We thank Beate Nitschke for her help with protein preparation and Robin Ghosh, Michael Schweikert (Stuttgart), and Maximilan Ulbrich (Freiburg) for insightful discussions. This work was supported by the Stuttgart Research Center Systems Biology (SRCSB) and a grant from the Baden W&#252;rttemberg Foundation (BiofMO-6 to SN).

We declare no conflicts of interest.

The protocol presented here provides an introduction to the use of DIB membranes to study the interplay between lateral ion channel movement and channel function using single-molecule TIRF microscopy. To obtain the best possible data, the preparation of stable DIB membranes with as many well-separated channels as possible is crucial for obtaining time series of individual particles, which can be analyzed satisfactorily.
Critical parameters to be optimized include the choice of lipid, the lipid concentration in the oil phase, and the protein and detergent concentrations in the aqueous droplets. The lipids employed are unusual, in that they show no clear phase transition at low temperatures. DPhPC is a commonly used lipid to produce stable membrane systems40. In principle, any lipid which maintains its fluid environment at low temperatures may be suitable for this application. In addition, the lipid should not be sensitive to oxidation. The detergent concentration in the droplets should be as low as possible to avoid membrane rupture. Stable membranes and good protein incorporation rates are generally achieved with detergent concentrations below the critical micelle concentration (cmc), given that the membrane protein does not precipitate.
If the DIB membranes do not tolerate specific detergents21,41, or if the proteins do not integrate from low detergent solution into the DIB membranes, the protein channels can first be reconstituted into small unilamellar lipid vesicles (SUVs), which are then fused to the DIB membranes from the droplet side, as has been successfully shown for E. coli MscL42. Sometimes, DIB membranes do not form because the lipid concentration in the oil phase is too low. To prevent DIB membranes from bursting, one should also be aware that the osmotic pressure between the hydrogel and the droplet must be precisely balanced without affecting the Ca2+-flux from cis to trans excessively. Optimized agarose thickness and mesh size seem to be crucial to observe the diffusion of membrane proteins. Any drying of the agarose layer should be avoided. The thickness can be determined using atomic force microscopy17. By varying the agarose concentration, volume, and rotation speed during spin coating, the mesh size and thickness of the hydrogel can be optimized. Note, however, that hydrogel layer thickness affects image contrast. To capture membrane proteins in DIBs, the agarose hydrogel can be replaced by custom-synthesized, non-crosslinked, Ni-NTA-modified, low-melting agarose to trap them via a His-tag17. An excessively high fluorescence background is often caused by rupture of the DIB membranes. This is particularly a problem with multi-well chambers, as the Ca2+-sensitive dye diffuses into the hydrogel. In this case, adjacent wells should be avoided. Fluorescence bleaching of the Ca2+-sensitive dye above the membrane should not be a significant limiting factor, as it is exchanged by unexcited dyes in the bulk of the droplet (Figure 3A) outside the TIRF evanescent field. The localization precision for the protein is given by the accuracy of fitting the spots and the pixel size.
Weak fluorescence signals can be caused by low Ca2+-flux through the channel. Possible reasons include: (i) inaccurate TIRF settings (e.g., laser intensity), (ii) the osmotic Ca2+ pressure across the membrane, or (iii) the intrinsic Ca2+-permeability of the channels is too low. To cope with the first issue, laser intensity, TIRF angle, and camera gain need to be optimized. The latter two issues can be overcome by the application of an electrical potential across the membrane2,43. However, the application of external voltages can distort the result, as electrical effects can influence the channel opening of ligand-gated or mechanosensitive ion channels that are actually not voltage-controlled. Examples of such channels are the mitochondrial protein translocase TOM-CC27, and its channel-forming subunit Tom4026,44,45,46. Finally, it should be noted that, inserting membrane proteins into DIB membranes in a specific orientation to achieve a desired functionality is tricky, and quantitative studies are rare47,48. In some cases, the orientation of the integrated proteins is random. This is a serious problem for studying membrane proteins, because certain membrane proteins are activated on only one side of the membrane.
TIRF microscopy is a powerful method for addressing single-molecule events in planar supported membranes49. Examples include assembly and folding pathway elucidation of channel proteins such as &#945;-hemolysin50, perfringolysin O51, and OmpG52. These studies included FRET as an additional technique. In addition, activation of the mechanosensitive ion channel MscL has previously been studied by mechanical stimulation of supported DIB bilayers42 using current measurements. Based on this work, future studies could combine the platform described here with single-molecule FRET experiments to address mechanosensitive channels at the single-molecule level in an optical manner17. Injection of buffer into the droplet, stretching the inner DIB monolayer, or targeted binding of individual channels to the underlying hydrogel can be used to further study not only the physical mechanism of mechanically activated channels, which respond to membrane tension and/or curvature as shown for MscL and MscS, the two-pore domain K+-channels, TREK-1, TREK-2, and TRAAK, and PIEZO (for review, see53), but also local binding to the cellular cytoskeleton, as shown for the touch-sensitive ion channel NOMPC54,55.

