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

1. Preparation of a PDMS Block to Form the Microfluidic Channel

Figure 1. Microfabrication of flow cell template and PDMS block preparation. (A) Design of a four-channel flow cell that fits onto a 24 x 60 mm glass coverslip (bottom). Six identical designs are arranged to fit onto a 10 cm silicon wafer (top). (B) Cut out block of approximately 5-8 mm thick PDMS on a hole puncher. (C) Insertion of tubing into the punched hole using a pair of tweezers. Please click here to view a larger version of this figure.

1. Obtain a flow cell template such as the one in Figure 1A. Typical channels are 0.3-1 mm wide, 70-100 µm high, and 1-2 cm long. Fabricate a template using standard photolithography techniques in a cleanroom facility²⁵ or order if cleanroom access is not available. Alternatively, machine a template with larger dimensions from a suitable material.
   Note: Cleanroom staff can train and guide unexperienced users (access to a cleanroom facility is always restricted to users with proper training) in the design and ordering of a mask, wafer cleaning, and photolithography. Once a template is obtained, it can be used outside the cleanroom repeatedly, provided it is kept in a closed dish and care is taken to exclude dust.
2. Prepare a mixture of ~100 ml PDMS (silicone elastomer base) and ~10 ml of cross-linker (curing agent) in a disposable plastic cup. Break off the tip of a disposable pipette to handle the viscous PDMS more easily. Weigh the PDMS into the plastic cup as pipetting is not accurate.
3. Stir the mixture well. Remove the many air bubbles by degassing in a vacuum desiccator (about 20 min). As the cup is placed under vacuum, the bubbles will initially grow in size, increasing the volume of the mixture greatly. Control the vacuum that is applied and make sure the cup is deep enough to avoid a spill.
4. Pour a large drop of the degassed PDMS mixture into a glass Petri dish (150 mm x 20 mm) and press the wafer onto the PDMS with the template facing up. This avoids bubbles being trapped beneath the wafer. Trapped bubbles will expand and may tilt the wafer when the dish is placed in the oven. Now pour degassed PDMS onto the template on top of the wafer until it is covered by about 5-8 mm of PMDS. If an air bubble occurs gently remove it with a pipette tip.
5. Bake the PMDS in an oven at 60 °C for 3 hr. Ensure that the dish is level.
6. Use a new scalpel blade to cut out a PDMS block containing the molded channel structures. The cut out block needs to fit onto a coverslip (see step 4.1.9).
7. Peel out the cut out block from the dish and place it onto a piece of clean aluminum foil.
   Note: The cross-linked PMDS blocks can be kept for a few months. On a single wafer, there are 6 sets of flow channels (Figure 1A), so 6 PDMS blocks can be produced in a single PDMS molding. After cutting out all 6 of the PDMS blocks, clean loose pieces of PDMS, but otherwise do not remove the remaining PDMS before pouring and cross-linking the next batch, which will require only about half the PDMS used for the first batch. A good template can last a few years.
8. Use a hole puncher to drill through the PDMS block in one straight motion (Figure 1B). Start on the side of the channel grooves. Be sure to remove the punched out piece of PDMS. Repeat this for all eight holes of the four channel design.
9. Place the PDMS block channel side down onto a new and wrinkle-free piece of aluminum foil. Store the block for up to a few months in a dry box, ideally in a desiccator.
10. Push the tubing (0.25 mm ID, 0.76 mm OD) about one-third into the punched hole using a pair of tweezers (Figure 1C). Cut the tubing slanted for easier insertion. Leave tubes long enough to reach the SUV reservoir and the syringe pump, respectively. After placing the assembled chip onto the microscope, cut the tubes again if they are too long.
11. To connect to the syringes of the pump, cut a short piece of larger silicone tubing (0.51 mm ID, 2.1 mm OD) and insert the thinner tube into one side. This ready-to-use block of PDMS can be kept for a few months.

2. Coverslip Cleaning

1. Clean coverslips according to the protocol described in Karatekin and Rothman²⁵ using a strong oxidizing mixture of sulfuric acid (H₂SO₄) and hydrogen peroxide (H₂O₂). Perform this procedure in a cleanroom, and observe proper safety precautions. Alternatively, use a cleanroom cleaned coverslip that is commercially available (see Materials List).

