The authors acknowledge Prof. Benjamin Schuler for sharing the expression plasmid for ClyA protein. This work was supported by Human Frontier Science Program (RGP0047-2020).

1. Cleaning of the slide and coverslip for single-molecule experiments
<ol>
	<li>Before the assembly of the imaging chamber, clean and prepare both the coverslips and slides. Drill multiple pairs of holes on the glass slides using a drilling machine with diamond-coated drill bits (0.5-1 mm in diameter). If acrylic sheets are used, use a laser cutter to make precise holes (0.5 mm), as shown in Figure 1. 
	NOTE: Each pair of holes will act as an inlet and outlet for flow exchange for an individual microfluidic chamber. A representative CAD file for holes in an acrylic slide is available in Supplementary Coding File 1. Holes drilled on glass slides usually end up being 1.5x-2x larger than the drill bit size.</li>
	<li>Clean the slides and coverslips with detergent. Rinse them thoroughly with deionized water. Place the slides and coverslips in a separate slide staining (Coplin) jar with water and sonicate in a bath sonicator for 30 min. For all the sonication steps, use a 70 W power setting.</li>
	<li>Remove the water and fill the Coplin jar containing the glass slides and coverslips with acetone to the brim (50-100 mL depending on the jar volume). In the case of acrylic sheets, use the same volume of premixed 50% (v/v) acetone solution or else the slides will become frosty. Shake the Coplin jar to ensure all the slides and coverslips are completely immersed in the solution and sonicate in a bath sonicator for 30 min.</li>
	<li>Remove the acetone and discard it in a waste container, pour methanol into the Coplin jars (50%, v/v for acrylic slides), and sonicate for 30 min. Both acetone and methanol are used to remove organic particles on the slides.</li>
	<li>Rinse the slides and coverslips with copious amounts of type 1 water and place them in a glass Coplin jar. Pour 1 M KOH solution into the jar and sonicate for 1 h. For the acrylic slides, use 0.5 M of KOH.</li>
	<li>Discard the KOH in an inorganic waste container. Rinse the jar with plenty of water and place the Coplin jars in a fume hood for piranha treatment. Do not perform piranha treatment for acrylic slides; directly proceed to step 1.9. 
	CAUTION: Piranha treatment should be done in a fume hood with proper gloves, safety glasses, and an apron for handling acids. Piranha solution is a strong oxidizing agent, and it removes any leftover organic particles from the slides and coverslips.</li>
	<li>For preparing the piranha solution, gently pour a 2/3 volume of sulphuric acid (98%, v/v) into a glass beaker and fill the rest with hydrogen peroxide (30%, v/v). Mix the solution carefully with a glass rod, pour it into the Coplin jar, and leave the Coplin jar in the fume hood for at least 1 h.</li>
	<li>Decant the piranha solution from the Coplin jar in a discard container for acid waste kept inside the fume hood. Rinse the Coplin jar with a copious amount of type 1 water.</li>
	<li>Dry the slides and coverslips in a gentle stream of nitrogen gas and place them in a clean Coplin jar. The dried slides and coverslips are now ready for plasma treatment. Take a pair of slides and coverslips and place them in the plasma machine.</li>
	<li>Create a vacuum in the chamber. Turn the knob to the radio frequency of 8-12 MHz for generating plasma in the chamber. A purple glow inside the chamber will indicate the formation of a plasma in the chamber.</li>
	<li>Perform the plasma treatment for 8-10 min. Gently release the vacuum after turning off the plasma and proceed to step 2 for assembling the microfluidic imaging chamber.</li>
</ol>
2. Assembly of the microfluidic chamber
NOTE: The imaging chamber is created by sandwiching double-sided tape between a pre-cleaned coverslip and slide from the previous step&#160;as described below.
<ol>
	<li>Assemble the slides and coverslip after the plasma treatment as shown in Figure 1. Place the glass slide on a clean tissue paper. Cut the double-sided tape into ~2-3 mm wide strips and place them on each side of a pair of holes on the glass slide. Use a pipet tip to flatten out the tape or it will cause leakage from one channel to the other. 
	NOTE: Alternatively, cut the tape for the entire slide using a laser or automated plotter cutter (Figure 1).</li>
	<li>Place the plasma-treated coverslip on the taped slide to close the chamber. Use a pipet tip to gently press the coverslip onto the taped regions to make the channel watertight.</li>
	<li>If using glass slides, seal off the edges using epoxy resin. Finally, each imaging chamber made from a slide and coverslip has a dimension of 10 mm x 2 mm x 0.1 mm (length x width x depth). Store the prepared microfluidic imaging chambers for 1-2 weeks in desiccated conditions (preferably between 10-25 &#176;C). 
	&#8203;NOTE: For the laser cut tapes used with acrylic slides, it is not necessary to use epoxy as the tape edges form a tight leakage-proof seal.</li>
</ol>
3. Making supported bilayers on glass substrate by vesicle fusion
NOTE: Crowded supported lipid bilayers are generated on the walls of the imaging chamber by the fusion of lipid vesicles prepared with doped PEG-lipids.
<ol>
	<li>Take a clean glass vial, preferably pre-cleaned using piranha solution. Add chloroform solution of 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC) and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (DOPE-PEG2000) at the desired mole fraction of PEG2000 lipids such that the final concentration after adding buffer will be 3 mM lipids in step 3.4. Prepare other membrane compositions of lipids similarly.</li>
	<li>Dry out the chloroform from the vial using a gentle stream of nitrogen with a small swirl so that the dried lipids are uniformly coated on the surface of the vial. Put the vial in a vacuum desiccator for 1 h. This will remove any residual chloroform in the glass vial. Add 1 mL of phosphate-buffered saline (PBS) and incubate overnight at 37 &#176;C.</li>
	<li>Prepare small unilamellar vesicles (SUVs) as described below.
	<ol>
		<li>Gently vortex for 1-2 min after overnight incubation until the solution turns turbid and milky.</li>