The present protocol aims to describe how to study the correlation between membrane mobility and ion channel permeability of membrane proteins in polymer-supported droplet-interface bilayer (DIB) membranes1,2,3.
The present technique complements an impressive array of advanced optical and surface analytical tools, such as single particle tracking4,5, fluorescence correlation spectroscopy6,7, and high-speed atomic force microscopy8,9,10. These provide valuable insights into the dynamic composition and structure of membranes that influence membrane-based reactions11,12,13. While the movement and lateral diffusion of proteins depends on the local density of proteins in the membrane, individual protein molecules can also be trapped in lipid rafts14 and by protein-protein interactions15,16. Depending on the protein domains protruding from the membrane into the extracellular environment or cytosol, protein mobility can vary from highly mobile to completely immobile. However, due to the complexity of the membrane and its peripheral structures, it is often difficult to decipher the interplay between the nature of lateral mobility and protein function17.
DIB membranes have proven to be an efficient platform for biophysical single-molecule analyses of membrane proteins18,19,20,21,22. They are formed by lipid self-assembly through the contact of aqueous droplets with hydrogel-supported substrates in a lipid/oil phase. Similar to the commonly used supported lipid bilayers (SLBs)1,23,24,25, DIBs allow local tuning of the lateral mobility by temporary or permanent binding of proteins to the polymer matrix when functionalized with suitable ligands17. The latter can serve as a model system for biochemical processes in cell membranes with a heterogeneous protein distribution10.
The experimental approach described here relies on the fluorescence of Ca2+-sensitive dyes to measure the Ca2+ ion flux through individual channels in close proximity to the membrane2,22 using TIRF microscopy. This optical approach limits the illumination of the sample to a distance close to the membrane, which, due to the physical properties of the evanescent excitation light, leads to a significant reduction of the fluorescence background. The latter is a prerequisite if high spatial and temporal resolution is required for the detection of single molecules. In contrast to classical electrophysiological methods26,27, no membrane voltages are required to study ion flux through individual channels. Furthermore, the method does not require labeling with fluorescent dyes or molecules that could interfere with the lateral movement of the channels in the membrane.
The method is particularly useful for studying protein channels embedded in the membrane at the single molecule level without using classical electrophysiology. Using mitochondrial protein-conducting channel TOM-CC from Neurospora crassa28,29,30 and OmpF, which supports the diffusion of small hydrophilic molecules across the outer membrane of Escherichia coli17,31, we illustrate how the membrane mobilities and channel activities of the two proteins can be studied and correlated. We suggest that this approach, although optimized for TOM-CC and OmpF, can be readily applied to other protein channels.