3. Preparation of v-SUVS Containing Both Lipid and Content Labels

Figure 2. Schematic of SUV preparation. Lipids are mixed in a glass tube (1) and solvent is evaporated to form a lipid film by rotating the tube in a water bath (2). Remaining traces of solvent are removed under high vacuum (3). The lipid film is hydrated in reconstitution buffer containing detergent and protein while vortexed (4). If contents dye is to be encapsulated, it is included in this step as well as in the dilution step (5). Dilution of the detergent concentration below its critical micelle concentration leads to liposome formation. Detergent is dialyzed away overnight (6). For t-SUVs for SBL formation including NBD-PE (green), vesicles are floated in a density gradient and collected at the interface between two layers (7a). To separate v-SUVs with encapsulated content marker from free dye the sample is run over a size-exclusion column and collected in 0.5 ml fractions (7b). Please click here to view a larger version of this figure.

1. Prepare 4 L of reconstitution buffer, containing 25 mM HEPES-KOH, 140 mM KCl, 100 µM EGTA and 1 mM DTT, pH 7.4. Use most of the buffer for dialysis and 100 ml for other steps.
   Note: Other buffers may be used, but it is important to keep the buffer osmolarity consistent throughout the experiment and choose conditions in which the proteins are functional. For preparing v-SUVs containing lipid labels only and t-SUVs, follow Karatekin and Rothman²⁵.
2. As lipid stocks are stored at -20 °C in chloroform or chloroform:methanol (2:1, v/v), let vials reach room temperature before opening to avoid condensation.
3. Rinse a glass tube with chloroform to remove trace amounts of detergent and mix the lipids at the desired ratio with a final amount of 1 µmol in a mixture of chloroform and methanol (2:1, v/v). Only use glass syringes/tubes to handle organic solvents/dissolved lipids.
   Note: The lipids used here are POPC:SAPE:DOPS:Cholesterol:PEG2000-DOPE:DiD at a molar ratio of 57.4:15:12:10:4.6:1 for v-SNARE containing vesicles and POPC:SAPE:DOPS:Cholesterol:brain PI(4,5)P₂:PEG2000-DOPE:NBD-PE (54.9:15:12:10:3:4.6:0.5) for t-SNARE SUVs. See Material for details. Other lipid compositions can be used.
4. Evaporate the solvent under a gentle stream of nitrogen, or in a rotary evaporator. Immerse the tip of the tube in a water bath (37 °C) heated above the highest melting temperature of the lipids to yield a homogenous lipid film and to avoid large changes in temperature while evaporating the solvent. Start vacuum at around 300 mbar until the lipid film is formed, then continue for ~2 min at highest possible vacuum in the rotary evaporator.
5. Remove the glass tube and wrap it in aluminum foil to avoid light exposure and remove remaining traces of solvent by placing the tube in a desiccator under high vacuum for at least 2 hr. This step can also run overnight.
6. To rehydrate the dried lipid film, prepare a mixture of detergent (n-Octyl-β-D-glucopyranoside, OG), protein (v-SNAREs), and SRB in reconstitution buffer (500 µl final volume). Keep the final detergent concentration ~2 times above the critical micelle concentration (CMC, 20-25 mM for OG) and adjust the final volume such that the detergent to lipid ratio is >10. Dissolve Sulforhodamine B (SRB) powder in the protein-detergent solution to obtain a final concentration of 10-50 mM SRB by gently shaking the solution. Adjust the osmolarity of the reconstitution buffer when adding high concentrations of SRB by reducing the concentration of potassium chloride accordingly.
   Note: Lipid-to-protein ratio (LP) for t-SNAREs is high (LP ~20,000, ~70 t-SNAREs per sq. micron²⁵). Significantly higher t-SNARE densities in the SBL may lead to inactive protein aggregates with significantly reduced the fusion rates^17,24. LP for v-SNAREs is much lower (LP ~200, ~7,000 v-SNAREs per µm²), close to the v-SNARE densities found on synaptic vesicles^46,47. To ensure protein function after recombinant expression and purification of v- and t-SNAREs it is useful to perform a simple bulk fusion assay⁴⁸ before going through all the steps of the single vesicle fusion assay.
7. Gently shake the pre-warmed glass tube containing the dried lipid film while adding the protein-detergent-SRB solution from step 3.6. Try to avoid creating bubbles. Continue to shake for 15 min at 37 °C.
8. Dilute the detergent fivefold by adding reconstitution buffer containing SRB (add 2 ml, 2.5 ml final volume) while rapidly vortexing to avoid concentration gradients. Continue to shake for 15 min to 1 hr at 37 °C.
   Note: Longer incubation increase protein reconstitution efficiency. The rapid dilution decreases the detergent concentration below the CMC and leads to the formation of small liposomes.
9. Dialyze the vesicle suspension first against ~1 L of reconstitution buffer for 1-2 hr at ambient temperature and then against 3 L of reconstitution buffer overnight at 4 °C with 4 g of polystyrene adsorbent, using a 20,000 MWCO dialysis tube or cassette. Use different beakers for dialysis of vesicles with and without SRB to prevent cross contamination.
10. Equilibrate a gel filtration column with reconstitution buffer. Run the vesicle suspension through the column to separate free SRB from the v-SUVs with encapsulated SRB. Use reconstitution buffer without SRB as eluent once the entire sample has entered the column. Collect v-SUVs in 0.5 ml fractions.
11. Verify successful SRB encapsulation at self-quenched concentrations by measuring SRB fluorescence before and after addition of detergent to an aliquot of vesicles¹⁶. Using a fluorescence spectrometer, excite the sample at 550 nm and scan the SRB emission between 570 nm and 630 nm. Expect a 4-8 fold increase in SRB fluorescence depending on the encapsulated amount upon detergent addition as the membrane is solubilized and the released SRB is diluted.
12. Characterize the lipid and protein recovery using fluorescence spectroscopy²⁴ and SDS-PAGE gel electrophoresis, respectively. Use a sensitive staining method, especially for the high LP t-SUV samples (see Materials List). Typically about 50% of both lipid and protein input is lost during the preparation resulting in a LP close to the nominal value.
13. Characterize SUV sizes using dynamic light scattering²⁴ or electron microscopy⁴⁸. Store SRB containing v-SUVs at 4 °C up to ~3-4 days. Do not freeze, as freezing and thawing breaks the membrane and releases encapsulated SRB.

