This research used resources at the Spallation Neutron Source (BL-15, BL-6, Biology and Chemistry labs), a DOE Office of the Science User Facility operated by the Oak Ridge National Laboratory. This research also used resources at the MLZ-FRM2 reactor Garching (KWS-2, Phoenix-J-NSE) and the JCNS1 at Forschungszentrum J&#252;lich GmbH, Germany. The author acknowledges Dr. Ralf Biehl and Dr. Andreas Stadler for their help with modeling and their contribution to both IgG and MBP proteins research, Dr. Piotr A. &#379;o&#322;nierczuk for NSE data reduction support, Dr. Changwoo Do for support with SANS measurements, and Rhonda Moody and Dr. Kevin Weiss for SNS biochemistry lab support.

The present work characterizes two proteins: a regular human antibody protein IgG, and the intrinsically disordered MBP. The lyophilized form of the proteins was obtained from commercial sources (see Table of Materials).
1. Protein sample preparation
<ol>
	<li>Prepare a 50 mM sodium phosphate + 0.1 M NaCl buffer by weighing and dissolving the respective solid reagents in heavy water (D2O) (see Table of Materials). This is the deuterated buffer solvent for IgG.
	<ol>
		<li>Adjust the pH of the buffer solution to 6.6.</li>
	</ol>
	</li>
	<li>Prepare a 20 mM sodium phosphate + 6 M urea buffer by weighing and dissolving the respective solid components in heavy water (D2O). This is the deuterated buffer solvent for MBP.
	<ol>
		<li>Adjust the pH of the buffer solution to 4.7.</li>
	</ol>
	</li>
	<li>Filter the buffer solvents using 0.2 &#181;m pore size filters (see Table of Materials).</li>
	<li>Weigh and dissolve the purified protein lyophilized powders in the deuterated solvents for the respective proteins (Steps 1.1.-1.2.) at high protein concentration (~50 mg/mL).</li>
	<li>Load the protein solution in dialyze baskets with dialysis membranes of 3.5 K MWCO, and dialyze against the filtered buffer for 24 h at 10 &#176;C for MBP and 25 &#176;C for IgG (see Table of Materials) by slightly shaking the tubes to create a diffusion gradient.</li>
	<li>Dilute the protein solution using the dialysis buffer in a series of concentrations: 1, 2, 5, 10, and 50 mg/mL.</li>
	<li>Determine the exact concentrations using a Nanodrop spectrophotometer (see Table of Materials).</li>
</ol>
2. Preliminary sample characterization by dynamic light scattering (DLS)
<ol>
	<li>Load 80 &#181;L of each protein solution from the concentration series prepared above (Step 1.6.) into the DLS disposable cell (see Table of Materials) and determine the diffusion coefficients, averaging over 10 acquisitions.</li>
	<li>Plot the translational diffusion coefficients as a function of protein concentration and extrapolate to zero concentration.</li>
	<li>Load each protein solution from the concentration series into the capillary tubes of a viscometer (see Table of Materials) and measure the dynamic viscosity.</li>
	<li>Plot the dynamic viscosities measured as a function of protein concentration and extrapolate to zero concentration. 
	NOTE: The extrapolation of DLS diffusion to zero concentration yields the value of translational diffusion for one single protein. The extrapolation of dynamic viscosity to zero concentration must yield the dynamic viscosity value measured experimentally for the buffer solution.</li>
</ol>
3. Collection of small-angle scattering (neutron or x-ray)
<ol>
	<li>Measure small-angle neutron scattering (SANS) and/or small-angle x-ray scattering (SAXS) (see Table of Materials) on four protein concentrations, preferably 2 mg/mL, 5 mg/mL, 10 mg/mL, and 50 mg/mL.</li>
	<li>Normalize SANS and SAXS spectra by protein concentration.</li>
	<li>Fit protein form factor P(Q) to the SANS and SAXS spectra using ensemble optimization31 and/or SasView32 software.</li>
	<li>Calculate structure factor S(Q, c) by dividing SANS and SAXS signal with P(Q) for each concentration. 
	NOTE: Readers interested in how to measure and interpret small-angle scattering data as support for NSE measurements are encouraged to thoroughly consult the references23,28,31,32,33.</li>
</ol>
4. Measurement of NSE
<ol>
	<li>Set up for the experiment and mount the sample following the steps below.
	<ol>
		<li>Select the thickness of the cell for sample loading based on the concentration of the protein solution, the temperature needed for measurement, and the amount of solution available. 
		NOTE: The present study used top loader transparent quartz containers of 40 mm x 30 mm x 4 mm.</li>
		<li>Clean the cell repeatedly, alternating between phosphate-free dish detergent (see Table of Materials), deionized water, and 70% ethanol.</li>
		<li>Dry the cell in the convection oven; do not exceed 80 &#176;C for the quartz cells.</li>
		<li>Load 4 mL of protein solution into the cell and close with caps. Use wax film or any sealant (see Table of Materials) to seal the sample cells. 
		NOTE: In the present study, 4.8 mL of solution at ~50 mg/mL was used to obtain sufficient scattering intensity.</li>