		<li>Take 100 &#181;L of this solution in a small 500 &#181;L centrifuge tube and sonicate for about 1 h in a bath sonicator (set at a 70 W power setting). The solution will become clear; if not, then sonicate for an additional 30 min. This step will generate SUVs of 50-100 nm in diameter. 
		NOTE: If preparing for the first time, characterize the SUVs using dynamic light scattering22,23 (DLS) or an equivalent method.</li>
	</ol>
	</li>
	<li>Add calcium chloride (CaCl2, 3 M stock) to the sonicated vesicles such that its final concentration is 30 mM in the lipid solution.</li>
	<li>Cut a micropipette tip for injecting the sample solution such that it fits tightly in the hole. Mix the lipid solution and inject through one of the holes in the imaging chamber made in step 2. Wipe the excess solution that comes out of the chamber from the outlet with a clean tissue to prevent the contamination of adjacent channels.</li>
	<li>Leave this assembly in a humidified chamber for 90 min. Prepare a humidifying chamber by placing a wet tissue at the end of a 50 mL centrifuge tube. Place the slide sideways inside the tube with tube caps closed. The vesicles will fuse and generate a uniform bilayer on the glass surface.</li>
	<li>Wash the chamber thoroughly with a copious amount of PBS buffer (roughly 5 times the volume of the imaging chamber). Prevent the entry of air bubbles into the chamber while washing as this can generate defects in the membrane.</li>
</ol>
4. Microscope setup and single-particle imaging measurements
NOTE: Single-molecule experiments are carried out on an objective-based total internal reflection fluorescence24,25,26 (TIRF) microscope setup (Figure 2). TIRF imaging provides a better signal-to-noise ratio for single-molecule imaging, though epi-fluorescence microscopes can also be used under certain conditions (especially when the fluorescent biomolecules can be removed from the bulk solution by washing). The prism-type TIRF can be used but the objective-type is preferable for the ease of setting up microfluidics27. For objective-type TIRF, a high numerical aperture objective (100x magnification, usually commercially available as a TIRF objective) is recommended.
<ol>
	<li>Before conducting any experiment on mobility and assembly, align the TIRF microscope to ensure an optimal signal-to-noise ratio due to illumination by the evanescent field. During alignment, keep the laser power low, between 100 &#181;W and 1 mW. 
	CAUTION: Laser safety glasses should be used during the aligning of the laser beam.
	<ol>
		<li>To align the microscope, prepare a fluorescent bead sample by adding them into the microfluidic channel at low concentrations (at ~100 pM) such that fluorescent spots from single beads do not overlap in the images.</li>
		<li>First, visualize the beads in epifluorescence mode. While illumination is in epifluorescence mode, move the M5 mirror translation stage (the mirror that reflects the laser to the dichroic) such that the beam coming out of the objective bends until it eventually is in a total internal reflection configuration.</li>
		<li>Verify whether TIR illumination has been achieved. When the TIR evanescent field is illuminating the bead sample, check that only the beads on the surface are visible and free-floating beads away from the surface are not observed.</li>
	</ol>
	</li>
	<li>At the start of the experiment, set the laser power at the objective back focal plane to 5-10 mW to prevent photo-destruction (photobleaching) of the fluorophores.</li>
	<li>Keep the slide on the microscope stage and focus on the bare membrane (without any fluorophores) first. Usually, small traces of impurities in lipid membranes are sufficient to do this. Optional: Incorporate a small number of fluorescent beads or quantum dots (sub picomolar range) during step 3.1, such that only two to five fluorescent particles are present in the field of view to facilitate identifying the correct focus.</li>
	<li>Inject the sample (membrane-proteins, etc.) using a micropipette into the PEG-SLB-coated imaging chamber made in step 3.7.</li>
	<li>For measuring single-molecule binding kinetics, use a syringe pump to flow the labeled biomolecules through the inlet holes into the microfluidic chamber (Figure 3). Use a flow rate of 50-500 &#181;L/min.
	<ol>
		<li>Start movie acquisition before adding the solution into the imaging chamber. Acquire &#62;5,000 frames at a frame rate of 25-50 frames/s under continuous flow until there is no further increase in the appearance of new spots on the membrane surface (Video 1). For the analysis of binding kinetics, follow the steps from 6.1.</li>
	</ol>
	</li>
	<li>For single-particle tracking, add the labeled biomolecules at low concentrations (compared to the binding constant) to the channel using a micropipette. Optimize the concentration to a density of &#60;0.1 particles/&#181;m2 so that individual particles rarely cross paths (Video 2). This ensures the high fidelity of tracks recovered from the datasets. Incubate if needed (in case of poor membrane binding by biomolecules, ~10 min) in a humidifying environment and wash the chamber with the buffer. 
	NOTE: Washing is needed in case the binding is poor on the membrane and most of the fluorescent molecules remain in the solution. Otherwise, this may interfere with imaging and result in a poor signal-to-noise ratio.</li>
	<li>For single-particle trajectory analysis, acquire 200-500 frames at 10-100 frames/s. Maintain the objective lens focused on the bilayer plane with minimal stage drift during the acquisition interval.</li>
</ol>
5. Image acquisition for counting subunits in a protein assembly
NOTE: Image acquisition for estimating stoichiometry requires continuous bleaching of fluorophores and detecting the number of steps until no more fluorophores are emitting fluorescence.
<ol>
	<li>For measuring subunits, add the relevant concentration of labeled biomolecules (example: see results; ClyA was added at 25 nM concentration) to the microfluidic imaging chamber and incubate (wash if necessary). Incubate the slide in a humidifying chamber at the desired temperature for the required amount of time (example: 37 &#176;C and 60 min for the assembly of ClyA).</li>
	<li>Perform image acquisition for single-molecule photobleaching as described below.
	<ol>
		<li>Wash the imaging chamber with a buffer before imaging (if necessary). Place the slide on the microscope and adjust the focus to visualize the labeled molecules.</li>
		<li>Set the laser power to a level at which the bleaching of the fluorophore occurs gradually (~1-5 min). Ideally, for each assembled biomolecule, keep the photobleaching step rate &#62;10-20 imaging frames apart. Acquire the images until all the molecules are completely photobleached (Video 3). 
		NOTE: To determine the total duration of the photobleaching trajectory required, plot the average intensity of individual spots with the number of imaging frames and determine the time constant by fitting an exponential decay function. The total duration of the acquisition can be set to 5-10 times the time constant.</li>
	</ol>
	</li>
	<li>Perform a time-dependent assembly measurement by starting movie acquisition as soon as the sample is introduced into the chamber and acquiring new photobleaching movies at fixed time intervals after membrane binding from different segments of the PEG-SLB.</li>
</ol>
6. Image and data analysis
<ol>
	<li>Perform single-particle binding and tracking as described below.
	<ol>
		<li>Extract single-particle trajectories from the movies acquired above, as shown in Figure 4. For detecting single particles and tracking them, use the MATLAB package, u-track28, or the Trackmate29, a plugin in ImageJ, which also provides a robust single-particle tracking algorithm. Follow the steps for setting up u-track analysis as shown in Supplementary File 1. 
		NOTE: The critical parameters for single-particle tracking are the standard deviation of the particle size (in pixels) and the gap closing length. Use 1 and 0, respectively, for these parameters as the most stringent criteria for tracking.</li>
		<li>To calculate the binding rate constants, first estimate the number of particles binding to the membrane surface over time. Use the MATLAB file binding_SPT (Supplementary Coding File 2) with the u-track output folder to identify and plot the number of particles appearing on the membrane obtained from step 6.1. Fit the observed kinetics to appropriate kinetic models (example: single exponential fit to determine the time constant, the inverse of which can be assigned to the apparent rate constant)30 to extract the rate constants (see results, Figure 5).</li>
		<li>The mean-squared displacement (MSD) of the diffusing particles (such as DNA tracer in PEG-SLBs, Figure 6) indicates the nature of diffusion. Use the MATLAB package MSD_ISD (Supplementary Coding File 2) with the u-track output folder to obtain the MSD plot as a function of lag time (Figure 6B).</li>
		<li>To identify heterogeneous diffusing species, calculate instantaneous squared displacement (ISD) distributions as shown in Figure 6C. ISD2, defined as (Xt+&#964; - Xt)2 + (Yt+&#964;&#8722; Yt)2, where lag time, &#964; = 2, is preferred as it reduces noise. Filter out short trajectories with less than three frames to reduce noise.</li>
		<li>Use ISD_analyzer (Supplementary Coding File 2) in MATLAB with the u-track output to plot the ISDs of the tracked particles (Figure 6C).</li>
	</ol>
	</li>
	<li>Perform single-molecule photobleaching analysis as described below.
	<ol>
		<li>To estimate the stoichiometry of the membrane complexes, perform a photobleaching analysis on the time-dependent intensities of the fluorescent complexes (Video 3). For this, apply a drift correction algorithm (drift_correct_images, Supplementary Coding File 2) based on the autocorrelation of movie frames to extract the translation drift. This assumes that the assembled structures are largely immobile during image acquisition. Run the MATLAB package Extract_traces_1C (Supplementary Coding File 2) to detect particle intensity time traces from the movies.</li>
		<li>Use the MATLAB code Stepcount_immobile (Supplementary Coding File 2) to perform the step detection for the intensity time traces. In case the particles are not immobile, use the u-track package to extract the location of moving particles. This set of xy-coordinates can be mapped to the raw movies to extract the intensity values of the moving particle as it diffuses laterally on the membrane. Use the MATLAB package utrack_Int (Supplementary Coding File 2) with the output from u-track analysis for this option.</li>
		<li>Perform a correction for incomplete labeling of the biomolecules with fluorophore tags to estimate the correct protein complex stoichiometry. Determine the labeling efficiency with a UV-VIS spectrophotometer by measuring the absorption of dye and protein to estimate the concentration. Calculate the labeling efficiency as the molar ratio of the dye to the protein. 
		NOTE: Some dyes also absorb at wavelengths close to 280 nm, and, hence, this absorption contribution by the dye should be accounted for during the concentration calculation.</li>
		<li>Perform a binomial correction to account for the incomplete labeling efficiency of the fluorophore in the following manner by using the MATLAB code Step_correction (Supplementary Coding File 2) with the data obtained from step counting in step 6.2.2. For example, for a pore-forming toxin like Cytolysin A, which forms a dodecameric ring-like structure, apply a binomial correction using the following formula: 
		nk <img alt="Equation 1" src="/files/ftp_upload/64243/64243eq01.jpg" style="margin: auto;" /> 
		where <img alt="Equation 2" src="/files/ftp_upload/64243/64243eq02.jpg" style="margin: auto;" /> = labeling efficiency, n = number of photobleaching steps, and s&#160;= actual counts. Use the MATLAB routine Stepcount_immobile to recover the corrected oligomeric species. Convert the frequency of oligomers (si) to mass fractions (Figure 7C; Supplementary Coding File 2). 
		NOTE: A set of u-track analyzed files are included in&#160;Supplementary Coding File 3. The MATLAB codes in Supplementary Coding File 2&#160;can be run with these data sets.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Here, we demonstrate single-molecule experiments on supported lipid bilayers (SLBs) that manifest a crowded environment for membrane-embedded biomolecules. The crowded environment generates an excluded volume effect, leading to the enhancement of biomolecular reactions1,2,39,40. For the PEG-lipid system, where the polymer primarily occupies the volume outside the bilayer, this effect is especia...