4. SUV-SBL Fusion Assay to Monitor Lipid Release Only

1. Formation of the tethered supported bilayer within the microfluidic flow channels
   1. Place the PDMS block (step 1.11) under high vacuum for at least 20 min prior to the experiment to remove dissolved gasses. This greatly reduces the risk of air bubbles in the microfluidic channels during the fusion experiment.
   2. Turn on the microscope setup and heat the stage and sample holder (Figures 3 and 4) to the desired temperature.
   3. Filter the reconstitution buffer used to dilute the vesicles through a filter with 0.45 µm or smaller pore size.
   4. Dilute ~30 µl of the NBD-PE labeled t-SUVs or protein-free (pf-SUVs) control liposomes (stock solution 0.5-1 mM lipid) with ~60 µl of buffer. The final concentration is not critical here.
   5. Degas this mixture using a 3 ml syringe. Press out most of the air above the sample while holding the syringe vertically. Seal the (needle free) tip using paraffin film and create a vacuum by pulling down the plunger. Tap the syringe barrel to accelerate degassing of the solution. Repeat this process a few times until no more bubbles occur when vacuum is applied.
   6. Use a hypodermic needle with a diameter slightly larger than the tubing that is attached to the PDMS block to punch a hole in the cap of a microcentrifuge tube. Make sure that the punched out piece of plastic is not inside the tube as it might clog the tubing connected to the microfluidic channel inlet later.
   7. Fill the degassed SUV solution into the microcentrifuge tube having a hole in the cap and place it into the holder on the microscope stage to equilibrate to the set temperature.
   8. Place a previously cleaned coverslip (section 2) in a plasma cleaner and run air plasma for about 5 min. Put the plasma treated coverslip (treated side facing up) on top of a few lint-free tissues serving as a cushion.
   9. Remove the aluminum foil from the degassed PDMS block and place the block on top of the coverslip. When reusing the PDMS block put on and detach a piece of adhesive tape to the channel side to clean it. Press down the PDMS block onto the coverslip using a pair of tweezers to make it stick, but do not press too hard as the glass might break.
   10. Place the assembled flow cell onto the microscope stage and connect the tubing to the SUV reservoir and the syringe pump, respectively. Tape the coverslip to the stage (Figure 3B).
   11. Start aspirating SUVs at 3 µl/min until the solution fills the channels completely (~2.25 mm/sec for the 75 µm x 300 µm channel cross-section). When solutions for all channels start moving up the tubing on the exit side, reduce flow to 0.5 µl/min and incubate for 30-45 min.
       Note: The time between plasma treating the glass slide and flowing the SUVs into the channel should not exceed 10-20 min as the effect of the plasma treatment is transient.
   12. Check the channels for any leak. Use a 10-20X air objective to observe NBD-PE fluorescence from the NBD-PE containing t- or pf-SUVs. Excite NBD fluorophores using the 488 nm laser. Leaks are difficult to detect using bright field illumination.
   13. Put a tube with degassed reconstitution buffer into the holder and let the temperature equilibrate as fast changes in temperature might cause defects in the SBL. Stop the flow and wait ~1 min to make sure the flow has stopped completely and no air bubble will be aspirated into the tube before switching the inlet tubes to the degassed buffer.
   14. Rinse all channels with degassed buffer to wash away unbound SUVs.
   15. Switch to a higher magnification TIRF objective (60X, oil, NA 1.45-1.49) and verify that the bilayer looks homogeneous and is free of any obvious large-scale defects such as dark patches or lipid tubules extending out from the SBL.
2. Check the fluidity of the bilayer
   1. If a dedicated FRAP unit is not available, or if a FRAP sequence cannot be programmed, then qualitatively test membrane fluidity as follows.