		<li>Load 4 mL of dialysis buffer into an identical container as the protein sample and seal.</li>
		<li>Transport samples to the beamline, close the shutter, and enter the spectrometer enclosure cave area34.</li>
		<li>Mount the sample cell on the aluminum sample holder by tightening the screws and the holding plates (Figure 2, left panel). 
		NOTE: One can mount two sample cells at the same time, given that the same measurement protocol is needed for all samples.</li>
		<li>Mount the graphite sample and/or Al2O3 powder sample loaded in an identical container as the protein sample. These are standards provided by the SNS-NSE beamline support.</li>
		<li>Place the sample holder by gently sliding it into the can of the temperature forcing system (TFS, see Table of Materials). 
		NOTE: TFS is the most used sample environment at SNS-NSE and pumps dry air into the sample canister to achieve the desired temperature (Figure 2, middle and right panels).</li>
		<li>Close the TFS lid and set the temperature to the desired value by accessing the interactive screen of the TFS.</li>
		<li>Mount the neutron camera (see Table of Materials) for the alignment of the samples to the beam.</li>
		<li>Sweep the instrument enclosure, evacuate, close the doors, and open the beam shutter. 
		NOTE: Sample cells are provided by the SNS-NSE beamline support. For available sample cells and the sample environment library, please refer to the SNS-NSE beamline web page7,35.</li>
	</ol>
	</li>
	<li>Collect the NSE data following the steps below.
	<ol>
		<li>Align the sample in the neutron beam using the neutron camera and the four independent sample apertures.</li>
		<li>Open the SNS-NSE data collection software36 and collect sample statistics by running diffraction scans for the desired scattering angles and wavelength.</li>
		<li>Set up the measurement parameters based on the statistics collected for each sample by editing the measurement macros provided by the assisting Instrument Scientist.</li>
		<li>Start scanning by typing the protocol name at the command prompter and acquire echoes for the sample.</li>
		<li>Start scanning and acquire echoes also for the elastic reference and the buffered solvent. Perform intermittent beam shutter operation for the sample change.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61862/61862fig02.jpg" /> 
Figure 2: NSE measurement system. Left panel: protein solution samples in quartz container mounted with screws and plates on the aluminum (Al) sample holder. The Al sample holder offers the possibility to mount two samples simultaneously inside the sample environment. Middle panel: The temperature forcing system (TFS) sample can be mounted at the sample stage the neutron beam window is on the right, while the neutron camera used for alignment is visible on the left side. Right panel: placing the sample holder with two samples into the sample can work with silicon (Si) windows. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61862/61862fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. NSE data reduction
NOTE: SNS-NSE is equipped with a dedicated software named DrSpine (data reduction for spin-echo)37,38 available on the ORNL Neutron Sciences Remote Analysis Cluster, a Quick User Guide, and built-in help support.
<ol>
	<li>Log in to Neutron Sciences Remote Analysis Cluster (see Table of Materials) with the ORNL user credentials, and press the Launch Session button.</li>
	<li>Set up the data reduction software following the steps below.
	<ol>
		<li>In the user directory, open a terminal window and type:&#160;source/SNS/software/nse/etc/setup_nse.sh.</li>
		<li>Next, type: drspine_create_env.sh.</li>
	</ol>
	</li>
	<li>Create a folder for the data reduction in the home directory and copy the provided scripts and macros from the shared directory.</li>
	<li>Edit, rename, and save the reduction macro provided accordingly.</li>
	<li>Type drspine at the command prompter and press enter to start the software reduction environment.</li>
	<li>Type &#34;the name of the reduction macro&#34; edited at Step 5.4. in the command prompter within the software environment and press Enter.</li>
</ol>
6. NSE data fitting
<ol>
	<li>Edit the python script &#34;stapler-drspine.py&#34;, provided by the assisting Instrument Scientist, with the names of the reduced file data. 
	NOTE: The python script is freely available to users of the instrument.</li>
	<li>Edit the function to fit from the library provided.</li>
	<li>Type the name of the edited script &#34;stapler-drspine.py&#34; at the command prompter and press Enter to read, fit, and plot reduced NSE data. 
	NOTE: The Instrument Scientist will provide a template for the reduction macro and the &#34;stapler-drspine.py&#34; python script that can read and fit NSE reduced data. The final reduced NSE data are in ASCII format and can be read by various preferred software.</li>
</ol>

