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.remotexs.ntu.edu.sg/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 main limitations of the NSE technique are the extended measurement time and the large sample volumes needed to acquire a good statistical signal. This can pose a challenge to measuring relevant macromolecular systems that exhibit low concentrations of the mobile species and reduced lifetimes.
Internal dynamics of IgG 
IgG protein has a Y-shaped structure specific to antibodies that can be approximated by three large fragments connected by flexible linkers. A mathematical model of three sizable particles residing in harmonic potential was assumed for this kind of structure. The linkers were approximated by elastic springs that give a fixed equilibrium position but allow fluctuations as a form of Brownian motion. Three degrees of freedom were permitted for each fragment around the relative equilibrium28. The internal dynamics represented by Fint(Q, t) in Equation 2 were described by the Ornstein-Uhlenbeck process50,51. The model allows the calculation of the mean square displacement of the fragments in the potential dip, the timescale of different motions, the friction, and the force constants that occur. The amplitude of the internal motion was approximated by normal modes analysis, considering different degrees of motion for each fragment. An extensive description of the model and the resulting mathematical parameters is out of scope here but can be found in detail in reference28. In this case study of IgG, a clear signature of the internal dynamics was observed on the timescale of several nanoseconds. The calculated spring constant of the linker appears to be realistically comparable to an entropic spring of similar length. The observed effective friction seems close to the friction of a free unbound IgG fragment. The pre-existing equilibrium hypothesis was validated, with assumed coexisting &#34;open&#34; or &#34;closed&#34; configurations of IgGs that are in equilibrium exhibiting different binding sites and binding specificities28. This information may be relevant for understanding the mechanics of antibodies and whether that mechanical behavior can be used to improve biocompatibility and bioavailability.
Internal dynamics of MBP 
MBP structure has a compact globular core that allows strong stretching and bending motions with its flexible random coil ends. MBP dynamics were interpreted using flexible polymer models with and without the addition of internal friction28,39. As in the case of IgG, the observations of the internal dynamics of MBP can be reconciled within the theory of Brownian motion, in which a larger amplitude of motion results in longer relaxation time with influences from the internal friction within the protein chain. The increased relaxation time with a larger amplitude of motions but smaller friction can be described using the same Ornstein-Uhlenbeck process50,51 of Brownian motion in a harmonic potential. Internal motions with large amplitudes but slow relaxation times become active upon full denaturation of MBP from its native state by reducing the restoring forces of chain configuration (dihedral potentials) and by smoothing the local energy barriers. Detailed descriptions of the MBP internal dynamics in the native and denatured state can be further found in references28,39. In the study of MBP, the investigation using NSE revealed a dynamical behavior pointing toward intermediary compactness of the protein between random coil polymers and globular proteins. The significant contribution of internal protein dynamics with a relaxation rate of several nanoseconds was governed by low-frequency collective stretching and bending motions5. Description of the native MBP dynamics by models from polymer theory breakdown was provided due to a large value of protein internal friction. In denatured MBP, the internal friction within the protein chain is reduced, and the polymer chain character of the relaxation spectrum prevails. The high flexibility of the structural ensemble motions might help increase the accessible protein surface, thereby facilitating the interaction with different binding partners.
Dynamical models 
While NSE as a technique is experimentally unique in assessing low-frequency harmonic motions in biological macromolecules and molecular subunits, the analysis of the intermediate scattering function requires a comparison with neutron spectra derived from various mathematical models. The model presented here and developed by Biehl et al.28 for concentrated protein solutions uses an elastic network approximation, in which the intermediate scattering function is a convolution of proper modular functions, and Brownian oscillators represent the internal dynamics. This is one of the very few available models for analyzing the segmental dynamics of proteins. Another well-established model for dilute protein solutions is the model proposed by Bu et al.52, which is a robust model that uses statistical mechanics and does not require complex fits or multiple parameters. A similar approach based on dynamic decoupling approximation can also be applied to dilute systems to characterize the effective diffusion as a sum of self-translational and internal motions53. Several of our references can provide comprehensive reporting on these models and their limitations. We strongly recommend Fitter et al.1 and Liu et al.54.

