Name: neutron spin echo spectroscopy as a unique probe for lipid membrane dynamics and membrane-protein interactions
Uploaded: 2021-05-27T14:30:00
Description: 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

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.

1. Deuteration scheme required for the experiment
<ol>
	<li>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 X<su...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chloroform (biotech grade)</td>
			<td>Sigma Aldrich</td>
			<td>496189</td>
			<td>Biotech. grade, &#8805;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 (D2O) (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&#174;&#160;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&#160; 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 ...

This protocol describes sample preparation and data reduction required for successful neutron spin echo studies of collective membrane fluctuations of relevance to biological functions and practical applications of lipid membranes. NSE directly accesses membrane dynamics on the nanosecond timescales of key membrane functions with the unique advantage of isotope sensitivity for probing selective membrane features inaccessible with other techniques. Studies of membrane dynamics on the nanoscale are essential to understanding the molecular mechanism's underlying membrane properties and membrane-protein interactions implicated in various cell pathologies.The method provides researchers new to NSE with detailed guidelines on designing, preparing, and characterizing lipid vesicles for successful experiments and subsequent data reduction analysis and interpretation. Demonstrating the procedure will be Teshani Kumarage and Julie Nguyen, a graduate student and an undergraduate student from the laboratory. Working inside a hood, prepare the lipid suspension by dissolving accurately-weighed lipids in one milliliter of solvent with manual mixing.Dry the lipid solution by gently streaming an inert gas in the vial while slowly rotating it at an angle. To thoroughly remove the residual solvent, place the vials overnight in a vacuum oven at 35 degrees Celsius. On the next day, hydrate the lipid film with two milliliters of heavy water to obtain a lipid concentration of 50 milligrams per milliliter and vortex the hydrated lipid suspension until the lipid film is fully dissolved.Next, perform five freeze-thaw cycles by storing the vial of hydrated lipid suspension at minus 80 degrees Celsius until frozen. And then transferring it to a 35 degrees Celsius water bath to thaw the lipid suspension. Vortex the thawed suspension until homogenous before proceeding to the next cycle.Before starting the experiment, assemble the extruder setup using a poly-bicarbonate membrane between two membrane supports and add two paper filters on each side to provide additional support. Use airtight glass syringes to hydrate the polycarbonate membrane by passing 0.3 milliliters of heavy water through the membrane assembly several times. After hydrating the membrane, insert a one milliliter gas-tight syringe with a prepared milky white color lipid solution into one end and an empty syringe into the opposite end of the extruder apparatus.Once the syringes are connected, place the assembly into the extruder block. Program the pump by holding down the Rate button to enter the extrusion rate and press the Diameter button to enter the syringe diameter. Then, press Withdraw until the light turns on.Press start and wait for the sample to start dispensing into the empty syringe. Hit the Stop button just before the sample syringe is fully empty. Record the dispensed volume, then hold the Rate button until phase one appears on the screen.Keeping the Withdraw light off, press the volume button to enter the dispensed volume recorded earlier. Press the Rate button again and use the right most up arrow to access phase two. Press volume to enter the same value of the dispensed volume recorded earlier.In this phase, press the Withdraw button until the Withdraw light is on. Repeat the cycle for phase three by pressing the volume button until LP:SE appears on the screen and set it to 20. Finally, press the Rate button, access phase four, and hit the volume button to get to the Stop function to finish the pump setup.After the pump is programmed, press Start to begin the extrusion cycle. Perform 15 to 20 extrusion cycles before collecting the transparent opal blue extruded lipid suspension in a clean vial for measurements. Open the DAVE software and select reduce NSE data from the Data Reduction menu.Upload the data files over different Q-values using the Open echo Files from the file menu. The uploaded files will show up under the available data sets. Right click on the selected file and label it according to the measurement it corresponds to like Sample, Cell, or Resolution.Using the Data Set tab, group the detector pixels in 2 X 2 to improve the signal to noise ratio. Apply the same binning to all files of Resolution, Cell, and Sample. Inspect the data over all pixel groups and mask those with poor signals by pressing the End key on the keyboard.Press Enter to access a pop-up window to apply the same mask to all four of your times. Masked pixels will turn green. Ensure that the collected data is in the form of an echo signal.Start fitting the resolution file from the uploaded file list by right clicking on the desired file and selecting Fit Operations, Fit Echoes Resolution from the pop-up menu. Ensure that the fits of the echo signals yield reasonable fitting parameters. To inspect the error associated with each fitting parameter over the entire detector, select Image Options, and then select the fitting parameter of interest.Then, right click on the detector image to access a pop-up window showing an error bar map. If the fit over a specific pixel is unsatisfactory, refit the signal over that pixel by selecting it, pressing the Fitting tab, and then pressing Fit Pixel. Input new starting parameters for the phase and period in the Fitting tab.Reduce the sample file by selecting the corresponding file from the uploaded and labeled file list. Inspect all pixels and mask the poor ones as described above. Then, right click on the file and select Fit Operations, Import Phases.For the resolution file, fit echo signals as described before with the values of the period unchanged and echo phase point imported from the resolution fits. Input the beam center for all data files by accessing the General tab and entering X and Y beam center values recorded from the experiment. Once the fits are complete, calculate the normalized intermediate scattering function by clicking right on the desired sample file from the list of fitted files and selecting Calculate I of Q from the pop-up menu.Enter the required information for the resolution and cell files and the number of Q-arcs in the pop-up window, then press OK to view results. Notice that the data in cyan shows a poor signal due to detector edge effects and should be eliminated when compiling different Q data sets. Finally, save the reduced data sets as ASCII files and save the entire session as a DAVE project in the desired folder.In this study, the NSE measurements of liposomal samples prepared with different deuteration schemes were carried out. NSE measurements of membrane bending fluctuations are performed on fully contrasted liposomes. This deuteration scheme results in a large scattering length difference between the membrane core and deuterated fluid environment, significantly enhancing the scattering signal from the liposomal membranes and improving the measurement statistics of bending dynamics.On the other hand, NSE measurements of membrane thickness fluctuations of liposomes show deviations relative to the Q to the third dependence of bending fluctuations. To isolate the thickness fluctuation signal, the obtained signal is divided by Q to the third and the excess dynamics are fitted to a Lorentzian function in Q.This method is contingent on good quality data, which is more achievable with samples of high concentrations and strong scattering signals. The dynamics probed by NSE can be synergistically explored by deuterium NMR relaxometry and molecular-dynamic simulations to illustrate how molecular lipid structures and packing motifs influence membrane functions.NSE studies on lipid membranes shed new light on membrane biophysics, the intricate relations of membrane structure and dynamics, and how they impact membrane functions and membrane-protein interactions.