The author declares no competing financial interests and no conflicts of interest.The manuscript's content is based on the lecture the author presented in the HANDS-Neutron Scattering Applications in Structural Biology School between 2019-2021.

NSE spectroscopy delivers a unique and detailed view of the dynamics of proteins, which other spectroscopic techniques cannot produce. Measurements over an extended time scale provide observations of both the proteins&#39; translational and rotational diffusion, as presented here. The segmental dynamics and other internal oscillations reveal themselves as a strong decay of the coherent scattering function S(Q, t) at a short time scale and are well separated from the overall diffusional relaxation processes. The ...

Probing dynamics of soft matter with neutrons 
Investigating the dynamical properties of proteins and peptides is a major part of biophysical research, and many well-developed methods exist today to access a wide range of energy landscapes1. Relating the experimentally revealed dynamics of the proteins to their biological function is a far more difficult task, requiring complex mathematical models and computer-aided dynamics simulations. The importance of neutron spectroscopy for the analysis of protein motions has been emphasized in several well-received and widely recognized studies1,2,3,4,5. Before exploring the diverse energy landscape of internal protein dynamics, a short overview of the dynamical processes in soft matter and how neutrons can access them is required.
The sensitivity of neutrons to isotopic configuration and the type of interactions they display with soft matter makes neutron scattering one of the most versatile investigation techniques6. There is a broad spectrum of correlation length scales and correlation times that neutrons can access, from nuclear excitations and atomic vibrations to collective motions and slow relaxation processes like isotropic rotations and diffusive motions. When investigating the scattered neutrons for their energy transfer, three main interactions can be distinguished: the elastic scattering, in which there is no energy exchange between incoming neutron and particle in the sample; the inelastic scattering, with a large, quantifiable energy exchange between neutron and particle; and the peculiar case of quasi-elastic scattering that designates a very small energy transfer compared to the incident neutron energy1,7. These interactions give precise information about the material investigated and form the theoretical basis of a wide variety of neutron scattering techniques.
In elastic scattering, the detector records the directions of the neutrons as a diffraction pattern, which shows the position of the sample atoms relative to one another. Information about the correlations of atomic positions is acquired (i.e., integrated intensity S(Q) concerning the momentum transfer Q, which pertains to structural information alone). This principle forms the basis of neutron diffraction8.
Complexity arises when the energy transfer is no longer zero due to excitations and internal fluctuations in the sample material. This forms the basis of neutron spectroscopy, in which the scattered neutrons are investigated as a function of both the energy transfer E and the momentum transfer Q. Dynamical and structural information is obtained. Neutron spectroscopy measures the same integrated intensity S(Q) for energy transfer (i.e., velocity change of the neutrons due to sample scattering, S(Q,&#969;) = S(Q, E), which is also referred to as the dynamic structure factor)9.
For calculating the scattering from a material, it is more adequate to use the pair correlation function7,10. In the diffraction case, the static pair correlation function G(r) gives the probability of finding the center of a particle at a given distance r from the center of another particle. The spectroscopy generalizes the static pair correlation function and includes energy/frequency/time in the scattering equation. The pair correlation function G(r) becomes a function of time G(r, t), which may be decomposed into a distinct atom pair correlation function GD(r, t), and a self-correlation function GS(r, t). These describe two types of correlations: pair-correlated motions of atoms that govern the coherent scattering, and self-correlation that governs the incoherent scattering10.
Coherent scattering is the scattering from &#34;the average&#34; and depends on the relative phase of the scattered waves. In the small-angle scattering regime, the scattered neutron waves from different scattering centers (different atoms) interfere constructively (have similar phases), and the collective motion of the atoms is observed with strong intensity enhancement. Coherent scattering essentially describes the scattering of a single neutron from all the nuclei in the sample10.
When no constructive interference occurs between the scattered neutron waves from different centers, a single atom is followed in time, and the self-correlation between the position of the atom at time t = 0 and the same atom at time t is observed. Thus, the information on the relative positions of atoms is lost, and the focus is only on local fluctuations. Scattering from local fluctuations governs incoherent scattering. Incoherent scattering is isotropic, contributes to the background signal, and degrades the signal-to-noise10,11.
Combining all of the above, we distinguish four major neutron scattering processes10: (1) elastic coherent (measures the correlations of atomic positions), (2) inelastic coherent (measures collective motions of atoms), (3) elastic incoherent (contributes to the background, reduces scattering intensity by Debye-Waller factor (DWF) and measures elastic incoherent structure factor (EISF), describing the geometry of diffusive motions in confined geometry, and (4) inelastic incoherent (measures single atom dynamics and self-correlation).