      1. Close the field diaphragm to a small size (~40 µm diameter) and adjust the 488 nm excitation light intensity through the software (20-80 µW, or 15-60 nW/µm²) to bleach the NBD-PE fluorescence in the exposed area significantly, but not completely. For a fluid bilayer, at steady state, the fluorescence intensity in the middle of the exposed area should be lower than at the edges, as intact NBD-PE molecules enter the exposed area and diffuse a certain distance before bleaching. In contrast, if the surface-adhered SUVs failed to burst, or for some other reason the supported bilayer is not fluid, all fluorophores in the exposed area should bleach.
         Note: Laser intensity values in this and later steps are given as a rough starting point and should be optimized for a given set of conditions.
      2. Stop the illumination and start it again a few minutes later to verify the results of the steady state measurements
   2. Program a FRAP sequence for a more quantitative measurement, if possible. See supplemental files and the corresponding legend for details.
      Note: Sometimes SUVs will adhere onto the glass coverslip, but fail to burst and form a fluid bilayer. If this occurs, rinse the channels with degassed reconstitution buffer containing 10 mM Mg²⁺ to help supported bilayer formation. Use fluorescence bleaching to assess bilayer fluidity as in 4.2. Once a fluid bilayer is formed, rinse with Mg²⁺-free reconstitution buffer.
3. Introducing v-SUVs into the microfluidic flow channels
   1. Degas reconstitution buffer and use it to dilute the v-SUV stock solution by a factor of about 10³ to 10⁵ depending on the v-SUV stock concentration. Aim for a dilution that results in about 10-100 fusion events within 60 sec in the field of view.
      Note: Too many fusions increase the background fluorescence (since each event deposits LR-PE or DiD lipid labels into the SBL) and make detection and analysis of fusion events difficult. In contrast, a fusion rate that is too low results in poor statistics¸ or requires acquisition of many more movies. For a v-SUV concentration of 0.1 mM lipid, start by diluting 5 µl SUV stock in 995 µl reconstitution buffer and then dilute 5-50 µl of this in 950-995 µl reconstitution buffer.
   2. Let temperature equilibrate before stopping the flow and inserting the inlet tube into the diluted v-SUV solution.
   3. Adjust the TIRF angle and polarization as follows.
      1. After setting the desired polarization by rotating the excitation beam, adjust the tilt of the mirror commanding the position of the beam via a stepper motor through the software. Slowly move the position of the laser beam from the center of the objective at the back focal plane to one side off-center. Observe the light from the objective at the front side emerge with an increasing angle with respect to the objective axis as the position is moved further off-center.
      2. Note the motor position when the exiting beam first disappears into the objective, i.e., when TIR is first achieved.
      3. Slowly keep moving the beam position further off-center while monitoring the fluorescence from the surface. Note the motor position when the surface fluorescence disappears when the beam has moved too far off-center.
      4. Choose a beam position between the two limits determined above. For best signal-to-noise ratio and more signal enhancement upon fusion, choose a shallow penetration depth (beam position closer to the edge of the objective) that still provides uniform illumination of the viewfield.
         Note: It is best to keep the same TIR beam position (same penetration depth) for all experiments after settings are optimized. Make sure rotating the polarization does not result in changes in beam position.
   4. Flow v-SUVs into the channel at a flow rate of 2 µl/min, corresponding to an average linear flow velocity of ~1.5 mm/sec for a cross section of 75 µm x 300 µm. Switch to the excitation/emission settings to monitor lipid mixing (LR or DiD only).
4. Observing fusion between single v-SUVs and the SBL

