R. Ashkar thanks M. Nagao, L.-R. Stingaciu, and P. Zolnierczuk for many useful discussions and for their frequent assistance with NSE experiments on their respective beamlines. The authors acknowledge the use of neutron spin echo spectrometers at NIST and ORNL. The NSE spectrometer at NIST is supported by the Center for High Resolution Neutron Scattering, a partnership between the National Institute of Standards and Technology and the National Science Foundation under agreement no. DMR-1508249. The NSE spectrometer at ORNL&#39;s Spallation Neutron Source is supported by the Scientific User Facilities Division, Office of Basic Energy Sciences, US Department of Energy. Oak Ridge National Laboratory is managed by UT-Battelle, LLC under US DOE Contract No. DE-AC05-00OR22725.

The authors declare no conflicts of interest and have nothing to disclose.

NSE is a powerful and unique technique in measuring mesoscopic dynamics of lipid membranes under various conditions. The effective utilization of NSE depends on sample quality, neutron contrast, and the range of accessible dynamics that can be probed for a given sample. Thus, several critical steps are required for performing successful NSE experiments and collecting high-quality data. A key step in ensuring the effective use of neutron beam time during an NSE experiment is to characterize the liposomal suspensions with ...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62396">Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions</a>. The Introduction, Protocol, and Representative Results sections have been updated.
In the Introduction, the fith pargraph was updated from:
Besides direct access to the length and time scale of membrane dynamics, NSE has the inherent capabilities of neutron isotope sensitivity52. Specifically, the ability of neutrons to interact differently with the isotopes of hydrogen, the most abundant element in biological systems, results in a different neutron scattering length density,34 or NSLD (the equivalent of the optical index of refraction50), when protium is substituted by deuterium. This enables an approach known as contrast variation, which is commonly used to highlight specific membrane features or conceal others &#8212;&#160;the latter scenario is referred to as contrast matching. A frequent application of contrast variation/matching is the substitution of water (NSLD = -0.56 &#215; 10-6&#160;&#197;-2) by heavy water or D2O (NSLD = 6.4&#160;&#215; 10-6&#160;&#197;-2) to amplify the neutron signal from protiated lipid membranes (NSLD ~ 2 &#215; 10-6&#160;&#197;-2). This approach is highly effective in studies of membrane structure because the penetration of D2O into the headgroup region of the membrane allows accurate determination of the membrane thicknesses (see Figure 2A, left panel) and of the location of different lipid subgroups when more sophisticated models are applied53,54. This paper highlights some examples on the use of contrast variation for studies of collective dynamics in biomimetic membranes and select membrane features.
to:
Besides direct access to the length and time scale of membrane dynamics, NSE has the inherent capabilities of neutron isotope sensitivity52. Specifically, the ability of neutrons to interact differently with the isotopes of hydrogen, the most abundant element in biological systems, results in a different neutron scattering length density,34 or NSLD (the equivalent of the optical index of refraction50), when protium is substituted by deuterium. This enables an approach known as contrast variation, which is commonly used to highlight specific membrane features or conceal others &#8212;&#160;the latter scenario is referred to as contrast matching. A frequent application of contrast variation/matching is the substitution of water (NSLD = -0.56 &#215; 10-6&#160;&#197;-2) by heavy water or D2O (NSLD = 6.4&#160;&#215; 10-6&#160;&#197;-2) to amplify the neutron signal from protiated lipid membranes (NSLD ~ 0&#160;&#215; 10-6&#160;&#197;-2). This approach is highly effective in studies of membrane structure because the penetration of D2O into the headgroup region of the membrane allows accurate determination of the membrane thicknesses (see Figure 2A, left panel) and of the location of different lipid subgroups when more sophisticated models are applied53,54. This paper highlights some examples on the use of contrast variation for studies of collective dynamics in biomimetic membranes and select membrane features.
In the Protocol, step 1.1 was updated from:
For bending fluctuation measurements, make fully protiated liposomes in D2O (D 99.9%) or D2O-buffer (e.g., phosphate buffer prepared with D2O instead of H2O). Use fully protiated DMPC (C36H72NO8P) and DSPC (C44H88NO8P) with <img alt="Equation 1" src="/files/ftp_upload/62396/62396eq01v2.jpg" /> 133.4 mg, where XDMPC and XDSPC are the mole fractions of DMPC and DSPC, here set to 0.7 and 0.3, respectively, and MwDMPC&#160;and MwDSPC are the molar weights given by 677.9 g/mol and 790.1 g/mol, respectively. Similarly, mDSPC = 66.6 mg. This deuteration scheme increases the scattering contrast between the membrane (NSLD&#160;~ 2&#160;&#215; 10-6&#160;&#197;-2) and the deuterated buffer (NSLD ~ 6.4 &#215; 10-6&#160;&#197;-2) and amplifies the signal from membrane undulations (see Figure 2A left panel).
to:
For bending fluctuation measurements, make fully protiated liposomes in D2O (D 99.9%) or D2O-buffer (e.g., phosphate buffer prepared with D2O instead of H2O). Use fully protiated DMPC (C36H72NO8P) and DSPC (C44H88NO8P) with <img alt="Equation 1" src="/files/ftp_upload/62396/62396eq01v2.jpg" /> 133.4 mg, where XDMPC and XDSPC are the mole fractions of DMPC and DSPC, here set to 0.7 and 0.3, respectively, and MwDMPC&#160;and MwDSPC are the molar weights given by 677.9 g/mol and 790.1 g/mol, respectively. Similarly, mDSPC = 66.6 mg. This deuteration scheme increases the scattering contrast between the membrane (NSLD&#160;~ 0&#160;&#215; 10-6&#160;&#197;-2) and the deuterated buffer (NSLD ~ 6.4 &#215; 10-6&#160;&#197;-2) and amplifies the signal from membrane undulations (see Figure 2A left panel).
In the Representative Results, the fist pagargaph was updted&#160;from:
NSE studies accessing bending fluctuations are typically performed over a Q-range of ~ (0.04 - 0.2) &#197;-1. This Q-range corresponds to intermediate length scales between the membrane thickness and the liposomal radius, where bending dynamics dominate. Measurement over an extended Q-range can give access to additional dynamic modes, including liposomal diffusion and intramembrane dynamics. For more details on the cross-over in membrane dynamics accessed by NSE, check these relevant publications25,71. It is important to emphasize that NSE signals are proportional to: <img alt="Equation 5" src="/files/ftp_upload/62396/62396eq05v2.jpg" />, where&#160;Icoh and Iinc&#160;are, respectively, the coherent and incoherent scattering intensity from the sample. Therefore, it is advisable to prepare NSE liposomal samples in deuterated buffers (i.e., buffers prepared with D2O instead of H2O) to minimize the incoherent scattering signal, mainly contributed by the hydrogen content of the sample. However, in some cases intermediate deuteration schemes (i.e., using mixtures of D2O and H2O) might be necessary to obtain optimal contrast conditions. Typically, NSE measurements of membrane bending fluctuations are performed on fully protiated liposomes in deuterated buffer, referred to as fully contrasted liposomes in Figure 5. This deuteration scheme results in a large NSLD difference between the membrane core (~2 &#215; 10-6 &#197;-2) and its deuterated fluid environment (~6.4 &#215; 10-6 &#197;-2), which significantly enhances the scattering signal from the liposomal membranes and improves the measurement statistics of bending dynamics. This contrast scheme (Figure 2A left panel) is frequently utilized in studies of bending rigidity of lipid membranes with single38,72 and multiple39,66 lipid components and in studies of membrane softening/stiffening by biological inclusions (e.g., cholesterol, drug molecules, peptides/proteins)36,37,73,74,75, and synthetic additives (e.g., nanoparticles)76,77.
to:
NSE studies accessing bending fluctuations are typically performed over a Q-range of ~ (0.04 - 0.2) &#197;-1. This Q-range corresponds to intermediate length scales between the membrane thickness and the liposomal radius, where bending dynamics dominate. Measurement over an extended Q-range can give access to additional dynamic modes, including liposomal diffusion and intramembrane dynamics. For more details on the cross-over in membrane dynamics accessed by NSE, check these relevant publications25,71. It is important to emphasize that NSE signals are proportional to: <img alt="Equation 5" src="/files/ftp_upload/62396/62396eq05v2.jpg" />, where&#160;Icoh and Iinc&#160;are, respectively, the coherent and incoherent scattering intensity from the sample. Therefore, it is advisable to prepare NSE liposomal samples in deuterated buffers (i.e., buffers prepared with D2O instead of H2O) to minimize the incoherent scattering signal, mainly contributed by the hydrogen content of the sample. However, in some cases intermediate deuteration schemes (i.e., using mixtures of D2O and H2O) might be necessary to obtain optimal contrast conditions. Typically, NSE measurements of membrane bending fluctuations are performed on fully protiated liposomes in deuterated buffer, referred to as fully contrasted liposomes in Figure 5. This deuteration scheme results in a large NSLD difference between the membrane core (~0&#160;&#215; 10-6 &#197;-2) and its deuterated fluid environment (~6.4 &#215; 10-6 &#197;-2), which significantly enhances the scattering signal from the liposomal membranes and improves the measurement statistics of bending dynamics. This contrast scheme (Figure 2A left panel) is frequently utilized in studies of bending rigidity of lipid membranes with single38,72 and multiple39,66 lipid components and in studies of membrane softening/stiffening by biological inclusions (e.g., cholesterol, drug molecules, peptides/proteins)36,37,73,74,75, and synthetic additives (e.g., nanoparticles)76,77.
In the Representative Reults, Figure 2 was updated from:
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62396/62396fig2v4.jpg" /> 
Figure 2:&#160;Examples of possible deuteration schemes in NSE experiments on lipid membranes. (A) Left: Fully contrasted membranes, e.g., protiated membranes in deuterated buffer, showing the NSLD profile along the normal to the membrane surface. The difference in the NSLD between the headgroup (~2&#160;&#215; 10-2 &#197;-2)&#160;and tail region (~4.5 &#215; 10-6 &#197;-2) of the membrane is due to the headgroup hydration with deuterated buffer. Right: Tail-contrast matched membranes such that the hydrocarbon tail region of the membrane has the same NSLD as the buffer, as shown in the corresponding NSLD profile along the membrane normal. (B) Domain-forming membranes with two neutron contrast schemes where the domains (center) or the matrix (left) are contrast-matched to the buffer, enabling selective studies of matrix or domain dynamics, respectively. This figure has been modified from Nickels et al., JACS 201541. (C) Asymmetric membranes prepared by cyclodextrin exchange between protiated and deuterated lipid vesicles, resulting in the deuteration of one membrane leaflet while keeping the other leaflet protiated. This allows studies of the bending dynamics of the protiated leaflet and provides insights into the mechanical coupling between opposing leaflets in asymmetric membranes. This figure has been modified from Rickeard et al., Nanoscale 202040. <a href="https://www.jove.com/files/ftp_upload/62396/62396fig2v4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
to:
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62396/62396fig2v5.jpg" /> 
Figure 2:&#160;Examples of possible deuteration schemes in NSE experiments on lipid membranes. (A) Left: Fully contrasted membranes, e.g., protiated membranes in deuterated buffer, showing the NSLD profile along the normal to the membrane surface. The difference in the NSLD between the tail region&#160;(~0&#160;&#215; 10-2 &#197;-2)&#160;and headgroup&#160;region (~4.5 &#215; 10-6 &#197;-2) of the membrane is due to the headgroup hydration with deuterated buffer. Right: Tail-contrast matched membranes such that the hydrocarbon tail region of the membrane has the same NSLD as the buffer, as shown in the corresponding NSLD profile along the membrane normal. (B) Domain-forming membranes with two neutron contrast schemes where the domains (center) or the matrix (left) are contrast-matched to the buffer, enabling selective studies of matrix or domain dynamics, respectively. This figure has been modified from Nickels et al., JACS 201541. (C) Asymmetric membranes prepared by cyclodextrin exchange between protiated and deuterated lipid vesicles, resulting in the deuteration of one membrane leaflet while keeping the other leaflet protiated. This allows studies of the bending dynamics of the protiated leaflet and provides insights into the mechanical coupling between opposing leaflets in asymmetric membranes. This figure has been modified from Rickeard et al., Nanoscale 202040. <a href="https://www.jove.com/files/ftp_upload/62396/62396fig2v5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62396">Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions</a>. The Introduction, Protocol, and Representative Results sections have been updated.