<table><tbody><tr><td>1,2-diphytanoyl-sn-glycero-3-phosphocholine</td><td>Avanti Polar Lipids</td><td>850356C</td><td></td></tr><tr><td>100x Oil objective Apochromat N.A. 1.49</td><td>Nikon</td><td>MRD01991</td><td>TIRF microscope&amp;nbsp;</td></tr><tr><td>10x Hoffmann modulation contrast objective NA 0.25&amp;nbsp;</td><td>Nikon</td><td></td><td>Microscope to assess DIB membrane formation&amp;nbsp;&amp;nbsp;</td></tr><tr><td>10x objective N.A. 0.25</td><td>Nikon</td><td>MRL00102</td><td>TIRF microscope&amp;nbsp;</td></tr><tr><td>40x Hoffmann modulation contrast objective NA 0.55</td><td>Nikon</td><td></td><td>Microscope to assess DIB membrane formation&amp;nbsp;&amp;nbsp;</td></tr><tr><td>488 nm laser, 100 mW&amp;nbsp;&amp;nbsp;</td><td>Visitron</td><td></td><td>TIRF microscope&amp;nbsp;</td></tr><tr><td>Adhesive tape</td><td></td><td></td><td></td></tr><tr><td>&amp;Auml;kta pure&amp;nbsp;&amp;nbsp;</td><td>Cytiva</td><td></td><td>Protein purification system</td></tr><tr><td>Bradford assay kit, Pierce</td><td>Thermo Fisher&amp;nbsp;</td><td>23236</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Roth</td><td>5239.2</td><td></td></tr><tr><td>Chelax 100 resin</td><td>Biorad</td><td>143-2832</td><td></td></tr><tr><td>Chloroform&amp;nbsp;</td><td>Sigma-Aldrich</td><td>MC1024452500</td><td></td></tr><tr><td>Dialysis cassettes Slide-A-Lyzer 20 k MWCO</td><td>Thermo Fisher&amp;nbsp;</td><td>87735</td><td></td></tr><tr><td>DIB chamber&amp;nbsp;</td><td>Custom made</td><td></td><td>PMMA chamber for DIB membranes</td></tr><tr><td>Digital power meter and energy console</td><td>Thorlabs</td><td>PM100D</td><td>Laser power meter&amp;nbsp;</td></tr><tr><td>Dimethyl sulfoxide</td><td>Roth&amp;nbsp;</td><td>4720.1</td><td></td></tr><tr><td>Double distilled H&lt;sub&gt;2&lt;/sub&gt;O</td><td></td><td></td><td></td></tr><tr><td>Eclipse TS 100 Hoffmann modulation contrast microscope&amp;nbsp;</td><td>Nikon</td><td></td><td>Microscope to assess DIB membrane formation&amp;nbsp;&amp;nbsp;</td></tr><tr><td>EDTA</td><td>Roth</td><td>8042.2</td><td></td></tr><tr><td>EMCCD camera iXon Ultra 897</td><td>Andor</td><td></td><td>TIRF microscope&amp;nbsp;</td></tr><tr><td>Ethanol</td><td>Sigma-Aldrich</td><td>32205-M</td><td></td></tr><tr><td>Fixed angle rotor Ti70</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Fluo-8, CalciFluor&lt;sup&gt;TM&lt;/sup&gt;</td><td>Santa Cruz Biotechnology</td><td>SC-362561</td><td>Ca&lt;sup&gt;2+&lt;/sup&gt;-sensitive dye</td></tr><tr><td>French press cell disruption homogenizer</td><td>Igneus</td><td>Igneus 40000 psi</td><td></td></tr><tr><td>GFP filter</td><td>AHF</td><td></td><td>Filter seeting used for excitation of DIB-membranes by epifluorescence with white light source&amp;nbsp;</td></tr><tr><td>Glass capillaries</td><td>World Precision Instruments</td><td>4878</td><td></td></tr><tr><td>Glass coverslips 40 mm x 24 mm x 0.13 mm</td><td>Roth</td><td>1870.2</td><td></td></tr><tr><td>Glycerol</td><td>Roth</td><td>3783.2</td><td></td></tr><tr><td>Hamilton syringe 10 mL</td><td>Roth</td><td>X033.1</td><td></td></tr><tr><td>Hamilton syringe 100 mL</td><td>Roth</td><td>X049.1</td><td></td></tr><tr><td>Hamilton syringe 500 mL</td><td>Roth</td><td>EY49.1</td><td></td></tr><tr><td>Heating block&amp;nbsp;</td><td>Eppendorf</td><td>Thermomixer comfort</td><td></td></tr><tr><td>Heating