Figure 3. The experimental pTIRF setup. (A) Schematic representation of a v-SUV and a t-SBL on a glass substrate. False-color TIRFM images of single SUV-SBL fusion event showing lipid docking (1) and release of lipid dye into the SBL (2) followed by bleaching and decrease of the fluorescence intensity (3). The total fluorescence intensity (sum of pixel values in the 5.3 µm x 5.3 µm box) signal is shown. (B) The coverslip bonded to the PDMS block is taped onto the heated stage. Inlet tube for microfluidic channels draw samples from tube in metal sample holder (right) aspirated by the syringe pump (left). Beneath the pump is the dual emission unit. Please click here to view a larger version of this figure.

Figure 4. Schematic of the experimental setup. The evanescent wave is created at the glass-buffer interface in the microfluidic channel. The SBL is formed on the glass and v-SUV are aspirated from the metal sample holder (top right) through the channel into the syringe (top left). M, mirror; DM, dichroic mirror; L, lens; F, filter; P, polarizer. Please click here to view a larger version of this figure.

1. Continuously excite and monitor v-SUV fluorescence using a 561 nm (for LR-PE) or 638 nm (for DiD) laser depending on the fluorophore incorporated in v-SUVs. As v-SUVs reach the flow channel and dock onto and fuse with the SBL, the background fluorescence signal starts to accumulate.
2. Adjust the excitation laser intensity through the software to continuously bleach the background fluorescence such that at steady state new docking and fusion events can readily be observed.
   Note: If bleaching is too slow, background fluorescence will be too high. If bleaching is too fast, signals from docked SUVs and from fluorophores released into the SBL will fade rapidly, shortening the window during which fusion of docked SUVs can be detected or a fluorophore can be tracked. For LR-PE excited at 561 nm, 2.5-7.5 mW power illuminating a 190 µm diameter circle (100-250 nW/µm²) is a reasonable value to start. For 638 nm excitation of DiD, 0.8-1.6 mW power over a 190 µm diameter circle (30-60 nW/µm²) can be used for initial tests.
3. Acquire several movies at different positions in a given channel. Check NBD-PE fluorescence to verify that the SBL does not have defects at these positions. Acquire full-frame movies at the maximum rate (~50 frames/sec) or a cropped region of interest at higher frame rate (up to ~100 Hz) for 60 sec.
4. Move to another microfluidic channel and repeat the recordings for other conditions. Include negative controls such as a protein-free SBL or SUVs, add the soluble cytoplasmic domain of the v-SNARE VAMP2 (CDV) as an inhibitor or treat v-SUV with tetanus neurotoxin (30 min at 37 °C) in one or more of the channels on the same coverslip.