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

The understanding of cell membranes and their function has remarkably evolved over the last few decades. The former view of cell membranes as passive lipid bilayers that define cell boundaries and house membrane proteins1 has gradually transformed into a dynamic model in which lipid bilayers play an important role in regulating vital biological processes, including cellular signaling, molecular exchange, and protein function &#8212;&#160;to name a few2,3,4,5,6. This realization that cell membranes are highly dynamic, constantly undergoing remodeling and molecular redistribution, has urged scientific explorations beyond equilibrium structures of membranes7,8,9. Accordingly, multiple approaches have been developed to study the various dynamic modes in biological and bioinspired lipid membranes. To date, the majority of these studies have primarily focused on diffusive molecular motions10,11,12,13 and macroscopic shape fluctuations14,15,16, leaving a significant gap in understanding intermediate membrane dynamics, i.e., collective fluctuations of lipid assemblies consisting of few 10-100s of lipid molecules. These dynamics occur over length scales of few tens to few 100 &#197; and over time scales of sub-ns to few hundred ns (see Figure 1), referred to here as mesoscopic scales. It is indeed on these scales that key biological activity takes place at the membrane level17. This includes viral budding18, channel gating19, and membrane-protein interactions20. It is also important to point out that the energy landscape of membrane proteins21,22 shows that conformational changes in proteins &#8212;&#160;necessary for their regulatory role &#8212;&#160;happen over the ns time scales23 of collective membrane fluctuations, further emphasizing the importance of mesoscopic dynamics in the biological function of cell membranes and their bioinspired analogs20. This paper focuses on the two primary mesoscopic dynamic modes in lipid membranes, namely, bending fluctuations and thickness fluctuations.
The main challenge in directly probing these fluctuation modes is the difficulty in simultaneously accessing their spatial and temporal scales using standard spectroscopy methods. The other challenge is that direct contact techniques could impact the same fluctuations they are meant to measure16. This is further exacerbated by the compositional and structural complexity of biological membranes24,25, which results in non-homogeneous membrane features, including lipid domain formation26,27,28,29,30 and membrane asymmetry31,32,33&#8212; demanding selective probes to understand the dynamics of different membrane features. Fortunately, these challenges can be overcome with non-invasive neutron spectroscopy methods, such as neutron spin echo (NSE), which inherently access the required length and time scales, and further enable studies of selective membrane features without changing their physicochemical environment34. Indeed, over the last few years NSE spectroscopy has evolved into a unique and powerful probe of collective membrane dynamics35. Results from NSE studies on lipid membranes have produced new insights into mechanical36,37 and viscoelastic38,39 properties of lipid membranes and have shed new light on their potential role in biological function40,41.
The NSE spectroscopy technique is based on an interferometric instrument design, first proposed by Mezei42, using a series of spin-flippers and magnetic coils to control the precession of the neutron spin as neutrons traverse the instrument. The design rests on magnetic mirroring of the magnetic field elements with respect to the sample position (Figure 1A). This implies that in the absence of energy exchange between the neutron and the sample, the neutron performs the same number of spin precessions, in opposite directions, in the first and second half of the instrument (notice the &#960;-flipper between the two precession coils). As a result, the final spin state of the neutron remains unchanged relative to the initial state - a phenomenon referred to as spin-echo (see transparent neutron in Figure 1A). However, when the neutron energetically interacts with the sample, the energy exchange modifies the number of spin precessions in the second half of the instrument, leading to a different final spin state (see Figure 1A). This is experimentally detected as a loss in polarization, as will be shown later in this paper. For more details on the NSE technique, the reader is referred to dedicated technical papers42,43,44,45.