Dynamics processes that neutrons can access in biology range from the damping of low frequency atomic and molecular vibrations, the interaction of solvent molecules with bio-surfaces, and diffusion processes in the hydration layer of macromolecules and confined geometry, to short-range translational, rotational, and tumbling diffusive motions, and protein domains and allosteric motions1. The wide diversity of neutron methods and instruments for measuring protein dynamics is based on how the achromatization of the incident or outgoing neutron beam is achieved and how the energy analysis of the scattered neutrons is performed. From triple-axis to time-of-flight, backscattering, and spin-echo spectrometers, one can explore dynamical processes with characteristic times between 1 x 10-14 s and 1 x 10-6 s (femtoseconds to microseconds)12.
Oak Ridge National Laboratory, with its two renowned neutron sources, the Spallation Neutron Source - SNS13 and the High Isotope Flux Reactor - HFIR14, has one of the best suites of spectrometers for investigating dynamics in bio-materials. Some of the most eloquent examples include the use of the cold neutron chopper spectrometer (CNCS) at SNS15&#160;to investigate the dynamical perturbation of hydration water around green fluorescent protein in solution16 or the sub-picosecond collective vibrations of several proteins17. A recurring problem of inelastic neutron scattering investigations is that some biological processes are too slow to be observed. Without extreme setups that lead to a huge loss of neutron intensity, time-of-flight spectrometers are limited to 10 &#181;eV energy resolution, corresponding to a maximum time scale of ~200 ps10,11. This is not sufficient to observe large-scale motions in proteins. Therefore, instruments with higher energy resolution like the backscattering spectrometers are often needed. Combining the time-of-flight and backscattering techniques has proven powerful for investigating the change in internal dynamics of Cytochrome P450cam (CYP101), an enzyme that catalyzes the hydroxylation camphor18.
Microscopic diffusivity measured by the backscattering spectrometer at SNS-BASIS19 was surprisingly well defined and could be separated into the diffusivity of water (hydration, cytoplasmic, and bulk-like water) and the diffusivity of cell constituents in planarian flatworms, the first living animal to be studied by neutron scattering20. Backscattering is a high-resolution spectroscopic technique, but it is also limited to several &#181;eV = several nanoseconds, while the slow dynamics in biomaterials also manifest as the survival time of correlation between atomic position or spin orientations (e.g., relaxation processes, which regularly happen in the time range of ten to hundreds of nanoseconds).
Neutron spin echo spectroscopy (NSE) is the only neutron scattering technique to reach such high resolution. Unlike other neutron techniques, NSE does not require achromatization of the beam since it uses the quantum mechanical phase of the neutrons, which is their magnetic moments. The manipulation of magnetic moments allows the use of a broad neutron beam wavelength distribution, while the technique is sensitive to very small neutrons velocity changes in the order of 1 x 10-4. NSE has been successfully used to investigate the slow dynamics of proteins in solution for many proteins. Among these many pioneer studies, we acknowledge the study of the segmental flexibility of pig immunoglobulin21; the coupled domain motions in Taq polymerase22; the domain motions in the tetramer of yeast alcohol dehydrogenase23; the change of conformation in phosphoglycerate kinase upon substrate binding3; the activation of domain motions and the dynamic propagation of allosteric signals in the Na+/H+ exchange regulatory cofactor 1 (NHERF1) protein4,24,25; the dynamics of a compact state of mercuric ion reductase26; and the diffusion of hemoglobin in red blood cells27. Two more recent studies in protein dynamics have exposed the flexibility of human antibody Immunoglobulin G (IgG) as an entropic spring28 and the characteristics of solvent contribution to the dynamics of intrinsically disordered myelin basic protein (MBP)5.
The present article explains the basic principles of NSE, the multiple preparatory methods recommended for a thorough protein dynamics investigation, as well as the methodology and the experimental protocol for NSE data acquisition at the NSE spectrometer at SNS, SNS-NSE. The protocol characterizes two proteins: IgG, a regular human antibody protein, and the intrinsically disordered protein MBP. The biophysical implications, the research relevance of the examples, and the limitations of the technique are discussed briefly.
NSE spectroscopy, the method for slow dynamics measurements 
NSE is a polarized technique that uses neutron&#160;time-of-flight to measure the exchange of energy (loss of polarization) due to the quasi-elastic interaction between neutrons and atoms in a sample. At the core of NSE spectroscopy lie two basic principles: (1) the ability of the neutron spin to precess in the magnetic field with a frequency proportional to magnetic strength <img alt="Equation 1" src="/files/ftp_upload/61862/61862eq01.jpg" style="margin: auto;" />, namely the Larmor frequency29, and (b) the spin-echo or Hann echo, representing the manipulation and refocusing of the polarization signal when applying a series of radiofrequency pulses30.