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JoVE Journal

JoVE Core

Cell Biology

Biology

Unit 1

Membrane Structure and Components

SCIENTISTS IN ACTION

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. ...

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

Detection of Nitric Oxide and Superoxide Radical Anion by Electron Paramagnetic Resonance Spectroscopy from Cells using Spin Traps

Reactive nitrogen/oxygen species (ROS/RNS) at low concentrations play an important role in regulating cell function, signaling, and immune response but in unregulated concentrations are detrimental to cell viability1, 2. While living systems have evolved with endogenous and dietary antioxidant defense mechanisms to regulate ROS generation, ROS are produced continuously as natural by-products of normal metabolism of oxygen and can cause oxidative damage to biomolecules resulting in loss of protein function, DNA cleavage, or lipid peroxidation3, and ultimately to oxidative stress leading to cell injury or death4. 
Superoxide radical anion (O2&bull;-) is the major precursor of some of the most highly oxidizing species known to exist in biological systems such as peroxynitrite and hydroxyl radical. The generation of O2&bull;- signals the first sign of oxidative burst, and therefore, its detection and/or sequestration in biological systems is important. In this demonstration, O2&bull;- was generated from polymorphonuclear neutrophils (PMNs). Through chemotactic stimulation with phorbol-12-myristate-13-acetate (PMA), PMN generates O2&bull;- via activation of nicotinamide adenine dinucleotide phosphate (NADPH) oxidase5. 
Nitric oxide (NO) synthase which comes in three isoforms, as inducible-, neuronal- and endothelial-NOS, or iNOS, nNOS or eNOS, respectively, catalyzes the conversion of L- arginine to L-citrulline, using NADPH to produce NO6. Here, we generated NO from endothelial cells. Under oxidative stress conditions, eNOS for example can switch from producing NO to O2&bull;- in a process called uncoupling, which is believed to be caused by oxidation of heme7 or the co-factor, tetrahydrobiopterin (BH4)8.
There are only few reliable methods for the detection of free radicals in biological systems but are limited by specificity and sensitivity. Spin trapping is commonly used for the identification of free radicals and involves the addition reaction of a radical to a spin trap forming a persistent spin adduct which can be detected by electron paramagnetic resonance (EPR) spectroscopy. The various radical adducts exhibit distinctive spectrum which can be used to identify the radicals being generated and can provide a wealth of information about the nature and kinetics of radical production9.
The cyclic nitrones, 5,5-dimethyl-pyrroline-N-oxide, DMPO10, the phosphoryl-substituted DEPMPO11, and the ester-substituted, EMPO12 and BMPO13, have been widely employed as spin traps--the latter spin traps exhibiting longer half-lives for O2&bull;- adduct. Iron (II)-N-methyl-D-glucamine dithiocarbamate, Fe(MGD)2 is commonly used to trap NO due to high rate of adduct formation and the high stability of the spin adduct14.