plate</td><td>Minitube&amp;nbsp;</td><td>HT200</td><td></td></tr><tr><td>Hepes</td><td>Roth&amp;nbsp;</td><td>9205.3</td><td></td></tr><tr><td>Hexadcane</td><td>Sigma-Aldrich</td><td>296317</td><td></td></tr><tr><td>His Trap HP 1 mL</td><td>Cytiva</td><td>29051021</td><td>Ni-NTA column</td></tr><tr><td>Imidazole</td><td>Sigma-Aldrich</td><td>1.04716.1000</td><td></td></tr><tr><td>KCl</td><td>Honeywell</td><td>10314243</td><td></td></tr><tr><td>KLM spin coater&amp;nbsp;</td><td>Schaefer Tec</td><td>SCV-10</td><td></td></tr><tr><td>List medical L/M-3P-A vertical pipette puller</td><td>Artisan Technology Group</td><td>57761-1</td><td></td></tr><tr><td>Low melting point agarose</td><td>Sigma-Aldrich&amp;nbsp;</td><td>A9414</td><td></td></tr><tr><td>M8 Stereomicroscope</td><td>Wild</td><td></td><td>Stereomicrosope&amp;nbsp;</td></tr><tr><td>Matlab&amp;nbsp;</td><td>MathWorks</td><td>R2022a</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>34860</td><td></td></tr><tr><td>MicroFil pipette tips</td><td>World Precision Instruments</td><td>MF34G-5</td><td></td></tr><tr><td>N&lt;sub&gt;2&lt;/sub&gt; gas</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td>Roth</td><td>3957.1</td><td></td></tr><tr><td>Nanoliter 2010 injector&amp;nbsp;</td><td>World Precision Instruments</td><td>Nanoliter 2010&amp;nbsp;</td><td></td></tr><tr><td>n-dodecyl-b-D-maltoside</td><td>Glycon Biochemicals</td><td>D97002-C</td><td></td></tr><tr><td>Ni-NTA agarose, non-crosslinked</td><td>Cube Biotech</td><td>124115393</td><td>Custom made</td></tr><tr><td>NIS-Elements AR software</td><td>Nikon</td><td>MQS31100/MQS42560/MQS42580/MQS42780/MQS41930</td><td>Imaging software</td></tr><tr><td>n-octyl-polyoxyethylene</td><td>Sigma-Aldrich</td><td>40530</td><td></td></tr><tr><td>O&lt;sub&gt;2&lt;/sub&gt; gas</td><td></td><td></td><td></td></tr><tr><td>Phenylmethylsulfonyl fluoride</td><td>Roth&amp;nbsp;</td><td>6367.3</td><td></td></tr><tr><td>Photodiode sensor&amp;nbsp; Si, 400 - 1100 nm, 500 mW</td><td>Thorlabs</td><td>S130C</td><td>Sensor for laser power meter&amp;nbsp;</td></tr><tr><td>Plasma cleaner</td><td>Diener Electronics</td><td>Zepto</td><td></td></tr><tr><td>Preparative ultracentrifuge Optima</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Quad-band TIRF-filter 446/523/600/677 HC</td><td>AHF</td><td></td><td>Filter setting used for excitation of DIB-membranes with 488 nm laser&amp;nbsp;</td></tr><tr><td>Resource Q 1 mL</td><td>Cytiva</td><td>17117701</td><td>Anion exchange column</td></tr><tr><td>Silicon oil AR 20</td><td>Sigma-Aldrich</td><td>10836</td><td></td></tr><tr><td>Sodium dodecyl sulfate</td><td>Roth</td><td>2326.2</td><td></td></tr><tr><td>Super LoLux camera</td><td>JVC</td><td></td><td>Stereomicrosope&amp;nbsp;</td></tr><tr><td>Thermoshaker</td><td>Gerhardt</td><td>THL 500/1</td><td></td></tr><tr><td>Ti-E Fluorescence microscope&amp;nbsp;&amp;nbsp;</td><td>Nikon</td><td>MEA53100</td><td></td></tr><tr><td>Tris-HCl</td><td>Sigma-Aldrich</td><td>9090.3</td><td></td></tr><tr><td>Tryptone</td><td>Roth</td><td>8952.2</td><td></td></tr><tr><td>Ultrasonic bath&amp;nbsp;</td><td>Bandelin Sonorex</td><td>RK 100</td><td></td></tr><tr><td>Vaccum pump</td><td>Vacuubrand</td><td>MD 4C NT</td><td></td></tr><tr><td>White light source for epifluorescence illumination (100 W)</td><td>Nikon</td><td>MBF72655</td><td>TIRF microscope&amp;nbsp;</td></tr><tr><td>Yeast extract</td><td>Roth</td><td>2363.2</td><td></td></tr><tr><td>&amp;beta;-mercaptoethanol</td><td>Sigma-Aldrich</td><td>M3148</td><td></td></tr></tbody></table>

single-molecule imaging of lateral mobility and ion channel activity in lipid bilayers using total internal reflection fluorescence (tirf) microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Real-time electrode-free optical single-channel recording reveals the interplay between lateral protein movement and the function of individual ion channels in DIB membranes. Reconstitution of the mitochondrial TOM core complex (Figure 1A) into a DIB membrane (Figure 4D) illustrates a strong temporal correlation between lateral mobility and ion permeability (Figure 5A). The TOM-CC gating appears to be sensitive to the mode of lateral movement17. Moving channels show Ca2+ flux through their pores and high fluorescence point intensities. Trapped, non-moving molecules show low and medium fluorescence intensities. For the general protein import pore of mitochondria TOM-CC, this single molecule approach revealed a strong temporal correlation between lateral mobility and ion permeability, suggesting that the TOM-CC channel has mechanosensitive properties17. A lateral stop of freely moving TOM-CC molecules is accompanied by a partial or complete closure of the TOM-CC channel. Imaging DIB membranes with OmpF (Figure 1E and Figure 4F), which is almost fully embedded in the membrane, shows no stop-and-go effects (Figure 5B). Random stops of OmpF are not associated with a change in intensity and, thus, with the closing of its pores. Based on the fluorescence signals38, the positional accuracy of the individual channels can be estimated in the range between 5 and 10 nm. However, it should be noted that this accuracy cannot be achieved if the channels wobble slightly due to mobile anchoring with the agarose hydrogel, as shown, for example, for the TOM-CC molecules in an intermediate state with a root mean displacement of 120 nm (Figure 5A).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64970/64970fig01.jpg" /> 
Figure 1: Isolation of TOM-CC. (A) Cryo EM structure of N. crassa TOM-CC30,39. Mitochondria from a N. crassa strain containing a Tom22 with a 6xHis tag were solubilized in DDM and subjected to Ni-NTA affinity chromatography (B) and anion-exchange chromatography (C). (D) SDS-PAGE of isolated TOM-CC. (E) Crystal structure (PDB, 1OPF) and (F) SDS-PAGE of purified E. coli OmpF. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64970/64970fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64970/64970fig02.jpg" /> 
Figure 2: Flow chart of PMMA chamber assembly. Step 1: A glass coverslip is spin-coated with an agarose hydrogel. Step 2: The spin-coated coverslip is mounted in a custom-made PMMA microscopy chamber. Step 3: Additional low melt agarose is added into the inlet port of the PMMA chamber on a 35 &#176;C hot plate. Step 4: Lipid monolayers are formed around aqueous droplets at a buffer/oil interface (left) and on the agarose hydrogel/oil interface (right). Step 5: Individual aqueous droplets are pipetted into the wells of the PMMA chamber to form a lipid bilayer upon contact of the two lipid monolayers. Step 6: Formation of the DIB membranes is validated by Hofmann modulation contrast microscopy. Step 7: Images of selected areas of the DIB membranes with inserted ion channels are acquired by TIRF microscopy. Green: 0.75% agarose; yellow: 2.5% agarose containing Ca2+ ions; magenta: lipid/oil phase; dark blue: aqueous droplet buffer containing Ca2+-sensitive dye and protein. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64970/64970fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64970/64970fig03.jpg" /> 
Figure 3: Experimental setup. (A) Schematic representation of a DIB membrane in a PMMA well. The bilayer rests on an ultrathin 0.75% agarose film to allow TIRF imaging of Ca2+-flux through an ion channel over time using a Ca2+-sensitive fluorescence dye (Fluo-8) in trans. (B) The Ca2+-flux is controlled exclusively by the osmotic pressure from cis to trans. This allows both the determination of the position in the membrane and the open-closed state of the channel. The channel shown here is the protein-conducting channel TOM-CC of N. crassa mitochondria30. (C) Trajectory of a single TOM-CC channel. Green: moving channel; yellow: non-moving channel. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64970/64970fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64970/64970fig04.jpg" /> 
Figure 4: Imaging TOM-CC and OmpF in DIB membranes. (A) DIB membrane imaged by Hoffmann modulation contrast microscopy showing the bilayer contact area between the hydrogel and the droplet. (B) Broken DIB membrane imaged as in (A). Arrow, edge of PMMA chamber. (C) DIB membrane imaged by TIRF microscopy without protein channels. (D) DIB membrane with reconstituted TOM-CC, imaged by TIRF microscopy. The white squares mark spots of high (SH), intermediate (SI), and low (SL) intensity. (E) Fitting the fluorescence intensity profile of the three spots marked in (A) to two-dimensional Gaussian functions reveals the position of individual TOM-CCs and the Ca2+ flux through the channel. (F) DIB membrane with reconstituted OmpF. (G) Gaussian fit of the fluorescent spot marked in (F). In contrast to the two-pore &#946;-barrel protein complex TOM-CC, the three-pore &#946;-barrel OmpF reveals only one permeability state. Pixel size, 0.16 &#956;m.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64970/64970fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64970/64970fig05.jpg" /> 
Figure 5: Channel activity correlates with lateral mobility of TOM-CC. (A) The fluorescent amplitude trace (top) and corresponding trajectory (bottom) of TOM-CC indicate that the open-closed channel activity of TOM-CC correlates with the lateral membrane mobility of the complex. The trajectory displays three permeability states. Green: fully open state; yellow: intermediate permeability state; red: closed channel state; red star: TOM-CC in the intermediate state wobbles around its mean position by about &#177;60 nm. The positional accuracies37 based on the fluorescence signals in the fully open and intermediate states range between 5 nm and 10 nm. (B) Fluorescent amplitude trace (top) and corresponding trajectory (bottom) of OmpF. OmpF reveals only one intensity level, regardless of whether it is in motion or trapped. The trajectory segments corresponding to the time periods of trapped molecules are marked in grey. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64970/64970fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Buffer&#160;</td>
			<td colspan="3">Reagent concentrations</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>A1*</td>
			<td colspan="3">20 mM Tris-HCl pH 8.5, 0.1 % (w/v) n-dodecyl-&#946;-D-maltoside (DDM), 10% (v/v) glycerol, 300 mM NaCl, and 1 mM phenylmethylsulfonyl fluoride (PMSF)</td>
			<td>100 mL</td>
		</tr>
		<tr>
			<td>A2*</td>
			<td colspan="3">20 mM Tris-HCl pH 8.5, 0.1% (w/v) DDM, 10% (v/v) glycerol, 1 M Imidazole and 1 mM PMSF</td>
			<td>100 mL</td>
		</tr>
		<tr>
			<td>B1*</td>
			<td colspan="3">20&#8201;mM HEPES pH 7.2, 0.1% (w/v) DDM, 2% (v/v) dimethyl sulfoxide (DMSO)</td>
			<td>100 mL&#160;</td>
		</tr>
		<tr>
			<td>B2*</td>
			<td colspan="3">20&#8201;mM HEPES pH 7.2, 0.1% (w/v) DDM, 1 M KCl, 2% (v/v) dimethyl sulfoxide (DMSO)</td>
			<td>100 mL</td>
		</tr>
		<tr>
			<td colspan="5">* Degas and pass through a 0.22 &#956;m filter before use.</td>
		</tr>
	</tbody>
</table>
Table 1: Buffer solutions for TOM-CC isolation.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Buffer&#160;</td>
			<td colspan="3">Reagent concentrations</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>LB*</td>
			<td colspan="3">1% (w/v) tryptone, 1% (w/v) NaCl and 0.5% (w/v) yeast extract&#160;</td>
			<td>1100 mL</td>
		</tr>
		<tr>
			<td>C1</td>
			<td colspan="3">2 mM MgCl2, and ~750 units DNAse and 50 mM Tris-HCl pH 7.5&#160;&#160;</td>
			<td>20 mL</td>
		</tr>
		<tr>
			<td>C2</td>
			<td colspan="3">50 mM Tris-HCl, pH 7.5</td>
			<td>50 mL&#160;</td>
		</tr>
		<tr>
			<td>C3</td>
			<td colspan="3">4% (w/v) sodium dodecyl sulfate (SDS), 2 mM &#946;-mercaptoethanol and 50 mM Tris-HCl pH 7.5</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>C4</td>
			<td colspan="3">2% (w/v) SDS, 500 mM NaCl and 50 mM Tris-HCl pH 7.5</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>C5</td>
			<td colspan="3">0.5% (w/v) octyl polyoxyethylene (octyl POE), 1 mM EDTA and 20 mM Tris pH 8.5&#160;</td>
			<td>1000 mL</td>
		</tr>
		<tr>
			<td colspan="5">* Sterilize before use.</td>
		</tr>
	</tbody>
</table>
Table 2: Buffer solutions for OmpF isolation.

Watch this Scientific Journal Video about Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy at JoVE.co

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...

single-molecule-imaging-lateral-mobility-ion-channel-activity-lipid

Nanyang Technological University

Research

JoVE Journal

Biochemistry

2.9K Views.  University of Stuttgart. Ionkanalen zijn niet statisch in biologische membranen. Hier presenteren we een optische benadering met één molecuul om het verband tussen laterale membraandiffusie en ionkanaalfunctie te ontrafelen. Het belangrijkste voordeel van deze techniek ten opzichte van andere methoden is dat fluorescerende etikettering van eiwitten die hun laterale beweging en functie kunnen verstoren, niet vereist is.De methode kan worden toegepast op elk membraaneiwitkanaal waar vrije of beperkte diffusie ...

Video: Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

A TIRF Microscopy Technique for Real-time, Simultaneous Imaging of the TCR and its Associated Signaling Proteins

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex (pMHC) proteins expressed on the surface of antigen presenting cells (APCs). The TCR complex is composed of the ligand binding TCR&alpha;&beta; heterodimer that associates non-covalently with CD3 dimers (the &#949;&delta; and &#949;&gamma; heterodimers and the &zeta;&zeta; homodimer)1. Upon engagement of the receptor, the CD3 &zeta; chains are phosphorylated by the Src family kinase, Lck. This leads to the recruitment of the Syk family kinase, Zap70, which is then phosphorylated and activated by Lck. After that, Zap70 phosphorylates the adapter proteins LAT and SLP76, initiating the formation of the proximal signaling complex containing a large number of different signaling molecules2. 
The formation of this complex eventually results in calcium and Ras-dependent transcription factor activation and the consequent initiation of a complex series of gene expression programs that give rise to T cell differentiation2. TCR signals (and the resulting state of differentiation) are modulated by many other factors, including antigen potency and crosstalk with co-stimulatory/co-inhibitory, chemokine, and cytokine receptors 3-4. Studying the spatial and temporal organization of the proximal signaling complex under various stimulation conditions is, therefore, key to understanding the TCR signaling pathway as well as its regulation by other signaling pathways.
One very useful model system to study signaling initiated by the TCR at the plasma membrane in T cells is glass-supported lipid bilayers, as described previously5-6. They can be utilized to present antigenic pMHC complexes, adhesion, and co-stimulatory molecules to T cells-serving as artificial APCs. By imaging the T cells interacting with the lipid bilayer using total internal reflection fluorescence microscopy (TIRFM), we can restrict the excitation to within 100 nm of the space between the glass and the cell surface 7-8. This allows us to image primarily the signaling events occurring at the plasma membrane. As we are interested in imaging the recruitment of signaling proteins to the TCR complex, we describe a two-camera TIRF imaging system wherein the TCR, labeled with fluorescent Fab (fragment antigen binding) fragments of the H57 antibody (purified from hybridoma H57-597, ATCC, ATCC Number:HB-218) which is specific for TCR&beta;, and signaling proteins, tagged with GFP, may be imaged simultaneously and in real time. This strategy is necessary due to the highly dynamic nature of both the T cells and of the signaling events that are occurring at the TCR. This imaging modality has allowed researchers to image single ligands 9-11 as well as recruitment of signaling molecules to activated receptors and is an excellent system to study biochemistry in-situ12-16.