The basics of the NSE process can be summarized in a few simple steps6,11 using Figure 1. (1) The neutron beam produced by the source (position 1) is polarized (position 2), guided, and transported (position 3), and arrives at the entrance of the NSE spectrometer, where it gets rotated by 90&#176; by the first pi-half flipper (position 4). (2) The polarized beam (e.g., neutron magnetic moments) becomes perpendicular to the first magnet&#39;s magnetic field lines (first precession zone, position 5) and starts to precess. (3) At the end of the magnet, neutron spins accumulate a certain precession angle proportional to the magnetic field strength and the time-of-flight spent inside (basically inversely proportional to the neutron velocity). The individual neutron velocities are encoded within their precession angle at the end of the first precession zone. (4) Close to the sample position, the pi-flipper (position 6) reverses the orientation of the spin by 180&#176;, changing the sign of the precession angle. (5) The neutrons interact with the sample&#39;s molecules (position 7) and get scattered. (6) The scattered neutrons enter and precess in the second precession zone (position 8) but become reversed-oriented. (7) Another pi-half flipper (position 9) is used to rotate the orientation of the spin from perpendicular to the horizontal direction. This will stop the precession, translating the precession angle &#966; into polarization proportional to cos(&#966;). (8) The analyzer (position 10) selects the neutrons based on one orientation. If the interaction with the sample is elastic, the neutron&#39;s velocity will not change. The neutrons will spend an identical amount of time flying in the first and second precession zones, and the accumulated precession angles are fully recovered. The full polarization is restored on the detector (position 11) as an echo of the original polarization (i.e., spin-echo). (9) However, in NSE, the scattering is quasi-elastic, so a small energy exchange between neutrons and sample molecules leads to different neutron velocities after scattering by the sample. Due to the different velocities, the neutrons will spend an additional time flying through the second precession zone and will not have properly recovered their precession angle. A partial polarization is retrieved on the detector, and the loss of polarization due to spin relaxation is proportional to the cos-Fourier-transform of the spectral function S(Q, &#969;), the intermediate scattering function F(Q, t). (10) The time parameter of the function F(Q, t) is proportional to the precession magnetic field strength. Scanning the loss of polarization as a function of magnetic field strength yields, therefore, a relaxation function that depends on the dynamical processes within the sample.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61862/61862fig01.jpg" /> 
Figure 1: Photograph of the NSE spectrometer at SNS (SNS-NSE) and neutron fly path schematic with the most important functional components. From right to left: 1 = neutron source; 2 = choppers-bender-polarizer-secondary shutter system; 3 = beam transport guides; 4 = pi/2 flipper for first 90&#176; spin-turn; 5 = first precession zone; 6 = pi flipper for 180&#176; spin-turn; 7 = sample area and sample environment (here, the cryo-furnace is shown); 8 = second precession zone; 9 = pi/2 flipper for second 90&#176; spin-turn; 10 = analyzer; 11 = detector. (Note that portions of 3, as well as 2 and 1 ,are situated behind the blue wall inside shielding; the choppers are replaced by a velocity selector for reactor-based NSE). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61862/61862fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>Bovine MBP protein solution</td><td>Sigma-Aldrich</td><td>M1891</td><td>lyophilized powder reconstituted in&amp;nbsp; D&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>D&lt;sub&gt;2&lt;/sub&gt;O - heavy water</td><td>Sigma-Aldrich</td><td>Product No. 151882</td><td>liquid</td></tr><tr><td>Dionized water</td><td>in house</td><td>-</td><td>for washing / cleanning cells</td></tr><tr><td>DLS instrument</td><td>Zetasizer Nano ZS, FZ-J&amp;uuml;lich</td><td>-</td><td>dynamic light scattering instrument</td></tr><tr><td>Elastic scattering standards</td><td>SNS-NSE, ORNL</td><td>-</td><td>Al&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;3&lt;/sub&gt;&amp;nbsp; and Graphite powders</td></tr><tr><td>Ethanol</td><td>Sigma-Aldrich</td><td>65350-M</td><td>70% ethanol for cleaning cells</td></tr><tr><td>IgG protein solution</td><td>Sigma-Aldrich</td><td>I4506</td><td>lyophilized powder reconstituted in&amp;nbsp; D&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>KWS-2 instrument</td><td>JCNS outstation at the MLZ,&amp;nbsp; Garching, Germany</td><td>-</td><td>small angle neutron instrument</td></tr><tr><td>Liquinox dish detergent</td><td>Alconox</td><td>-</td><td>Phosphate-free liquid lab glassware cleaner</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>Product No.S9390</td><td>disodium phosphate heptahydrate salt</td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>Product No.S9888</td><td>sodium chloride salt</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>Product No. S9638</td><td>monosodium phosphate monohydrate salt</td></tr><tr><td>Nanodrop spectrophotometer</td><td>Thermo Scientific</td><td>Catalog number:&amp;nbsp;ND-2000</td><td>NanoDrop 2000/2000c Spectrophotometer</td></tr><tr><td>Neutron alignment camera</td><td>NeutronOptics, Grenoble</td><td>NOG210222</td><td>100 x 100 mm camera with Sony IMX249 CMOS sensor</td></tr><tr><td>Parafilm M - wax parafilm</td><td>Bemis</td><td>Parafilm M - 5259-04LC PM996</td><td>all-purpose laboratory film in cardboard dispenser</td></tr><tr><td>Phoenix-J-NSE Spectrometer</td><td>JCNS outstation at the MLZ,&amp;nbsp; Garching, Germany</td><td>-</td><td>neutron spectrometer</td></tr><tr><td>SasView</td><td>https://www.sasview.org/</td><td></td><td></td></tr><tr><td>SAXSpace, Anton Paar instrument</td><td>FZ-J&amp;uuml;lich</td><td>-</td><td>small angle x-ray instrument</td></tr><tr><td>Slide-A-Lyzer dialysis membranes</td><td>Thermo Scientific</td><td>88400-88405</td><td>Slide-A-Lyzer mini dialysis devices tubes of 3.5 K MWCO</td></tr><tr><td>SNS Remote Analysis Cluster</td><td>Neutron Science Remote Analysis (sns.gov)</td><td>https://analysis.sns.gov</td><td></td></tr><tr><td>SNS-NSE spectrometer</td><td>ORNL, Oak Ridge, TN, USA</td><td>-</td><td>neutron spectrometer</td></tr><tr><td>Sterile syringe filters</td><td>VWR</td><td>N.A. PN:28145-501</td><td>0.2 &amp;micro;m pore size filters</td></tr><tr><td>Temperature Forcing System (TFS)</td><td>SP Scientific</td><td>Part Number 100004055</td><td>sample environment equipment</td></tr><tr><td>Urea -d4</td><td>Sigma-Aldrich</td><td>Product No. 176087</td><td>deuterated Urea salt</td></tr><tr><td>Viscometer</td><td>FZ-J&amp;uuml;lich</td><td>-</td><td>falling ball viscometer</td></tr></tbody></table>