Here, a simplified description is presented to provide a rough estimate of the length and time scales accessible with NSE. The length scales are determined by the range of achievable wavevector transfers, Q = 4&#960; sin &#952;/&#955;, where 2&#952; is the scattering angle and&#160;&#955; is the neutron wavelength. One can see that Q&#160;is set by the wavelength range and the extent of rotation of the second arm of the spectrometer (see Figure 1A). A typical Q-range on NSE spectrometers is ~0.02-2 &#197;-1&#160;46,47, and up to 0.01-4 &#197;-1 with recent upgrades48,49, corresponding to spatial scales of ~1-600 &#197;. On the other hand, the accessible time scale is calculated from the total precession angle (or phase) acquired by the neutron within the magnetic precession coils, and is found to be50: <img alt="Equation 12" src="/files/ftp_upload/62396/62396eq12v5.jpg" />. In this expression, t is the Fourier time defined as <img alt="Equation 13" src="/files/ftp_upload/62396/62396eq13v2.jpg" />, where&#160;<img alt="Equation 50" src="/files/ftp_upload/62396/62396eq50v2.jpg" />&#160;is the neutron gyromagnetic ratio, <img alt="Equation 51" src="/files/ftp_upload/62396/62396eq51v6.jpg" />&#160;is the coil length, and <img alt="Equation 52" src="/files/ftp_upload/62396/62396eq52v6.jpg" />&#160;is the strength of the coil&#39;s magnetic field. It is worth pointing out that the Fourier time is a quantity that is strictly dependent on the instrument geometry, magnetic field strength, and neutron wavelength. For instance, using neutrons of wavelength&#160;<img alt="Equation 70" src="/files/ftp_upload/62396/62396eq70v6.jpg" /> = 8&#160;&#197; and instrument settings of <img alt="Equation 51" src="/files/ftp_upload/62396/62396eq51v6.jpg" />&#160;= 1.2 m and <img alt="Equation 52" src="/files/ftp_upload/62396/62396eq52v6.jpg" /> = 0.4 T, the Fourier time is calculated to be t ~ 50 ns. Experimentally, the Fourier time is tuned by changing the current in the precession coils (i.e., magnetic field strength) or using different neutron wavelengths, resulting in typical NSE time scales of ~ 1 ps to 100 ns. However, recent upgrades in NSE spectrometers have enabled access to longer Fourier times, up to ~400 ns on the J-NSE-Phoenix spectrometer at the Heinz Maier-Leibnitz Zentrum51 and the SNS-NSE spectrometer at Oak Ridge National Lab48, and up to ~1,000 ns at the IN15 NSE spectrometer at the Institut Laue-Langevin (ILL)49.
Besides direct access to the length and time scale of membrane dynamics, NSE has the inherent capabilities of neutron isotope sensitivity52. Specifically, the ability of neutrons to interact differently with the isotopes of hydrogen, the most abundant element in biological systems, results in a different neutron scattering length density,34 or NSLD (the equivalent of the optical index of refraction50), when protium is substituted by deuterium. This enables an approach known as contrast variation, which is commonly used to highlight specific membrane features or conceal others &#8212;&#160;the latter scenario is referred to as contrast matching. A frequent application of contrast variation/matching is the substitution of water (NSLD = -0.56 &#215; 10-6&#160;&#197;-2) by heavy water or D2O (NSLD = 6.4&#160;&#215; 10-6&#160;&#197;-2) to amplify the neutron signal from protiated lipid membranes (NSLD ~ 0&#160;&#215; 10-6&#160;&#197;-2). This approach is highly effective in studies of membrane structure because the penetration of D2O into the headgroup region of the membrane allows accurate determination of the membrane thicknesses (see Figure 2A, left panel) and of the location of different lipid subgroups when more sophisticated models are applied53,54. This paper highlights some examples on the use of contrast variation for studies of collective dynamics in biomimetic membranes and select membrane features.
Here, the effectiveness of NSE in providing unique insights into dynamical and functional membrane properties is illustrated through tangible examples of NSE studies on model and biologically relevant lipid membrane systems with emphasis on mesoscale dynamics in free-standing membranes, in the form of liposomal suspensions. For NSE measurements of in-plane membrane dynamics, the reader is referred to dedicated publications on grazing-incidence neutron spin-echo spectroscopy (GINSES)55,56 and other studies of aligned multilamellar membrane stacks57,58,59,60.
For simplicity, this paper highlights three different schemes of membrane deuteration illustrated on a well-studied domain-forming, or phase separating, lipid bilayer system of 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) mixtures61,62. The two lipids are characterized by a mismatch in their hydrocarbon chain length (14 carbons/tail in DMPC vs 18 carbons/tail in DSPC) and their gel-fluid transition temperature&#160;(Tm, DMPC = 23 &#176;C vs Tm, DSPC = 55 &#176;C). This results in lateral phase-separation in DMPC:DSPC membranes at temperatures between the upper and lower transition temperatures of the mixture63. The deuteration schemes considered here are chosen to demonstrate the different dynamic modes accessible in NSE measurements on liposomal membranes, namely, bending fluctuations, thickness fluctuations, and selective bending/thickness fluctuations of lateral domains. All lipid compositions are reported for DMPC:DSPC bilayers prepared at a mole fraction of 70:30, using commercially available protiated and perdeuterated variants of DMPC and DSPC. All sample preparation steps are based on 4 mL of liposomal suspension, in D2O, with a lipid concentration of 50 mg/mL, for a total lipid mass of Mtot&#160;= 200 mg per sample.