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.remotexs.ntu.edu.sg/files/ftp_upload/61862/61862fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bovine MBP protein solution</td>
			<td>Sigma-Aldrich</td>
			<td>M1891</td>
			<td>lyophilized powder reconstituted in&#160; D2O</td>
		</tr>
		<tr>
			<td>D2O - 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&#252;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>Al2O3&#160; 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&#160; D2O</td>
		</tr>
		<tr>
			<td>KWS-2 instrument</td>
			<td>JCNS outstation at the MLZ,&#160; 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>Na2HPO4&#183;7H2O</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>NaH2PO4&#183;H2O</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:&#160;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,&#160; 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&#252;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 &#181;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&#252;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 performed on a concentration series for each protein to account for crowding effects. The extrapolation to zero concentration of the translational diffusion coefficients measured by DLS yields the value of translational diffusion for a single protein. The dynamic viscosity as a function of protein concentration must also be measured prior to NSE to account for the hydrodynamic interactions. The extrapolation to zero concentration of the dynamic viscosities as a function of concentration needs to yield the value that can be experimentally measured for the buffer solution of each prepared protein sample.
The shape and structure of proteins in concentrated solution were assessed prior to any NSE experiment to gain insights on the protein form factor and the strength of the structure factor, which might influence how the proteins move in solution. Small-angle scattering was the method of choice for these investigations. In the present study, the structural conformation of IgG protein in solution was observed by SAXS, and the structure of MBP was assessed by SANS. The scattering intensity (Q) measured (Steps 3.3.-3.4.) is proportional to the product between form factor P(Q) and the structure factor S(Q,c) weighted by the number of particles (Equation 1)5,28,39:
I (Q) = N &#8729; P (Q) &#8729;S (Q, c)&#160; &#160;(1)
SAXS was measured on a Kratky-type SAXS instrument40 with an X-ray wavelength of 0.154 nm for IgG, and SANS was measured41 for MBP with a neutron wavelength of 4.5 &#197;. The experimental small-angle intensities I(Q) were background and solvent corrected, scaled by solution concentration, and the form factor was calculated using freely available software packages like Ensemble modeling-EOM and SasView24,25,29,39,42,43. Further, the structure factor for each protein was obtained from Equation 1.
For the present study, the NSE experiments were performed with two spin-echo spectrometers: the SNS-NSE instrument34 in Figure 1 and the Phoenix-J-NSE instrument44. Incident wavelengths between 8 &#197; - 12 &#197;, with Fourier times between 0.1 ns &#8804; tmax&#160;&#8804; 130 ns for several Q measurements between 0.05 &#197;&#8722;1- 0.2 &#197;&#8722;1, were measured. The difference between the two spectrometers is a vital point in the understanding of NSE science, and they represent both the old concept of a so-called &#34;classic spin echo&#34; for the static neutron source of a reactor and the new &#34;time-of-flight concept&#34; for the pulsating neutron source. The Phoenix-J-NSE spectrometer is a classic-type NSE, in which a velocity selector selects a particular wavelength. To allow variations in the neutron velocities, a &#177;10% wavelength bandwidth is introduced. Despite the very narrow energy band used for a single scan, the situation of the Phoenix-J-NSE spectrometer at a high flux reactor source ensures space-vector and correlations time ranges are covered in relatively short measurement times. The SNS-NSE spectrometer is the new-generation, ultra-high resolution, choppers neutron spectrometer, with a wavelength span of 2 &#197; &#60; &#955; &#60; 14 &#197; and a simultaneous wavelength bandwidth of 2.4 &#197; - 3.6 &#197;, depending on the position of the spectrometer. Due to this wide bandwidth, high data collection efficiency is accomplished, allowing nearly gapless coverage of a broad wave-vector time range with only few scattering angle settings. The selection of wavelength band in the SNS-NSE is made by a chopper system consisting of four choppers. Both NSE spectrometers discussed here have precession fields based on superconducting technology with high magnetic field homogeneity, novel state-of-the-art field correction elements for stray field corrections, and novel polarizing benders41,42.
The NSE spectra were measured at 10 &#176;C for MBP protein and 25 &#176;C for IgG1 protein. The coherent intermediate scattering intensity I(Q, t) / I(Q, t = 0) measured by NSE is the contributive result of all the dynamic processes in the sample that happen within the investigated time scale. This contains the internal protein dynamics, the overall translational diffusion, the rotational and tumbling diffusion, and the segmental motions within the protein molecule itself. A simplified model assumes that the internal dynamics and the translational and rotational diffusions are totally decoupled5,45,46,47,48 and can be characterized separately. For the single particle, the coherent scattering function can be written as the product (Equation 2)23,48:
F (Q, t) = Ftrans(Q, t) &#8729;Frot(Q, t) &#8729;Fint(Q, t)&#160; &#160;(2)
where the translational diffusion term for a single rigid protein is an exponential function of the translational diffusion coefficient DT (Equation 3)23,48:
Ftrans(Q, t) = exp(&#8722;Q2&#8729; DT&#8729; t)&#160; &#160;(3)
For large concentrations of proteins, the translational diffusion coefficient DT is influenced by protein-protein interactions, quantified by the structure factor S(Q) and by hydrodynamic interactions described by the hydrodynamic function HT(Q) (Equation 4)23,48:
DT(Q) = DT0 &#8729;HT(Q, c) / S(Q, c)&#160; &#160;(4)
In Equation 4, DT0 is the extrapolated diffusion value to zero concentration from the DLS measurements (Step 2.2.). HT is calculated as HT = 1 -c&#8729; [&#951;], where c is the protein concentration, [&#951;]&#160;is the intrinsic viscosity calculated by [&#951;] = (&#951; - &#951;0) /&#951;0 &#8729; c, and &#951;0 is the measured dynamic viscosity (Steps 2.3.-2.4.). The structure factor S(Q, c) was obtained from SAXS measurements for the IgG protein and SANS measurements for the MBP protein.
Figure 3&#160;displays examples of intermediate scattering functions I(Q, t) / I(Q, t = 0), as measured by NSE for IgG and MBP proteins. A clear deviation from a simple diffusion-like relaxation process was observed on the short Fourier time scale, &#60;25 ns, indicating the accessibility of protein internal dynamics by NSE and the need for a more complex model to describe the observed dynamical processes.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61862/61862fig03.jpg" /> 
Figure 3: NSE relaxation spectra for IgG and MBP proteins. The IgG data are shown here fitted just by a single-exponential function to reveal the deviation from diffusion at shorter Fourier times. The deviation was observed for both proteins. In contrast, the MBP data are shown here in the full-relaxation range, fitted by the model developed byBiehl45. The model implies coarse-graining and enables the simultaneous fit of short, intermediate, and long Fourier time regimes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61862/61862fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Therefore, the scattering functions were fitted by atomic modeling of both proteins using models developed by Biehl, described in references18,25,38,43,44,46. The intermediate scattering function was calculated by coarse-graining to a couple of hundred individual grains, as opposed to all-atom simulation, to reduce computational load and time, and the MMTK49 software library (freely accessible) was used to access atomic coordinates and implement trans-rotational motions on protein domains and fragments. The results of the intermediate scattering function calculation for both proteins were in excellent agreement with the experimental NSE data and proved that the slower dynamics observed at long Fourier times can be attributed to the overall translational and rotational diffusion processes, while the fast dynamics observed at short time scales can be attributed to the dynamics of protein domains.

Watch this Scientific Journal Video about Study of Protein Dynamics via Neutron Spin Echo Spectroscopy at JoVE.com

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 ...

study-of-protein-dynamics-via-neutron-spin-echo-spectroscopy

Nanyang Technological University

Research

JoVE Journal

Engineering

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 ...

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 ...

Proton Transfer and Protein Conformation Dynamics in Photosensitive Proteins by Time-resolved Step-scan Fourier-transform Infrared Spectroscopy

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards understanding its mechanism. Protonation and conformational changes affect the vibration pattern of amino acid side chains and of the peptide bond, respectively, both of which can be probed by infrared (IR) difference spectroscopy. For proteins whose function can be repetitively and reproducibly triggered by light, it is possible to obtain infrared difference spectra with (sub)microsecond resolution over a broad spectral range using the step-scan Fourier transform infrared technique. With ~102-103 repetitions of the photoreaction, the minimum number to complete a scan at reasonable spectral resolution and bandwidth, the noise level in the absorption difference spectra can be as low as ~10-4, sufficient to follow the kinetics of protonation changes from a single amino acid. Lower noise levels can be accomplished by more data averaging and/or mathematical processing. The amount of protein required for optimal results is between 5-100 &#181;g, depending on the sampling technique used. Regarding additional requirements, the protein needs to be first concentrated in a low ionic strength buffer and then dried to form a film. The protein film is hydrated prior to the experiment, either with little droplets of water or under controlled atmospheric humidity. The attained hydration level (g of water / g of protein) is gauged from an IR absorption spectrum. To showcase the technique, we studied the photocycle of the light-driven proton-pump bacteriorhodopsin in its native purple membrane environment, and of the light-gated ion channel channelrhodopsin-2 solubilized in detergent.

Key steps of protein function, in particular backbone conformational changes and proton transfer reactions, often take place in the microsecond to millisecond time scale. These dynamical processes can be studied by time-resolved step-scan Fourier-transform infrared spectroscopy, in particular for proteins whose function is triggered by light.

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards ...