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.

IgG protein from human serum and bovine MBP proteins were reconstituted at high concentrations (~50 mg/mL) in D2O-base buffers. Since the proteins were dissolved in high concentrations, the solutions obtained were crowded proteins solutions. The dynamics investigated using NSE suffer from the crowded environment that the proteins reside in (structure factor interactions and hydrodynamics effects)5,28,39. DLS was perfo...

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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.

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 ...

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2.0K Views.  NSD, Oak Ridge National Laboratory. 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.

Video: Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

Echo Particle Image Velocimetry

The transport of mass, momentum, and energy in fluid flows is ultimately determined by spatiotemporal distributions of the fluid velocity field.1 Consequently, a prerequisite for understanding, predicting, and controlling fluid flows is the capability to measure the velocity field with adequate spatial and temporal resolution.2 For velocity measurements in optically opaque fluids or through optically opaque geometries, echo particle image velocimetry (EPIV) is an attractive diagnostic technique to generate "instantaneous" two-dimensional fields of velocity.3,4,5,6 In this paper, the operating protocol for an EPIV system built by integrating a commercial medical ultrasound machine7 with a PC running commercial particle image velocimetry (PIV) software8 is described, and validation measurements in Hagen-Poiseuille (i.e., laminar pipe) flow are reported.
For the EPIV measurements, a phased array probe connected to the medical ultrasound machine is used to generate a two-dimensional ultrasound image by pulsing the piezoelectric probe elements at different times. Each probe element transmits an ultrasound pulse into the fluid, and tracer particles in the fluid (either naturally occurring or seeded) reflect ultrasound echoes back to the probe where they are recorded. The amplitude of the reflected ultrasound waves and their time delay relative to transmission are used to create what is known as B-mode (brightness mode) two-dimensional ultrasound images. Specifically, the time delay is used to determine the position of the scatterer in the fluid and the amplitude is used to assign intensity to the scatterer. The time required to obtain a single B-mode image, t, is determined by the time it take to pulse all the elements of the phased array probe. For acquiring multiple B-mode images, the frame rate of the system in frames per second (fps) = 1/&delta;t. (See 9 for a review of ultrasound imaging.)
For a typical EPIV experiment, the frame rate is between 20-60 fps, depending on flow conditions, and 100-1000 B-mode images of the spatial distribution of the tracer particles in the flow are acquired. Once acquired, the B-mode ultrasound images are transmitted via an ethernet connection to the PC running the PIV commercial software. Using the PIV software, tracer particle displacement fields, D(x,y)[pixels], (where x and y denote horizontal and vertical spatial position in the ultrasound image, respectively) are acquired by applying cross correlation algorithms to successive ultrasound B-mode images.10 The velocity fields, u(x,y)[m/s], are determined from the displacements fields, knowing the time step between image pairs, &Delta;T[s], and the image magnification, M[meter/pixel], i.e., u(x,y) = MD(x,y)/&Delta;T. The time step between images &Delta;T = 1/fps + D(x,y)/B, where B[pixels/s] is the time it takes for the ultrasound probe to sweep across the image width. In the present study, M = 77[&mu;m/pixel], fps = 49.5[1/s], and B = 25,047[pixels/s]. Once acquired, the velocity fields can be analyzed to compute flow quantities of interest.

An echo particle image velocimetry (EPIV) system capable of acquiring two-dimensional fields of velocity in optically opaque fluids or through optically opaque geometries is described, and validation measurements in pipe flow are reported.

The transport of mass, momentum, and energy in fluid flows is ultimately determined by spatiotemporal distributions of the fluid velocity field.1 ...

Angle-resolved Photoemission Spectroscopy At Ultra-low Temperatures

The physical properties of a material are defined by its electronic structure. Electrons in solids are characterized by energy (&omega;) and momentum (k) and the probability to find them in a particular state with given &omega; and k is described by the spectral function A(k, &omega;). This function can be directly measured in an experiment based on the well-known photoelectric effect, for the explanation of which Albert Einstein received the Nobel Prize back in 1921. In the photoelectric effect the light shone on a surface ejects electrons from the material. According to Einstein, energy conservation allows one to determine the energy of an electron inside the sample, provided the energy of the light photon and kinetic energy of the outgoing photoelectron are known. Momentum conservation makes it also possible to estimate k relating it to the momentum of the photoelectron by measuring the angle at which the photoelectron left the surface. The modern version of this technique is called Angle-Resolved Photoemission Spectroscopy (ARPES) and exploits both conservation laws in order to determine the electronic structure, i.e. energy and momentum of electrons inside the solid. In order to resolve the details crucial for understanding the topical problems of condensed matter physics, three quantities need to be minimized: uncertainty* in photon energy, uncertainty in kinetic energy of photoelectrons and temperature of the sample.
In our approach we combine three recent achievements in the field of synchrotron radiation, surface science and cryogenics. We use synchrotron radiation with tunable photon energy contributing an uncertainty of the order of 1 meV, an electron energy analyzer which detects the kinetic energies with a precision of the order of 1 meV and a He3 cryostat which allows us to keep the temperature of the sample below 1 K. We discuss the exemplary results obtained on single crystals of Sr2RuO4 and some other materials. The electronic structure of this material can be determined with an unprecedented clarity.

The overall goal of this method is to determine the low-energy electronic structure of solids at ultra-low temperatures using Angle-Resolved Photoemission Spectroscopy with synchrotron radiation.

The physical properties of a material are defined by its  electronic structure. Electrons in solids are characterized by energy (&omega;)  and momentum ...

Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique amino acid sequence, it is only realized by the folding of the polypeptide chain into a single defined three-dimensional arrangement or more commonly into an ensemble of interconverting conformations. Investigating the connection between protein conformation and its function is therefore essential for a complete understanding of how proteins are able to fulfill their great variety of tasks. One possibility to study conformational changes a protein undergoes while progressing through its functional cycle is hydrogen-1H/2H-exchange in combination with high-resolution mass spectrometry (HX-MS). HX-MS is a versatile and robust method that adds a new dimension to structural information obtained by e.g. crystallography. It is used to study protein folding and unfolding, binding of small molecule ligands, protein-protein interactions, conformational changes linked to enzyme catalysis, and allostery. In addition, HX-MS is often used when the amount of protein is very limited or crystallization of the protein is not feasible. Here we provide a general protocol for studying protein dynamics with HX-MS and describe as an example how to reveal the interaction interface of two proteins in a complex. &#160;&#160;

Protein conformation and dynamics are key to understanding the relationship between protein structure and function. Hydrogen exchange coupled with high-resolution mass spectrometry is a versatile method to study the conformational dynamics of proteins as well as characterizing protein-ligand and protein-protein interactions, including contact interfaces and allosteric effects.