Cellular membranes are highly crowded and complex systems1. Molecular crowding can have a considerable impact on the diffusion of membrane-bound entities like protein and lipids2,3,4. Similarly, bimolecular reactions on lipid membranes like receptor dimerization or the oligomerization of membrane complexes are influenced by crowding5,6,7. The nature, configuration, and concentration of crowders can govern the membrane binding, diffusivity, and protein-protein interaction in several ways8,9. Since controlling membrane crowding on cellular membranes and interpreting its influence on embedded biomolecules is challenging, researchers have tried to establish alternate in vitro systems10.
A popular approach for artificial crowded membranes is doping the bilayer membranes with polymer (such as polyethylene glycol, PEG)-grafted lipids11,12. During the visualization of protein and lipid dynamics on supported lipid bilayers (SLBs), these polymers additionally shield the membrane-embedded components from the underlying negatively charged substrate (such as glass) by effectively lifting the bilayer away from the underlying support. By varying the size and concentration of the polymer, one can control the extent of molecular crowding, as well as its separation from the underlying solid support13,14. This is clearly an advantage over lipid bilayers supported on solid substrates without polymer cushions15,16, where transmembrane biomolecules can lose their activity17,18,19. More importantly, it allows us to recapitulate the crowded environment of the cell membrane in vitro, which is critical for many membrane processes.
Surface-grafted polymers on membranes also undergo changes in their configuration depending on their grafting density12. At low concentrations, they remain in an entropically coiled configuration, known as a mushroom, above the membrane surface. With increasing concentration, they start to interact and tend to uncoil and extend, finally yielding a dense brush-like formation on the membrane21. Since the transition from the mushroom to the brush regime is highly heterogeneous and manifests in poorly characterized conditions of the polymer, it is important to use well-characterized conditions for crowding on polymer-grafted membranes. Compared to a recent study20, we identify and report crowded membrane compositions that maintain the diffusive transport and activity of transmembrane biomolecules.
In this protocol, we discuss how to generate PEGylated lipid membranes and provide recommendations for PEG densities that mimic crowding in two different regimes of polymer configuration (namely, mushroom and brush). The protocol also describes single-molecule binding, particle tracking, and photobleaching data acquisition and analysis for molecules embedded in these crowded membranes. First, we describe the thorough cleaning steps, the assembly of the imaging chamber, and the generation of PEG-SLBs. Second, we provide details for the single-molecule binding, particle tracking, and photobleaching experiments. Third, we discuss i) extracting the relative binding affinities, ii) characterizing molecular diffusion, and iii) counting subunits in a protein assembly from movies of single molecules on the membrane.
While we characterized this system with single-molecule imaging, the protocol is useful for all membrane biophysicists interested in understanding the effect of crowding on biomolecular reactions on lipid membranes. Overall, we present a robust pipeline for making crowded and supported lipid bilayers, along with various single-molecule assays conducted on them and the corresponding analysis routines.

<table><tbody><tr><td>2.5 ml Syringes</td><td>HMD Healthcare</td><td>Dispo Van, 2.5 ml Tuberculin</td><td>Plastic syringe</td></tr><tr><td>Acetone</td><td>Finar Chemicals</td><td>10020LL025</td><td></td></tr><tr><td>Acrylic Sheet</td><td></td><td></td><td>2 mm thick</td></tr><tr><td>Acrylic Sheet</td><td>BigiMall</td><td></td><td>2 mm, Clear</td></tr><tr><td>Bath Sonicator</td><td>Branson</td><td>CPX-1800</td><td></td></tr><tr><td>Calcium Chloride</td><td></td><td></td><td></td></tr><tr><td>Chloroform</td><td>Sigma</td><td>528730</td><td>&amp;nbsp;HPLC grade</td></tr><tr><td>Cholesterol</td><td>Avanti</td><td>700100</td><td></td></tr><tr><td>Coplin Jar</td><td>Duran Wheaton Kimble</td><td>S6016</td><td>8 Slide Jar with Glass Cover</td></tr><tr><td>Coverslips</td><td>VWR</td><td>631-1574</td><td>24 mm X 50 mm</td></tr><tr><td>Cy3-DNA Strand</td><td>IDT</td><td></td><td>GCTGCTATTGCGTCCGTTTGGTT&lt;br/&gt; GGTGTGGTTGG-Cy3</td></tr><tr><td>Cyanine Dye (Cy3)</td><td>Cytiva Life Sciences</td><td>PA23001</td><td></td></tr><tr><td>DiI</td><td>Invitrogen</td><td>D3911</td><td>Dil Stain (1,1&#039;-Dioctadecyl-3,3,3&#039;,3&#039;-Tetramethylindocarbocyanine Perchlorate (&#039;DiI&#039;; DiIC18(3)))</td></tr><tr><td>DNA Connector Strand 1</td><td>Sigma Aldrich</td><td></td><td>GCTGCTATTGCGTCCGTTTAGCT&lt;br/&gt; GGGGGAGTATTGCGGAGGAAGC&lt;br/&gt; T</td></tr><tr><td>DNA Connector Strand 2</td><td>Sigma Aldrich</td><td></td><td>CGGACGCAATAGCAGCTCACAG&lt;br/&gt; TCGGTCACAT</td></tr><tr><td>DNA Tocopherol Strand</td><td>Biomers</td><td></td><td>Toco-CCCAATGTGACCGACTGTGA</td></tr><tr><td>DOPE-PEG2000</td><td>Avanti</td><td>880130</td><td>1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (ammonium salt)</td></tr><tr><td>Double Sided Tape</td><td>3M</td><td>LF93010LE</td><td></td></tr><tr><td>Drill Bits (Diamond Coated)</td><td></td><td></td><td>0.5 - 1 mm</td></tr><tr><td>Drilling Machine</td><td>Dremel</td><td>220</td><td>Workstation</td></tr><tr><td>EMCCD</td><td>Andor</td><td>DU-897U-CS0-#BV</td><td></td></tr><tr><td>Fluorescence Beads</td><td>Invitrogen</td><td>F10720</td><td></td></tr><tr><td>Glass Slides</td><td>Blue Star</td><td></td><td>Micro Slides, PIC-1</td></tr><tr><td>Glass Vials</td><td>Sigma</td><td>854190</td><td></td></tr><tr><td>Hydrogen Peroxide</td><td>Lobachemie</td><td>00182</td><td>30% Solution, AR Grade</td></tr><tr><td>Labolene</td><td>Thermo-Fischer Scientific</td><td></td><td>&amp;nbsp;Detergent</td></tr><tr><td>Laser 532 nm</td><td>Coherent</td><td>Sapphire</td><td></td></tr><tr><td>Laser Cutter</td><td>Universal Laser Systems</td><td>ILS12.75</td><td></td></tr><tr><td>Lissamine Rhodamine DOPE</td><td>Avanti</td><td>810150</td><td>1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (ammonium salt)</td></tr><tr><td>Methanol</td><td>Finar Chemicals</td><td>30932LL025</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td>IX81</td><td></td></tr><tr><td>Phosphate Buffer Saline (PBS)</td><td></td><td></td><td>1X</td></tr><tr><td>Plasma Cleaner</td><td>Harrick Plasma Inc</td><td>PDC-002</td><td></td></tr><tr><td>POPC</td><td>Avanti</td><td>850457</td><td>1-palmitoyl-2-oleoyl-glycero-3-phosphocholine</td></tr><tr><td>Programmable Syringe Pump</td><td>New Era Pump Systems</td><td>NE1010</td><td>High Pressure Syringe Pump</td></tr><tr><td>PTFE Caps</td><td>Sigma</td><td>27141</td><td></td></tr><tr><td>PTFE Tubing</td><td>Cole-Parmer</td><td>WW-06417-21</td><td>Masterflex, 0.022&quot; ID x 0.042&quot; OD</td></tr><tr><td>Sulphuric Acid</td><td>SD Fine Chemicals</td><td></td><td>98%, AR Grade</td></tr><tr><td>TIRF Objective</td><td>Olympus</td><td>UPLAPO100XOHR</td><td></td></tr><tr><td>Vacuum Desiccator</td><td>Tarsons</td><td></td><td></td></tr><tr><td>Vortex Mixer</td><td>Tarsons</td><td></td><td></td></tr></tbody></table>

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.

Monitoring the binding of ClyA protein on PEGylated membranes 
After step 4.5, the binding kinetics are estimated by plotting the number of particles binding to the membrane surface over time (Video 1). As ClyA protein binds to a membrane with 5 mol% PEG2000 lipids,the particle density increases and reaches saturation (Figure 5). An exponential decay fit to the bound particles (cyan circles) gives the time constant (&#964;b) for the membrane ...

Watch this Scientific Journal Video about Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes at JoVE.com

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

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 ...

single-molecule-diffusion-assembly-on-polymer-crowded-lipid

Research

JoVE Journal

Biochemistry

2.2K Views.  Indian Institute of Science. Celmembranen zijn zeer druk vanwege de aanwezigheid van ingebedde eiwitten en suikers. We demonstreren beeldvorming van één molecuul op synthetische overvolle ondersteunde lipide bilayers. Dit platform is nuttig voor het onderzoeken van het effect van verdringing op membraanbiomoleculaire reacties, op moleculair niveau.Het protocol verklaart de analyse van binding, diffusie en assemblage van membraanbiomoleculen. Succes in het experiment hangt af van een rigoureuze reiniging van het ...

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

Synthesis of Cyclic Polymers and Characterization of Their Diffusive Motion in the Melt State at the Single Molecule Level

We demonstrate a method for the synthesis of cyclic polymers and a protocol for characterizing their diffusive motion in a melt state at the single molecule level. An electrostatic self-assembly and covalent fixation (ESA-CF) process is used for the synthesis of the cyclic poly(tetrahydrofuran) (poly(THF)). The diffusive motion of individual cyclic polymer chains in a melt state is visualized using single molecule fluorescence imaging by incorporating a fluorophore unit in the cyclic chains. The diffusive motion of the chains is quantitatively characterized by means of a combination of mean-squared displacement (MSD) analysis and a cumulative distribution function (CDF) analysis. The cyclic polymer exhibits multiple-mode diffusion which is distinct from its linear counterpart. The results demonstrate that the diffusional heterogeneity of polymers that is often hidden behind ensemble averaging can be revealed by the efficient synthesis of the cyclic polymers using the ESA-CF process and the quantitative analysis of the diffusive motion at the single molecule level using the MSD and CDF analyses.

A protocol for the synthesis and characterization of diffusive motion of cyclic polymers at the single molecule level is presented.

We demonstrate a method for the synthesis of cyclic polymers and a protocol for characterizing their diffusive motion in a melt state at the single ...