<table><tbody><tr><td>Chloroform (biotech grade)</td><td>Sigma Aldrich</td><td>496189</td><td>Biotech. grade, &amp;ge;99.8%, contains 0.5-1.0% ethanol as stabilizer</td></tr><tr><td>Circulating water bath</td><td>Julabo</td><td>SE-12</td><td>Heating Circulator with smart pump, programmable temperature settings, and external sensor connection for measurement and control</td></tr><tr><td>Deuterium Oxide</td><td>Cambridge Isotopes Laboratories</td><td>DLM-4</td><td>Deuterated water; Heavy water (D&lt;sub&gt;2&lt;/sub&gt;O) (D, 99.9%)</td></tr><tr><td>Digital Semi-Microbalance</td><td>Mettler Toledo</td><td>MS105</td><td>Semi-micro balance with 120 g capacity, 0.01 mg readability, high resolution weighing cell, ergonomic doors, and pipette-check application</td></tr><tr><td>Ethanol (molecular biology grade)</td><td>Sigma Aldrich</td><td>E7023</td><td>200 proof ethanol for molecular biology applications</td></tr><tr><td>Glass Pipets</td><td>VWR</td><td>36360-536</td><td>Disposable Soda Lime glass Pasteur pipets</td></tr><tr><td>Glass Vials</td><td>Thermo Scientific</td><td>B7990-1</td><td>Borosilicate glass vials with PTFE/Silione septum caps</td></tr><tr><td>Lab grade freezer</td><td>Fisher Scientific</td><td>IU2886D</td><td>Ultra-low temprature freezer (-86 to -50 C) for long-term storage of lipids and proteins</td></tr><tr><td>Lipids (protaited or perdeuterated)</td><td>Avanti Polar Lipids</td><td>varies by lipid</td><td>Lipids can be purchased from Avanti in powder form or in a chloroform solution with the required amounts and deuteration schemes.</td></tr><tr><td>Millipore water purifier</td><td>Millipore Sigma</td><td>ZRQSVP3US</td><td>Direct-Q&amp;reg;&amp;nbsp;3 UV Water Purification System which deliver both pure and ultrapure water with a built-in UV lamp to reduce the levels of organics for biological&amp;nbsp; applications</td></tr><tr><td>Mini Extruder Set</td><td>Avanti Polar Lipids</td><td>610020</td><td>Mini-extruder set includes mini-extruder, heating block, 2 GasTight Syringes, and 2 O-rings, Polycarbonate Membranes, and Filter Supports</td></tr><tr><td>Quick Connect Fittings</td><td>Grainger</td><td>2YDA1 and 2YDA7</td><td>Push-button tube fittings for QuickConnect water circulation applications, e.g. high temperature vesicle extrusion</td></tr><tr><td>Syringe Pump</td><td>SyringePump.com</td><td>New Era-1000</td><td>Fully programmable syringe pump for infusion and withdrawal; programs up to 41 pumping phases with adjustable pumping rates, dispensed volumes, and extrusion cycles</td></tr><tr><td>Ultrasonic bath</td><td>Fisher Scientific</td><td>CPX2800</td><td>Temperature controlled ultra sonic bath with programmable functionality for degassing and ultrasonic applications</td></tr><tr><td>Vacuum Oven</td><td>Thermo Scientific</td><td>3608</td><td>0.7 cu ft vaccum oven with built-in-high-limit thermostat guards against overheating</td></tr><tr><td>Vortex Mixer</td><td>Fisher Scientific</td><td>02-215-414</td><td>Variable speed, analog control that allows low rpm start-up for gentle shaking or high-speed mixing for vigorous vortexing of samples</td></tr></tbody></table>