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique ...

Chemistry

Gradient Echo Quantum Memory in Warm Atomic Vapor

Gradient echo memory (GEM) is a protocol for storing optical quantum states of light in atomic ensembles. The primary motivation for such a technology is that quantum key distribution (QKD), which uses Heisenberg uncertainty to guarantee security of cryptographic keys, is limited in transmission distance. The development of a quantum repeater is a possible path to extend QKD range, but a repeater will need a quantum memory. In our experiments we use a gas of rubidium 87 vapor that is contained in a warm gas cell. This makes the scheme particularly simple. It is also a highly versatile scheme that enables in-memory refinement of the stored state, such as frequency shifting and bandwidth manipulation. The basis of the GEM protocol is to absorb the light into an ensemble of atoms that has been prepared in a magnetic field gradient. The reversal of this gradient leads to rephasing of the atomic polarization and thus recall of the stored optical state. We will outline how we prepare the atoms and this gradient and also describe some of the pitfalls that need to be avoided, in particular four-wave mixing, which can give rise to optical gain.

The gradient echo memory is a protocol for storing optical quantum states of light in atomic ensembles. Quantum memory is a key element of a quantum repeater, which can extend the range of quantum key distribution. We outline the operation of the scheme when implemented in a 3-level atomic ensemble.

Gradient  echo memory (GEM) is a protocol for storing optical quantum states of light in  atomic ensembles. The primary motivation for such a technology ...

Measuring Material Microstructure Under Flow Using 1-2 Plane Flow-Small Angle Neutron Scattering

A new small-angle neutron scattering (SANS) sample environment optimized for studying the microstructure of complex fluids under simple shear flow is presented. The SANS shear cell consists of a concentric cylinder Couette geometry that is sealed and rotating about a horizontal axis so that the vorticity direction of the flow field is aligned with the neutron beam enabling scattering from the 1-2 plane of shear (velocity-velocity gradient, respectively). This approach is an advance over previous shear cell sample environments as there is a strong coupling between the bulk rheology and microstructural features in the 1-2 plane of shear. Flow-instabilities, such as shear banding, can also be studied by spatially resolved measurements. This is accomplished in this sample environment by using a narrow aperture for the neutron beam and scanning along the velocity gradient direction. Time resolved experiments, such as flow start-ups and large amplitude oscillatory shear flow are also possible by synchronization of the shear motion and time-resolved detection of scattered neutrons. Representative results using the methods outlined here demonstrate the useful nature of spatial resolution for measuring the microstructure of a wormlike micelle solution that exhibits shear banding, a phenomenon that can only be investigated by resolving the structure along the velocity gradient direction. Finally, potential improvements to the current design are discussed along with suggestions for supplementary experiments as motivation for future experiments on a broad range of complex fluids in a variety of shear motions.

A shear cell is developed for small-angle neutron scattering measurements in the velocity-velocity gradient plane of shear and is used to characterize complex fluids. Spatially resolved measurements in the velocity gradient direction are possible for studying shear-banding materials. Applications include investigations of colloidal dispersions, polymer solutions, and self-assembled structures.

A new small-angle neutron scattering (SANS) sample environment optimized for studying the microstructure of complex fluids under simple shear flow is ...

Using Neutron Spin Echo Resolved Grazing Incidence Scattering to Investigate Organic Solar Cell Materials

The spin echo resolved grazing incidence scattering (SERGIS) technique has been used to probe the length-scales associated with irregularly shaped crystallites. Neutrons are passed through two well defined regions of magnetic field; one before and one after the sample. The two magnetic field regions have opposite polarity and are tuned such that neutrons travelling through both regions, without being perturbed, will undergo the same number of precessions in opposing directions. In this case the neutron precession in the second arm is said to &#34;echo&#34; the first, and the original polarization of the beam is preserved. If the neutron interacts with a sample and scatters elastically the path through the second arm is not the same as the first and the original polarization is not recovered. Depolarization of the neutron beam is a highly sensitive probe at very small angles (&#60;50&#160;&#956;rad) but still allows a high intensity, divergent beam to be used. The decrease in polarization of the beam reflected from the sample as compared to that from the reference sample can be directly related to structure within the sample.
In comparison to scattering observed in neutron reflection measurements the SERGIS signals are often weak and are unlikely to be observed if the in-plane structures within the sample under investigation are dilute, disordered, small in size and polydisperse or the neutron scattering contrast is low. Therefore, good results will most likely be obtained using the SERGIS technique if the sample being measured consist of thin films on a flat substrate and contain scattering features that contains a high density of moderately sized features (30 nm to 5 &#181;m) which scatter neutrons strongly or the features are arranged on a lattice. An advantage of the SERGIS technique is that it can probe structures in the plane of the sample.

Progress has been made in utilizing spin echo resolved grazing incidence scattering (SERGIS) as a neutron scattering technique to probe the length-scales in irregular samples. Crystallites of [6,6]-phenyl-C61-butyric acid methyl ester have been probed using the SERGIS technique and the results confirmed by optical and atomic force microscopy.