The compartmentalization of proteins either within the plasma membrane or into intracellular locations is one regulatory mechanism that can greatly influence signaling outcomes; hence, to understand signaling it is important to study the spatial and temporal behavior of the proteins involved. We describe here a TIRF microscopy based system to study signal transduction in T cells, but is broadly applicable.

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex ...

Immunology and Infection

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Bioengineering

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

Imaging Plasma Membrane Deformations With pTIRFM

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during the fusion of vesicle and plasma membranes. These rapid and localized changes are achieved by dynamic interactions between lipids and specialized proteins that control membrane curvature. The absence of such interactions would not only have devastating consequences for vesicle fusion, but a host of other cellular functions that involve control of membrane shape. In recent years, the identity of a number of proteins with membrane-shaping properties has been determined. What remains missing is a roadmap of when, where, and how they act as fusion and content release progress.
Our understanding of the molecular events that enable membrane remodeling has historically been limited by a lack of analytical methods that are sensitive to membrane curvature or have the temporal resolution to track rapid changes. PTIRFM satisfies both of these criteria. We discuss how pTIRFM is implemented to visualize and interpret rapid, submicron changes in the orientation of chromaffin cell membranes during dense core vesicle (DCV) fusion. The chromaffin cells we use are isolated from bovine adrenal glands. The membrane is stained with a lipophilic carbocyanine dye,1,1&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or&#160;diD. DiD intercalates in the membrane plane with a &#34;fixed&#34; orientation and is therefore sensitive to the polarization of the evanescent field. The diD-stained cell membrane is sequentially excited with orthogonal polarizations of a 561&#160;nm laser (p-pol, s-pol). A 488&#160;nm laser is used to visualize vesicle constituents and time the moment of fusion. Exocytosis is triggered by locally perfusing cells with a depolarizing KCl solution. Analysis is performed offline using custom-written software to understand how diD emission intensity changes relate to fusion pore dilation.

Polarization-based Total Internal Reflection Fluorescence Microscopy (pTIRFM) enables real-time detection of cell membrane dynamics. This article describes the implementation of pTIRFM for the study of membrane remodeling during regulated exocytosis. The technique is generalizable to other processes in cell biology that directly or indirectly involve changes in membrane shape.

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during ...

Biology

Visualizing Clathrin-mediated Endocytosis of G Protein-coupled Receptors at Single-event Resolution via TIRF Microscopy

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological assays, measuring global changes in endocytosis, have identified over 30 known components participating in CME, and biochemical studies have generated an interaction map of many of these components. It is becoming increasingly clear, however, that CME is a highly dynamic process whose regulation is complex and delicate. In this manuscript, we describe the use of Total Internal Reflection Fluorescence (TIRF) microscopy to directly visualize the dynamics of components of the clathrin-mediated endocytic machinery, in real time in living cells, at the level of individual events that mediate this process. This approach is essential to elucidate the subtle changes that can alter endocytosis without globally blocking it, as is seen with physiological regulation. We will focus on using this technique to analyze an area of emerging interest, the role of cargo composition in modulating the dynamics of distinct clathrin-coated pits (CCPs). This protocol is compatible with a variety of widely available fluorescence probes, and may be applied to visualizing the dynamics of many cargo molecules that are internalized from the cell surface.

Clathrin-mediated endocytosis, a rapid and highly dynamic process internalizes many proteins, including signaling receptors. The protocol described here directly visualizes the kinetics of individual endocytic events. This is essential for understanding how core members of the endocytic machinery coordinate with each other, and how protein cargo influence this process.

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological ...

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

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