5. Clean up and PDMS recycling
   1. In order to reuse the PDMS block, rinse the microfluidic channels with ~200 µl of 70% ethanol at a flow rate of 5 µl/min. Finally, aspirate air through the tubing and channels.
   2. Detach the PDMS block gently from the coverslip by squeezing it slightly. Place it on a clean piece of aluminum foil. Store in a vacuum desiccator. It can be reused several times.
   3. For a more thorough cleaning rinse the channels with a detergent or sodium hydroxide solution prior to 70% ethanol. Alternatively, remove all tubes from the PDMS and sonicate the block for 30 min in isopropanol, before drying and inserting new tubing.

5. SUV-SBL Fusion Assay to Monitor Lipid and Content Release Simultaneously

1. For dual color monitoring of simultaneous lipid and soluble contents release, follow the same steps as in section 4, except use liposomes labeled with both a lipid (DiD) and a soluble contents (SRB) label, as explained in section 3. SRB is encapsulated at 10 mM and is initially highly self-quenched.
2. Using the configuration shown schematically in Figure 4, excite SRB and DiD fluorescence simultaneously using 561 nm and 638 nm lasers, respectively. A dichroic mirror (640 nm) splits the emission into two beams that run through a short (595/50 nm) and a long (700/75 nm) wavelength filter to detect the SRB and DiD emissions, respectively. The two emission beams are projected side-by-side onto the EM-CCD chip.