The spin echo resolved grazing incidence scattering (SERGIS) technique has been used to probe the length-scales associated with irregularly shaped ...

Fast Imaging Technique to Study Drop Impact Dynamics of Non-Newtonian Fluids

In the field of fluid mechanics, many dynamical processes not only occur over a very short time interval but also require high spatial resolution for detailed observation, scenarios that make it challenging to observe with conventional imaging systems. One of these is the drop impact of liquids, which usually happens within one tenth of millisecond. To tackle this challenge, a fast imaging technique is introduced that combines a high-speed camera (capable of up to one million frames per second) with a macro lens with long working distance to bring the spatial resolution of the image down to 10 &#181;m/pixel. The imaging technique enables precise measurement of relevant fluid dynamic quantities, such as the flow field, the spreading distance and the splashing speed, from analysis of the recorded video. To demonstrate the capabilities of this visualization system, the impact dynamics when droplets of non-Newtonian fluids impinge on a flat hard surface are characterized. Two situations are considered: for oxidized liquid metal droplets we focus on the spreading behavior, and for densely packed suspensions we determine the onset of splashing. More generally, the combination of high temporal and spatial imaging resolution introduced here offers advantages for studying fast dynamics across a wide range of microscale phenomena.

Drop impact of non-Newtonian fluids is a complex process since different physical parameters influence the dynamics over a very short time (less than one tenth of a millisecond). A fast imaging technique is introduced in order to characterize the impact behaviors of different non-Newtonian fluids.

In the field of fluid mechanics, many dynamical processes not only occur over a very short time interval but also require high spatial resolution for ...

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without labeling other than deuterium. In particular, neutron backscattering spectroscopy records the scattering signals at multiple scattering angles simultaneously and is well suited to study the dynamics of biological systems on the ps-ns timescale. By employing D2O-and possibly deuterated buffer components-the method allows monitoring of both center-of-mass diffusion and backbone and side-chain motions (internal dynamics) of proteins in liquid state.
Additionally, hydration water dynamics can be studied by employing powders of perdeuterated proteins hydrated with H2O. This paper presents the workflow employed on the instrument IN16B at the Institut Laue-Langevin (ILL) to investigate protein and hydration water dynamics. The preparation of solution samples and hydrated protein powder samples using vapor exchange is explained. The data analysis procedure for both protein and hydration water dynamics is described for different types of datasets (quasielastic spectra or fixed-window scans) that can be obtained on a neutron backscattering spectrometer.
The method is illustrated with two studies involving amyloid proteins. The aggregation of lysozyme into &#181;m sized spherical aggregates-denoted particulates-is shown to occur in a one-step process on the space and time range probed on IN16B, while the internal dynamics remains unchanged. Further, the dynamics of hydration water of tau was studied on hydrated powders of perdeuterated protein. It is shown that translational motions of water are activated upon the formation of amyloid fibers. Finally, critical steps in the protocol are discussed as to how neutron scattering is positioned regarding the study of dynamics with respect to other experimental biophysical methods.

Neutron backscattering spectroscopy offers a nondestructive and label-free access to the ps-ns dynamics of proteins and their hydration water. The workflow is presented with two studies on amyloid proteins: on the time-resolved dynamics of lysozyme during aggregation and on the hydration water dynamics of tau upon fiber formation.

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without ...

Biochemistry

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 ...

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral budding, among other vital cellular processes. For efficient biological activity, cell membranes should be rigid enough to maintain the integrity of the cell and its compartments yet fluid enough to allow membrane components, such as proteins and functional domains, to diffuse and interact. This delicate balance of elastic and fluid membrane properties, and their impact on biological function, necessitate a better understanding of collective membrane dynamics over mesoscopic length and time scales of key biological processes, e.g., membrane deformations and protein binding events. Among the techniques that can effectively probe this dynamic range is neutron spin echo (NSE) spectroscopy. Combined with deuterium labeling, NSE can be used to directly access bending and thickness fluctuations as well as mesoscopic dynamics of select membrane features. This paper provides a brief description of the NSE technique and outlines the procedures for performing NSE experiments on liposomal membranes, including details of sample preparation and deuteration schemes, along with instructions for data collection and reduction. The paper also introduces data analysis methods used to extract key membrane parameters, such as the bending rigidity modulus, area compressibility modulus, and in-plane viscosity. To illustrate the biological importance of NSE studies, select examples of membrane phenomena probed by NSE are discussed, namely, the effect of additives on membrane bending rigidity, the impact of domain formation on membrane fluctuations, and the dynamic signature of membrane-protein interactions.

This paper describes the protocols for sample preparation, data reduction, and data analysis in neutron spin echo (NSE) studies of lipid membranes. Judicious deuterium labeling of lipids enables access to different membrane dynamics on mesoscopic length and time scales, over which vital biological processes occur.

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral ...

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

Summary

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Chapters in this video

Echo Particle Image Velocimetry

Angle-resolved Photoemission Spectroscopy At Ultra-low Temperatures

Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

Gradient Echo Quantum Memory in Warm Atomic Vapor

Measuring Material Microstructure Under Flow Using 1-2 Plane Flow-Small Angle Neutron Scattering

Using Neutron Spin Echo Resolved Grazing Incidence Scattering to Investigate Organic Solar Cell Materials

Fast Imaging Technique to Study Drop Impact Dynamics of Non-Newtonian Fluids

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

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

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions