale

Protein conformational dynamics play fundamental roles in regulation of enzymatic catalysis, ligand binding, allostery, and signaling, which are important ...

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Biochemistry

5.1K Views.  Iowa State University. Here, a detailed description of the protocol implemented in the laboratory for acquisition and analysis of 15N relaxation dispersion profiles by solution NMR spectroscopy is provided.

Video: 15N CPMG Relaxation Dispersion for the Investigation of Protein Conformational Dynamics on the  µs-ms Timescale

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 ...

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the propagation of neurodegenerative disorders. Such assemblies consist in multiple protein subunits organized in a non-covalent way to form large macromolecular objects that can execute a variety of cellular functions or cause detrimental consequences. Atomic insights into the assembly mechanisms and the functioning of those macromolecular assemblies remain often scarce since their inherent insolubility and non-crystallinity often drastically reduces the quality of the data obtained from most techniques used in structural biology, such as X-ray crystallography and solution Nuclear Magnetic Resonance (NMR). We here present magic-angle spinning solid-state NMR spectroscopy (SSNMR) as a powerful method to investigate structures of macromolecular assemblies at atomic resolution. SSNMR can reveal atomic details on the assembled complex without size and solubility limitations. The protocol presented here describes the essential steps from the production of 13C/15N isotope-labeled macromolecular protein assemblies to the acquisition of standard SSNMR spectra and their analysis and interpretation. As an example, we show the pipeline of a SSNMR structural analysis of a filamentous protein assembly.

Structures of supramolecular protein assemblies at atomic resolution are of high relevance because of their crucial roles in a variety of biological phenomena. Herein, we present a protocol to perform high-resolution structural studies on insoluble and non-crystalline macromolecular protein assemblies by magic-angle spinning solid-state nuclear magnetic resonance spectroscopy (MAS SSNMR).

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the ...

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages for identifying biologically active chemical structures that can modify complex phenotypes such as lifespan. Described here is a simple protocol for producing hundreds of 96-well culture plates with fairly consistent numbers of C. elegans in each well. Next, we specified how to use these cultures to screen thousands of chemicals for effects on the lifespan of the nematode C. elegans. This protocol makes use of temperature sensitive sterile strains, agar plate conditions, and simple animal handling to facilitate the rapid and high throughput production of synchronized animal cultures for screening.

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Here we describe a simple protocol for rapidly producing hundreds of nematode growth media agar, 96-well culture plates with consistent numbers of Caenorhhabditis elegans per well. These cultures are useful for the phenotypic screening of whole organisms. We focus here on using these cultures to screen chemicals for pro-longevity effects.

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages ...

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 ...

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.

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the ...

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

Most human body proteins' activity and functionality are related to configurational changes of entire subdomains within the protein crystal structure. The crystal structures build the basis for any calculation that describes the structure or dynamics of a protein, most of the time with strong geometrical restrictions. However, these restrictions from the crystal structure are not present in the solution. The structure of the proteins in the solution may differ from the crystal due to rearrangements of loops or subdomains on the pico to nanosecond time scale (i.e., the internal protein dynamics time regime). The present work describes how slow motions on timescales of several tens of nanoseconds can be accessed using neutron scattering. In particular, the dynamical characterization of two major human proteins, an intrinsically disordered protein that lacks a well-defined secondary structure and a classical antibody protein, is addressed by neutron spin echo spectroscopy (NSE) combined with a wide range of laboratory characterization methods. Further insights into protein domain dynamics were achieved using mathematical modeling to describe the experimental neutron data and determine the crossover between combined diffusive and internal protein motions. The extraction of the internal dynamic contribution to the intermediate scattering function obtained from NSE, including the timescale of the various movements, allows further vision into the mechanical properties of single proteins and the softness of proteins in their nearly natural environment in the crowded protein solution.

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

The present protocol describes methods for investigating the structure and dynamics of two model proteins that have an important role in human health. The technique combines bench-top biophysical characterization with neutron spin echo spectroscopy to access the dynamics at time and length scales relevant for protein interdomain motions.

Most human body proteins' activity and functionality are related to configurational changes of entire subdomains within the protein crystal structure. The ...