Chemistry

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Bioengineering

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

Fluorescence Recovery after Merging a Droplet to Measure the Two-dimensional Diffusion of a Phospholipid Monolayer

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after merging (FRAM). FRAM adopts the same principles as the fluorescence recovery after photobleaching (FRAP) technique, especially for measuring fluorescence recovery after bleaching a specific area, but FRAM uses a drop coalescence instead of photobleaching dye molecules to induce a chemical potential gradient of dye molecules. Our technique has several advantages over FRAP: it only requires a fluorescence microscope rather than a confocal microscope equipped with high power lasers; it is essentially free from the selection of fluorescence dyes; and it has far more freedom to define the measured diffusion area. Furthermore, FRAM potentially provides a route for studying the mixing or inter-diffusion of two different surfactants, when the monolayers at a surface of droplet and at a flat air/water interface are prepared with different species, independently.

We present a new technique to measure the lateral diffusion of a surface active species at the fluid-fluid interface by merging a droplet monolayer onto a flat monolayer.

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after ...

Automated Lipid Bilayer Membrane Formation Using a Polydimethylsiloxane Thin Film

An artificial lipid bilayer, or black lipid membrane (BLM), is a powerful tool for studying ion channels and protein interactions, as well as for biosensor applications. However, conventional BLM formation techniques have several drawbacks and they often require specific expertise and laborious processes. In particular, conventional BLMs suffer from low formation success rates and inconsistent membrane formation time. Here, we demonstrate a storable and transportable BLM formation system with controlled thinning-out time and enhanced BLM formation rate by replacing conventionally used films (polytetrafluoroethylene, polyoxymethylene, polystyrene) to polydimethylsiloxane (PDMS). In this experiment, a porous-structured polymer such as PDMS thin film is used. In addition, as opposed to conventionally used solvents with low viscosity, the use of squalene permitted a controlled thinning-out time via slow solvent absorption by PDMS, prolonging membrane lifetime. In addition, by using a mixture of squalene and hexadecane, the freezing point of the lipid solution was increased (~16 &#176;C), in addition, membrane precursors were produced that can be indefinitely stored and readily transported. These membrane precursors have reduced BLM formation time of &#60; 1 hr and achieved a BLM formation rate of ~80%. Moreover, ion channel experiments with gramicidin A demonstrated the feasibility of the membrane system.

We demonstrate a storable, transportable lipid bilayer formation system. A lipid bilayer membrane can be formed within 1 hr with over 80% success rate when a frozen membrane precursor is brought to ambient temperature. This system will reduce laborious processes and expertise associated with ion channels.

An artificial lipid bilayer, or black lipid membrane (BLM), is a powerful tool for studying ion channels and protein interactions, as well as for ...

Self-Assembly of Hybrid Lipid Membranes Doped with Hydrophobic Organic Molecules at the Water/Air Interface

Because of their unique properties, including an ultrathin thickness (3-4 nm), ultrahigh resistivity, fluidity and self-assembly ability, lipid bilayers can be readily functionalized and have been used in various applications such as bio-sensors and bio-devices. In this study, we introduced a planar organic molecule: copper (II) 2,9,16,23-tetra-tert-butyl-29H,31H-phthalocyanine (CuPc) to dope lipid membranes. The CuPc/lipid hybrid membrane forms at the water/air interface by self-assembly. In this membrane, the hydrophobic CuPc molecules are located between the hydrophobic tails of lipid molecules, forming a lipid/CuPc/lipid sandwich structure. Interestingly, an air-stable hybrid lipid bilayer can be readily formed by transferring the hybrid membrane onto a Si substrate. We report a straightforward method for incorporating nanomaterials into a lipid bilayer system, which represents a new methodology for the fabrication of biosensors and biodevices.

We report a protocol for producing a hybrid lipid membrane at the water/air interface by doping the lipid bilayer with copper (II) 2,9,16,23-tetra-tert-butyl-29H,31H-phthalocyanine (CuPc) molecules. The resulting hybrid lipid membrane has a lipid/CuPc/lipid sandwich structure. This protocol can also be applied to the formation of other functional nanomaterials.

Because of their unique properties, including an ultrathin thickness (3-4 nm), ultrahigh resistivity, fluidity and self-assembly ability, lipid bilayers ...

Reconstitution of Septin Assembly at Membranes to Study Biophysical Properties and Functions

Most cells can sense and change their shape to carry out fundamental cell processes. In many eukaryotes, the septin cytoskeleton is an integral component in coordinating shape changes like cytokinesis, polarized growth, and migration. Septins are filament-forming proteins that assemble to form diverse higher-order structures and, in many cases, are found in different areas of the plasma membrane, most notably in regions of micron-scale positive curvature. Monitoring the process of septin assembly in vivo is hindered by the limitations of light microscopy in cells, as well as the complexity of interactions with both membranes and cytoskeletal elements, making it difficult to quantify septin dynamics in living systems. Fortunately, there has been substantial progress in the past decade in reconstituting the septin cytoskeleton in a cell-free system to dissect the mechanisms controlling septin assembly at high spatial and temporal resolutions. The core steps of septin assembly include septin heterooligomer association and dissociation with the membrane, polymerization into filaments, and the formation of higher-order structures through interactions between filaments. Here, we present three methods to observe septin assembly in different contexts: planar bilayers, spherical supports, and rod supports. These methods can be used to determine the biophysical parameters of septins at different stages of assembly: as single octamers binding the membrane, as filaments, and as assemblies of filaments. We use these parameters paired with measurements of curvature sampling and preferential adsorption to understand how curvature sensing operates at a variety of length and time scales.