In Situ Neutron Powder Diffraction Using Custom-made Lithium-ion Batteries

Li-ion batteries are widely used in portable electronic devices and are considered as promising candidates for higher-energy applications such as electric vehicles.1,2 However, many challenges, such as energy density and battery lifetimes, need to be overcome before this particular battery technology can be widely implemented in such applications.3 This research is challenging, and we outline a method to address these challenges using in situ NPD to probe the crystal structure of electrodes undergoing electrochemical cycling (charge/discharge) in a battery. NPD data help determine the underlying structural mechanism responsible for a range of electrode properties, and this information can direct the development of better electrodes and batteries.
We briefly review six types of battery designs custom-made for NPD experiments and detail the method to construct the &#8216;roll-over&#8217; cell that we have successfully used on the high-intensity NPD instrument, WOMBAT, at the Australian Nuclear Science and Technology Organisation (ANSTO). The design considerations and materials used for cell construction are discussed in conjunction with aspects of the actual in situ NPD experiment and initial directions are presented on how to analyze such complex in situ data.

In Situ Neutron Powder Diffraction Using Custom-made Lithium-ion Batteries

We describe the design and construction of an electrochemical cell for the examination of electrode materials using in situ neutron powder diffraction (NPD). We briefly comment on alternate in situ NPD cell designs and discuss methods for the analysis of the corresponding in situ NPD data produced using this cell.

Li-ion batteries are widely used in portable electronic devices and are considered as promising candidates for higher-energy applications such as electric ...

Recombination Dynamics in Thin-film Photovoltaic Materials via Time-resolved Microwave Conductivity

A method for investigating recombination dynamics of photo-induced charge carriers in thin film semiconductors, specifically in photovoltaic materials such as organo-lead halide perovskites is presented. The perovskite film thickness and absorption coefficient are initially characterized by profilometry and UV-VIS absorption spectroscopy. Calibration of both laser power and cavity sensitivity is described in detail. A protocol for performing Flash-photolysis Time Resolved Microwave Conductivity (TRMC) experiments, a non-contact method of determining the conductivity of a material, is presented. A process for identifying the real and imaginary components of the complex conductivity by performing TRMC as a function of microwave frequency is given. Charge carrier dynamics are determined under different excitation regimes (including both power and wavelength). Techniques for distinguishing between direct and trap-mediated decay processes are presented and discussed. Results are modelled and interpreted with reference to a general kinetic model of photoinduced charge carriers in a semiconductor. The techniques described are applicable to a wide range of optoelectronic materials, including organic and inorganic photovoltaic materials, nanoparticles, and conducting/semiconducting thin films.

A Time Resolved Microwave Conductivity technique for investigating direct and trap-mediated recombination dynamics and determining carrier mobilities of thin film semiconductors is presented here.

A method for investigating recombination dynamics of photo-induced charge carriers in thin film semiconductors, specifically in photovoltaic materials ...

Dielectric RheoSANS &#8212; Simultaneous Interrogation of Impedance, Rheology and Small Angle Neutron Scattering of Complex Fluids

A procedure for the operation of a new dielectric RheoSANS instrument capable of simultaneous interrogation of the electrical, mechanical and microstructural properties of complex fluids is presented. The instrument consists of a Couette geometry contained within a modified forced convection oven mounted on a commercial rheometer. This instrument is available for use on the small angle neutron scattering (SANS) beamlines at the National Institute of Standards and Technology (NIST) Center for Neutron Research (NCNR). The Couette geometry is machined to be transparent to neutrons and provides for measurement of the electrical properties and microstructural properties of a sample confined between titanium cylinders while the sample undergoes arbitrary deformation. Synchronization of these measurements is enabled through the use of a customizable program that monitors and controls the execution of predetermined experimental protocols. Described here is a protocol to perform a flow sweep experiment where the shear rate is logarithmically stepped from a maximum value to a minimum value holding at each step for a specified period of time while frequency dependent dielectric measurements are made. Representative results are shown from a sample consisting of a gel composed of carbon black aggregates dispersed in propylene carbonate. As the gel undergoes steady shear, the carbon black network is mechanically deformed, which causes an initial decrease in conductivity associated with the breaking of bonds comprising the carbon black network. However, at higher shear rates, the conductivity recovers associated with the onset of shear thickening. Overall, these results demonstrate the utility of the simultaneous measurement of the rheo-electro-microstructural properties of these suspensions using the dielectric RheoSANS geometry.

Here, we present a procedure for the measurement of simultaneous impedance, rheology and neutron scattering from soft matter materials under shear flow.

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Echo Particle Image Velocimetry

Angle-resolved Photoemission Spectroscopy At Ultra-low Temperatures

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

Proton Transfer and Protein Conformation Dynamics in Photosensitive Proteins by Time-resolved Step-scan Fourier-transform Infrared Spectroscopy

In Situ Neutron Powder Diffraction Using Custom-made Lithium-ion Batteries

Recombination Dynamics in Thin-film Photovoltaic Materials via Time-resolved Microwave Conductivity

Dielectric RheoSANS — Simultaneous Interrogation of Impedance, Rheology and Small Angle Neutron Scattering of Complex Fluids