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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>&#160;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 
			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&#39;-Dioctadecyl-3,3,3&#39;,3&#39;-Tetramethylindocarbocyanine Perchlorate (&#39;DiI&#39;; DiIC18(3)))</td>
		</tr>
		<tr>
			<td>DNA Connector Strand 1</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>GCTGCTATTGCGTCCGTTTAGCT 
			GGGGGAGTATTGCGGAGGAAGC 
			T</td>
		</tr>
		<tr>
			<td>DNA Connector Strand 2</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>CGGACGCAATAGCAGCTCACAG 
			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>&#160;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&#34; ID x 0.042&#34; 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

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

2.2K Views.  Indian Institute of Science. 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. 

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

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Conventional BODIPY Conjugates for Live-Cell Super-Resolution Microscopy and Single-Molecule Tracking

Single molecule localization microscopy (SMLM) techniques overcome the optical diffraction limit of conventional fluorescence microscopy and can resolve intracellular structures and the dynamics of biomolecules with ~20 nm precision. A prerequisite for SMLM are fluorophores that transition from a dark to a fluorescent state in order to avoid spatio-temporal overlap of their point spread functions in each of the thousands of data acquisition frames. BODIPYs are well-established dyes with numerous conjugates used in conventional microscopy. The transient formation of red-shifted BODIPY ground-state dimers (DII) results in bright single molecule emission enabling single molecule localization microscopy (SMLM). Here we present a simple but versatile protocol for SMLM with conventional BODIPY conjugates in living yeast and mammalian cells. This procedure can be used to acquire super-resolution images and to track single BODIPY-DII states to extract spatio-temporal information of BODIPY conjugates. We apply this procedure to resolve lipid droplets (LDs), fatty acids, and lysosomes in living yeast and mammalian cells at the nanoscopic length scale. Furthermore, we demonstrate the multi-color imaging capability with BODIPY dyes when used in conjunction with other fluorescent&#160;probes. Our representative results show the differential spatial distribution and mobility of BODIPY-fatty acids and neutral lipids in yeast under fed and fasted conditions. This optimized protocol for SMLM can be used with hundreds of commercially available BODIPY conjugates and is a useful resource to study biological processes at the nanoscale far beyond the applications of this work.&#160;

Conventional BODIPY conjugates can be used for live-cell single-molecule localization microscopy (SMLM) through exploitation of their transiently forming, red-shifted ground state dimers. We present an optimized SMLM protocol to track and resolve subcellular neutral lipids and fatty acids in living mammalian and yeast cells at the nanoscopic length scale.

Single molecule localization microscopy (SMLM) techniques overcome the optical diffraction limit of conventional fluorescence microscopy and can resolve ...

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...

Making Precise and Accurate Single-Molecule FRET Measurements using the Open-Source smfBox

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes measurements on freely diffusing biomolecules more accessible. This overview includes a step-by-step protocol for using this instrument to make measurements of precise FRET efficiencies in duplex DNA samples, including details of the sample preparation, instrument setup and alignment, data acquisition, and complete analysis routines. The presented approach, which includes how to determine all the correction factors required for accurate FRET-derived distance measurements, builds on a large body of recent collaborative work across the FRET Community, which aims to establish standard protocols and analysis approaches. This protocol, which is easily adaptable to a range of biomolecular systems, adds to the growing efforts in democratising smFRET for the wider scientific community.

This article provides step-by-step instructions for making fully-corrected accurate FRET measurements on individual, freely diffusing biomolecules using the open-source, inexpensive smfBox, from switch on, through alignment and focusing, to data collection and analysis.

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes ...

Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately 20 nm to accommodate large tRNA substrates at the A-, P- and E-sites. Consequently, the ribosome dynamics are naturally de-phased quickly. Single molecule method can detect each ribosome separately and distinguish inhomogeneous populations, which is essential to reveal the complicated mechanisms of multi-component systems. We report the details of a smFRET method based on the Nikon Ti2 inverted microscope to probe the ribosome dynamics between the ribosomal protein L27 and tRNAs. The L27 is labeled at its unique Cys 53 position and reconstituted into a ribosome that is engineered to lack L27. The tRNA is labeled at its elbow region. As the tRNA moves to different locations inside the ribosome during the elongation cycle, such as pre- and post- translocation, the FRET efficiencies and dynamics exhibit differences, which have suggested multiple subpopulations. These subpopulations are not detectable by ensemble methods. The TIRF-based smFRET microscope is built on a manual or motorized inverted microscope, with home-built laser illumination. The ribosome samples are purified by ultracentrifugation, loaded into a home-built multi-channel sample cell and then illuminated via an evanescent laser field. The reflection laser spot can be used to achieve feedback control of perfect focus. The fluorescence signals are separated by a motorized filter-turret and collected by two digital CMOS cameras. The intensities are retrieved via the NIS-Elements software.

Single molecule fluorescence energy transfer is a method that tracks the tRNA dynamics during ribosomal protein synthesis. By tracking individual ribosomes, inhomogeneous populations are identified, which shed light on mechanisms. This method can be used to track biological conformational changes in general to reveal dynamic-function relationships in many other complexed biosystems. Single molecule methods can observe non-rate limiting steps and low-populated key intermediates, which are not accessible by conventional ensemble methods due to the average effect.

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately ...

Single-Molecule Imaging of EWS-FLI1 Condensates Assembling on DNA

The fusion genes resulting from chromosomal translocation have been found in many solid tumors or leukemia. EWS-FLI1, which belongs to the FUS/EWS/TAF15 (FET) family of fusion oncoproteins, is one of the most frequently involved fusion genes in Ewing sarcoma. These FET family fusion proteins typically harbor a low-complexity domain (LCD) of FET protein at their N-terminus and a DNA-binding domain (DBD) at their C-terminus. EWS-FLI1 has been confirmed to form biomolecular condensates at its target binding loci due to LCD-LCD and LCD-DBD interactions, and these condensates can recruit RNA polymerase II to enhance gene transcription. However, how these condensates are assembled at their binding sites remains unclear. Recently, a single-molecule biophysics method-DNA Curtains-was applied to visualize these assembling processes of EWS-FLI1 condensates. Here, the detailed experimental protocol and data analysis approaches are discussed for the application of DNA Curtains in studying the biomolecular condensates assembling on target DNA.

Here, we describe the use of the single-molecule imaging method, DNA Curtains, to study the biophysical mechanism of EWS-FLI1 condensates assembling on DNA.

The fusion genes resulting from chromosomal translocation have been found in many solid tumors or leukemia. EWS-FLI1, which belongs to the FUS/EWS/TAF15 ...