Cell-free reconstitution has been a key tool to understand the cytoskeleton assembly, and work in the last decade has established approaches to study septin dynamics in minimal systems. Presented here are three complementary methods to observe septin assembly in different membrane contexts: planar bilayers, spherical supports, and rod supports.

Most cells can sense and change their shape to carry out fundamental cell processes. In many eukaryotes, the septin cytoskeleton is an integral component ...

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.

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.

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

Assembly of Cell Mimicking Supported and Suspended Lipid Bilayer Models for the Study of Molecular Interactions

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial protective barrier for all cell types, separating the internal cellular components from the extracellular environment. These membranes are composed largely of a lipid bilayer, which contains outer hydrophilic head groups and inner hydrophobic tail groups, along with various proteins and cholesterol. The composition and structure of the lipids themselves play a crucial role in regulating biological function, including interactions between cells and the cellular microenvironment, which may contain pharmaceuticals, biological toxins, and environmental toxicants. In this study, methods to formulate uni-lipid and multi-lipid supported and suspended cell mimicking lipid bilayers are described. Previously, uni-lipid phosphatidylcholine (PC) lipid bilayers as well as multi-lipid placental trophoblast-inspired lipid bilayers were developed for use in understanding molecular interactions. Here, methods for achieving both types of bilayer models will be presented. For cell mimicking multi-lipid bilayers, the desired lipid composition is first determined via lipid extraction from primary cells or cell lines followed by liquid chromatography-mass spectrometry (LC-MS). Using this composition, lipid vesicles are fabricated using a thin-film hydration and extrusion method and their hydrodynamic diameter and zeta potential are characterized. Supported and suspended lipid bilayers can then be formed using quartz crystal microbalance with dissipation monitoring (QCM-D) and on a porous membrane for use in a parallel artificial membrane permeability assay (PAMPA), respectively. The representative results highlight the reproducibility and versatility of in vitro cell membrane lipid bilayer models. The methods presented can aid in rapid, facile assessment of the interaction mechanisms, such as permeation, adsorption, and embedment, of various molecules and macromolecules with a cell membrane, helping in the screening of drug candidates and prediction of potential cellular toxicity.

This protocol describes the formation of cell mimicking uni-lipid and multi-lipid vesicles, supported lipid bilayers, and suspended lipid bilayers. These in vitro models can be adapted to incorporate a variety of lipid types and can be used to investigate various molecule and macromolecule interactions.

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial ...

On-Chip Octanol-Assisted Liposome Assembly for Bioengineering

Microfluidics is a widely used tool to generate droplets and vesicles of various kinds in a controlled and high-throughput manner. Liposomes are simplistic cellular mimics composed of an aqueous interior surrounded by a lipid bilayer; they are valuable in designing synthetic cells and understanding the fundamentals of biological cells in an in vitro fashion and are important for applied sciences, such as cargo delivery for therapeutic applications. This article describes a detailed working protocol for an on-chip microfluidic technique, octanol-assisted liposome assembly (OLA), to produce monodispersed, micron-sized, biocompatible liposomes. OLA functions similarly to bubble blowing, where an inner aqueous (IA) phase and a surrounding lipid-carrying 1-octanol phase are pinched off by surfactant-containing outer fluid streams. This readily generates double-emulsion droplets with protruding octanol pockets. As the lipid bilayer assembles at the droplet interface, the pocket spontaneously detaches to give rise to a unilamellar liposome that is ready for further manipulation and experimentation. OLA provides several advantages, such as steady liposome generation (&#62;10 Hz), efficient encapsulation of biomaterials, and monodispersed liposome populations, and requires very small sample volumes (~50 &#181;L), which can be crucial when working with precious biologicals. The study includes details on microfabrication, soft-lithography, and surface passivation, which are needed to establish OLA technology in the lab. A proof-of-principle synthetic biology application is also shown by inducing the formation of biomolecular condensates inside the liposomes via transmembrane proton flux. It is anticipated that this accompanying video protocol will facilitate the readers to establish and troubleshoot OLA in their labs.

The present protocol describes octanol-assisted liposome assembly (OLA), a microfluidic technique to generate biocompatible liposomes. OLA produces monodispersed, micron-sized liposomes with efficient encapsulation, allowing immediate on-chip experimentation. This protocol is anticipated to be particularly suitable for synthetic biology and synthetic cell research.

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

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