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.

NSE studies accessing bending fluctuations are typically performed over a Q-range of ~ (0.04 - 0.2) &#197;-1. This Q-range corresponds to intermediate length scales between the membrane thickness and the liposomal radius, where bending dynamics dominate. Measurement over an extended Q-range can give access to additional dynamic modes, including liposomal diffusion and intramembrane dynamics. For more details on the cross-over in membrane dynamics accessed by NSE, check these relevant publications<sup class="xr...

Watch this Scientific Journal Video about Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions at JoVE.com

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and 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 ...

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3.9K Views.  Virginia Tech. 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.

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

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Biochemistry

Site Directed Spin Labeling and EPR Spectroscopic Studies of Pentameric Ligand-Gated Ion Channels

Ion channel gating is a stimulus-driven orchestration of protein motions that leads to transitions between closed, open, and desensitized states. Fundamental to these transitions is the intrinsic flexibility of the protein, which is critically modulated by membrane lipid-composition. To better understand the structural basis of channel function, it is necessary to study protein dynamics in a physiological membrane environment. Electron Paramagnetic Resonance (EPR) spectroscopy is an important tool to characterize conformational transitions between functional states. In comparison to NMR and X-ray crystallography, the information obtained from EPR is intrinsically of lower resolution. However, unlike in other techniques, in EPR there is no upper-limit to the molecular weight of the protein, the sample requirements are significantly lower, and more importantly the protein is not constrained by the crystal lattice forces. Therefore, EPR is uniquely suited for studying large protein complexes and proteins in reconstituted systems. In this article, we will discuss general protocols for site-directed spin labeling and membrane reconstitution using a prokaryotic proton-gated pentameric Ligand-Gated Ion Channel (pLGIC) from Gloeobacter violaceus (GLIC) as an example. A combination of steady-state Continuous Wave (CW) and Pulsed (Double Electron Electron Resonance-DEER) EPR approaches will be described that will enable a complete quantitative characterization of channel dynamics.

This article describes methods for site-directed spin labeling and reconstitution of pentameric ligand-gated channels for Electron Paramagnetic Resonance studies. This protocol can be adapted for any membrane protein. The reconstitution method described here can also be used for patch-clamp measurements of macroscopic and single-channel currents in a defined lipid system.

Ion channel gating is a stimulus-driven orchestration of protein motions that leads to transitions between closed, open, and desensitized states. ...

Chemistry

Contrast-Matching Detergent in Small-Angle Neutron Scattering Experiments for Membrane Protein Structural Analysis and Ab Initio Modeling

The biological small-angle neutron scattering instrument at the High-Flux Isotope Reactor of Oak Ridge National Laboratory is dedicated to the investigation of biological materials, biofuel processing, and bio-inspired materials covering nanometer to micrometer length scales. The methods presented here for investigating physical properties (i.e., size and shape) of membrane proteins (here, MmIAP, an intramembrane aspartyl protease from Methanoculleus marisnigri) in solutions of micelle-forming detergents are well-suited for this small-angle neutron scattering instrument, among others. Other biophysical characterization techniques are hindered by their inability to address the detergent contributions in a protein-detergent complex structure. Additionally, access to the Bio-Deuteration Lab provides unique capabilities for preparing large-scale cultivations and expressing deuterium-labeled proteins for enhanced scattering signal from the protein. While this technique does not provide structural details at high-resolution, the structural knowledge gap for membrane proteins contains many addressable areas of research without requiring near-atomic resolution. For example, these areas include determination of oligomeric states, complex formation, conformational changes during perturbation, and folding/unfolding events. These investigations can be readily accomplished through applications of this method.

Contrast-Matching Detergent in Small-Angle Neutron Scattering Experiments for Membrane Protein Structural Analysis and Ab Initio Modeling

This protocol demonstrates how to obtain a low-resolution ab initio model and structural details of a detergent-solubilized membrane protein in solution using small-angle neutron scattering with contrast-matching of the detergent.

The biological small-angle neutron scattering instrument at the High-Flux Isotope Reactor of Oak Ridge National Laboratory is dedicated to the ...

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...

The Microfluidic Probe: Operation and Use for Localized Surface Processing

Microfluidic devices allow assays to be performed using minute amounts of sample and have recently been used to control the microenvironment of cells. Microfluidics is commonly associated with closed microchannels which limit their use to samples that can be introduced, and cultured in the case of cells, within a confined volume. On the other hand, micropipetting system have been used to locally perfuse cells and surfaces, notably using push-pull setups where one pipette acts as source and the other one as sink, but the confinement of the flow is difficult in three dimensions. Furthermore, pipettes are fragile and difficult to position and hence are used in static configuration only. 
The microfluidic probe (MFP) circumvents the constraints imposed by the construction of closed microfluidic channels and instead of enclosing the sample into the microfluidic system, the microfluidic flow can be directly delivered onto the sample, and scanned across the sample, using the MFP. . The injection and aspiration openings are located within a few tens of micrometers of one another so that a microjet injected into the gap is confined by the hydrodynamic forces of the surrounding liquid and entirely aspirated back into the other opening. The microjet can be flushed across the substrate surface and provides a precise tool for localized deposition/delivery of reagents which can be used over large areas by scanning the probe across the surface. 
In this video we present the microfluidic probe1 (MFP). We explain in detail how to assemble the MFP, mount it atop an inverted microscope, and align it relative to the substrate surface, and finally show how to use it to process a substrate surface immersed in a buffer.