Engineering

Protocol for NMR 15N Relaxation and hetNOE Experiments in Protein Dynamics Study

NMR 15N Relaxation Experiments for the Investigation of Picosecond to Nanoseconds Structural Dynamics of Proteins

Nuclear magnetic resonance (NMR) spectroscopy allows studying proteins in solution and under physiological temperatures. Frequently, either the amide groups of the protein backbone or the methyl groups in side chains are used as reporters of structural dynamics in proteins. A structural dynamics study of the protein backbone of globular proteins on 15N labeled and fully protonated samples usually works well for proteins with a molecular weight of up to 50 kDa. When side chain deuteration in combination with transverse relaxation optimized spectroscopy (TROSY) is applied, this limit can be extended up to 200 kDa for globular proteins and up to 1 MDa when the focus is on the side chains. When intrinsically disordered proteins (IDPs) or proteins with intrinsically disordered regions (IDRs) are investigated, these weight limitations do not apply but can go well beyond. The reason is that IDPs or IDRs, characterized by high internal flexibility, are frequently dynamically decoupled. Various NMR methods offer atomic-resolution insights into structural protein dynamics across a wide range of time scales, from picoseconds up to hours. Standard 15N relaxation measurements overview a protein&#39;s internal flexibility and characterize the protein backbone dynamics experienced on the fast pico- to nanosecond timescale. This article presents a hands-on protocol for setting up and recording NMR 15N R1, R2, and heteronuclear Overhauser effect (hetNOE) experiments. We show exemplary data and explain how to interpret them simply qualitatively before any more sophisticated analysis.

Author Spotlight: Exploring Intrinsically Disordered Protein Dynamics Through NMR Relaxation Experiments

Nuclear magnetic resonance (NMR) spectroscopy can characterize structural protein dynamics in a residue-specific manner. We provide a hands-on protocol for recording NMR 15N R1 and R2 relaxation and {1H}-15N heteronuclear Overhauser effect (hetNOE) experiments, sensitive to the picoseconds to nanoseconds timescale.

Nuclear magnetic resonance (NMR) spectroscopy allows studying proteins in solution and under physiological temperatures. Frequently, either the amide ...

Structure-Based Simulation and Sampling of Transcription Factor Protein Movements along DNA from Atomic-Scale Stepping to Coarse-Grained Diffusion

One-dimensional (1-D) sliding of transcription factor (TF) protein along DNA is essential for facilitated diffusion of the TF to locate target DNA site for genetic regulation. Detecting base-pair (bp) resolution of the TF sliding or stepping on the DNA is still experimentally challenging. We have recently performed all-atom molecular dynamics (MD) simulations capturing spontaneous 1-bp stepping of a small WRKY domain TF protein along DNA. Based on the 10 &#181;s WRKY stepping path obtained from such simulations, the protocol here shows how to conduct more extensive conformational samplings of the TF-DNA systems, by constructing the Markov state model (MSM) for the 1-bp protein stepping, with various numbers of micro- and macro-states tested for the MSM construction. In order to examine processive 1-D diffusional search of the TF protein along DNA with structural basis, the protocol further shows how to conduct coarse-grained (CG) MD simulations to sample long-time scale dynamics of the system. Such CG modeling and simulations are particularly useful to reveal the protein-DNA electrostatic impacts on the processive diffusional motions of the TF protein above tens of microseconds, in comparison with sub-microseconds to microseconds protein stepping motions revealed from the all-atom simulations.

The goal of this protocol is to reveal structural dynamics of one-dimensional diffusion of protein along DNA, using a plant transcription factor WRKY domain protein as an exemplary system. To do this, both atomistic and coarse-grained molecular dynamics simulations along with extensive computational samplings have been implemented.

One-dimensional (1-D) sliding of transcription factor (TF) protein along DNA is essential for facilitated diffusion of the TF to locate target DNA site ...

¹⁵N CPMG Relaxation Dispersion for the Investigation of Protein Conformational Dynamics on the µs-ms Timescale

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

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

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

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

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

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

NMR ¹⁵N Relaxation Experiments for the Investigation of Picosecond to Nanoseconds Structural Dynamics of Proteins

Structure-Based Simulation and Sampling of Transcription Factor Protein Movements along DNA from Atomic-Scale Stepping to Coarse-Grained Diffusion