6. Data Analysis

1. Analysis of FRAP data
   1. Use the MATLAB program provided in the Supplementary Information to estimate the lipid diffusion coefficient, D. The program reads a list of OME-TIFF files⁴⁹, detects the bleached area, plots the mean pixel values in the bleached area as a function of time, and fits the resulting recovery curve to a model by Soumpasis⁵⁰ to extract the recovery time, , where w is the radius of the bleached circle.
      Note: A MATLAB program was provided earlier²⁵ for analysis of FRAP movies using Nikon ND2 files. The current program reads OME-TIFF files⁴⁹, because most file formats can readily be converted to OME-TIFF. A quantitative analysis of FRAP data is easiest and most accurate when bleaching is instantaneous, the bleached region is a circle, and bleaching during read-out is negligible. Although these criteria are not strictly satisfied in the simple FRAP measurements described here, a reasonable estimate of the diffusion coefficient can be obtained. For a more accurate estimate, use tracking of single lipid dyes (section 6.3).
2. Analysis of docking rate, fusion rate and docking-to-fusion delay times
   1. Use ImageJ to open a movie to analyze and adjust brightness and contrast to clearly identify vesicle docking and fusing events. Start the SpeckleTrackerJ²⁶ plugin. See Smith et al.²⁶ and online documentation for SpeckleTrackerJ for instructions for usage.
   2. Identify all newly docked SUVs in SpeckleTrackerJ. To distinguish SUVs that dock tightly from those that bounce off the SBL, impose a minimum docking duration of a few frames. Save the tracks, and repeat for all movies.
      Note: For the docking rate, all that matters is to identify when a SUV docked, so only the first frame in which the SUVs docked needs to be recorded in these tracks. The rest of the tracks will not be taken into account in the analysis. However, auto-tracking each SUV until it bleaches or fuses helps mark SUVs that have already been tracked.
   3. Identify all fusing vesicles. For these, the tracks should include all frames from the first frame a SUV docked until the first frame in which fusion is evident by a sudden increase in the fluorescence of the tracked spot. The durations of these tracks are used to calculate the docking-to-fusion delays. Save all tracks. Repeat for all movies.
   4. For batch analysis of docking or fusion data, compile a list of trajectory files from the relevant movies and run the MATLAB programs provided with Karatekin and Rothman²⁵. Follow instructions that are provided together with the programs.
      Note: The programs plot the cumulative docking and fusion events as a function of time, based on information extracted from the trajectory files. Docking and fusion rates are estimated from the slopes of these plots. For fusion data, the docking-to-fusion delays are also calculated and their distribution plotted as a survival plot, i.e., the probability that fusion has not yet occurred by a given delay after docking.
3. Lipid diffusivity
   1. For each fusion event, track single fluorescent lipids as they become discernible when they have diffused sufficiently away from the fusion site (Figure 6). Use SpeckleTrackerJ for tracking, and save the tracks for further analysis using MATLAB. Depending on the size of the fusing vesicle, typically 3-30 single fluorophores can be tracked. Because longer tracks are desirable for the calculation of  , try to track single molecules as long as possible, using manual correction of trajectories if necessary.
   2. Calculate the mean squared displacement (MSD) for single lipid marker trajectories that last > 40-50 frames (about ~1.5 sec). Use the MSD to calculate the lipid diffusion coefficient, . See Smith et al.²⁶ and Stratton et al.²⁷ for details.
4. Single lipid dye intensity, SUV-SBL intensity reduction factor, and vesicle size
   1. Measure the sum of pixel values of a lipid marker in a 3 x 3 pixel (0.8 µm x 0.8 µm) area around the marker for ~15 frames before and after the marker bleaches in a single step. Subtract the background intensity averaged over the post-bleach frames from the pre-bleach intensity averaged before bleaching to obtain the intensity of the tracked lipid marker. Repeat the measurement for as many markers as practical for a given movie.
   2. Plot the distribution of single lipid label intensities, , and fit a Gaussian to estimate the mean.
   3. Calculate the delay between when fusion occurred and when the trajectory of a single lipid marker released in that event ended in a single-step bleaching. Obtain the bleaching time for a fluorophore in the SBL, , by plotting the survivor function of the delays and fitting to an exponential.
      Note: Because the polarized excitation field couples more weakly to the fluorophores on a SUV, the bleaching time of a SUV is usually much slower²⁷.
   4. Estimate , the intensity reduction factor for a lipid dye when in the SUV relative to when it is in the SBL, following Stratton²⁷ ( is the intensity of a single dye in the SUV).
      Note: If bleaching were negligible,  would be equal to the docked intensity (point (1) in Figure 3A, right panel), divided by the total intensity reached after all fluorophores are deposited into the SBL upon fusion (dashed line labeled で Figure 3). However, due to rapid bleaching in the SBL, is typically not reached and an accurate estimate of requires fitting the release kinetics to an expression²⁷ that includes both (see 6.4.3) and . Separate expressions are provided in Stratton et al.²⁷ for the cases of pore-limited and diffusion limited release kinetics; the best fit case varies from event to event (see 6.5.1 for the case of pore-limited kinetics).
   5.  Calculate the vesicle area for individual events from²⁷ , where is the docked SUV intensity, is the mean intensity of a single lipid dye in the SBL, is the intensity reduction factor for a lipid dye when it is in the SUV relative to when it is in the SBL, and is the known areal density of lipid dyes.
5. Fusion pore properties 
   1. Assuming release is pore-limited, fit the lipid label release kinetics to ²⁷  , where is the docked SUV intensity just before fusion and the other parameters are as defined earlier. Use the value of  obtained in 6.4.3 as a fixed parameter, and extract the best fit estimates for  and .
   2. Estimate the fraction of time the pore is open, P₀, assuming retardation of lipid release is due to pore flickering²⁷:  = , where A_ves is the vesicle area (section 6.4.5), b is the pore height (typically taken to be ~15 nm), r_p ≈ 3 nm is the effective pore radius as seen by the diffusing lipid labels and includes half the bilayer thickness (~2 nm), is the lipid diffusivity (calculated in 6.3), and is the time for the lipids to be released from the SUV into the SBL (from 6.5.1).
   3. To confirm that a nominal P₀ > 1 indicates a fully open pore, P₀ = 1, fit the intensity time course to equation 4 of Stratton et al.²⁷, the predicted kinetics for a permanently open pore. This fit should be better than fitting the expression in 6.5.1 for a permanently open pore.