In this video we present the microfluidic probe1 (MFP). We explain in detail how to assemble the MFP, mount it atop an inverted microscope, and align it relative to the substrate surface, and finally show how to use it to process a substrate surface immersed in a buffer solution.

Microfluidic devices allow assays to be performed using minute amounts of sample and have recently been used to control the microenvironment of cells. ...

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

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

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

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

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

Engineering

Measuring the Time-Evolution of Nanoscale Materials with Stopped-Flow and Small-Angle Neutron Scattering

This paper presents the use of a stopped-flow small-angle neutron-scattering (SANS) sample environment to quickly mix liquid samples and study nanoscale kinetic processes on time scales of seconds to minutes. The stopped-flow sample environment uses commercially available syringe pumps to mix the desired volumes of liquid samples that are then injected through a dynamic mixer into a quartz glass cell in approximately 1 s. Time-resolved SANS data acquisition is synced with the sample mixing to follow the evolution of the nanostructure in solution after mixing. 
To make the most efficient use of neutron beam time, we use a series of flow selector valves to automatically load, rinse, and dry the cell between measurements, allowing for repeated data collection throughout multiple sample injections. Sample injections are repeated until sufficient neutron scattering statistics are collected. The mixing setup can be programmed to systematically vary conditions to measure the kinetics at different mixing ratios, sample concentrations, additive concentrations, and temperatures. The minimum sample volume required per injection is approximately 150 &#181;L depending on the path length of the quartz cell. 
Representative results using this stopped-flow sample environment are presented for rapid lipid exchange kinetics in the presence of an additive, cyclodextrin. The vesicles exchange outer-leaflet (exterior) lipids on the order of seconds and fully exchange both interior and exterior lipids within hours. Measuring lipid exchange kinetics requires in situ mixing to capture the faster (seconds) and slower (minutes) processes and extract the kinetic rate constants. The same sample environment can also be used to probe molecular exchange in other types of liquid samples such as lipid nanoparticles, proteins, surfactants, polymers, emulsions, or inorganic nanoparticles. Measuring the nanoscale structural transformations and kinetics of exchanging or reacting systems will provide new insights into processes that evolve at the nanoscale.

This protocol presents the use of a stopped-flow sample environment to quickly mix multiple liquid solutions in situ during a small-angle neutron scattering measurement and to study kinetic processes on nanometer length scales and second time scales.

This paper presents the use of a stopped-flow small-angle neutron-scattering (SANS) sample environment to quickly mix liquid samples and study nanoscale ...

High-Resolution Studies of Proteins in Natural Membranes by Solid-State NMR

Membrane proteins are vital for cell function and thus represent important drug targets. Solid-state Nuclear Magnetic Resonance (ssNMR) spectroscopy offers a unique access to probe the structure and dynamics of such proteins in biological membranes of increasing complexity. Here, we present modern solid-state NMR spectroscopy as a tool to study structure and dynamics of proteins in natural lipid membranes and at atomic scale. Such spectroscopic studies profit from the use of high-sensitivity ssNMR methods, i.e., proton-(1H)-detected ssNMR and DNP (Dynamic Nuclear Polarization) supported ssNMR. Using bacterial outer membrane beta-barrel protein BamA and the ion channel KcsA, we present methods to prepare isotope-labeled membrane proteins and to derive structural and motional information by ssNMR.

This work details robust basic routines on how to prepare isotope-labeled membrane protein samples and analyze them at high-resolution with modern solid-state NMR spectroscopy methods.

Membrane proteins are vital for cell function and thus represent important drug targets. Solid-state Nuclear Magnetic Resonance (ssNMR) spectroscopy ...

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 ...

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

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Chapters in this video

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Site Directed Spin Labeling and EPR Spectroscopic Studies of Pentameric Ligand-Gated Ion Channels

Contrast-Matching Detergent in Small-Angle Neutron Scattering Experiments for Membrane Protein Structural Analysis and Ab Initio Modeling

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

The Microfluidic Probe: Operation and Use for Localized Surface Processing

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

Measuring the Time-Evolution of Nanoscale Materials with Stopped-Flow and Small-Angle Neutron Scattering

High-Resolution Studies of Proteins in Natural Membranes by Solid-State NMR

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