Electron paramagnetic resonance (EPR) spectroscopy was employed to detect nitric oxide from bovine aortic endothelial cells and superoxide radical anion from human neutrophils using iron (II)-N-methyl-D-glucamine dithiocarbamate, Fe(MGD)2 and 5,5-dimethyl-1-pyroroline-N-oxide, DMPO, respectively.

Reactive nitrogen/oxygen species (ROS/RNS) at low concentrations play an important role in regulating cell function, signaling, and immune response but in ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Imaging FITC-dextran as a Reporter for Regulated Exocytosis

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated exocytosis is fundamental for intercellular communication and is a key mechanism for the secretion of neurotransmitters, hormones, inflammatory mediators, and other compounds, by a variety of cells. At least three distinct mechanisms are known for regulated exocytosis: full exocytosis, where a single SG fully fuses with the plasma membrane, kiss-and-run exocytosis, where a single SG transiently fuses with the plasma membrane, and compound exocytosis, where several SGs fuse with each other, prior to or after SG fusion with the plasma membrane. The type of regulated exocytosis undertaken by a cell is often dictated by the type of secretory trigger. However, in many cells, a single secretory trigger can activate multiple modes of regulated exocytosis simultaneously. Despite their abundance and importance across cell types and species, the mechanisms that determine the different modes of secretion are largely unresolved. One of the main challenges in investigating the different modes of regulated exocytosis, is the difficulty in distinguishing between them as well as exploring them separately. Here we describe the use of fluorescein isothiocyanate (FITC)-dextran as an exocytosis reporter, and live cell imaging, to differentiate between the different pathways of regulated exocytosis, focusing on compound exocytosis, based on the robustness and duration of the exocytic events.

Here we detail a method for live cell imaging of regulated exocytosis. This method utilizes FITC-dextran, which accumulates in lysosome-related organelles, as a reporter. This simple method also allows distinguishing between different modes of regulated exocytosis in cells that are difficult to manipulate genetically.

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Detection of Protein Aggregation using Fluorescence Correlation Spectroscopy

Protein aggregation is a hallmark of neurodegenerative disorders such as amyotrophic lateral sclerosis (ALS), Alzheimer's disease (AD), Parkinson's disease (PD), Huntington's disease (HD), and so on. To detect and analyze soluble or diffuse protein oligomers or aggregates, fluorescence correlation spectroscopy (FCS), which can detect the diffusion speed and brightness of a single particle with a single molecule sensitivity, has been used. However, the proper procedure and know-how for protein aggregation detection have not been widely shared. Here, we show a standard procedure of FCS measurement for diffusion properties of aggregation-prone proteins in cell lysate and live cells: ALS-associated 25 kDa carboxyl-terminal fragment of TAR DNA/RNA-binding protein 43 kDa (TDP25) and superoxide dismutase 1 (SOD1). The representative results show that a part of aggregates of green fluorescent protein (GFP)-tagged TDP25 was slightly included in the soluble fraction of murine neuroblastoma Neuro2a cell lysate. Moreover, GFP-tagged SOD1 carrying ALS-associated mutation shows a slower diffusion in live cells. Accordingly, we here introduce the procedure to detect the protein aggregation via its diffusion property using FCS.

We here introduce a procedure to measure protein oligomers and aggregation in cell lysate and live cells using fluorescence correlation spectroscopy.

Protein aggregation is a hallmark of neurodegenerative disorders such as amyotrophic lateral sclerosis (ALS), Alzheimer's disease (AD), Parkinson's ...