Confocal Microscopy to Measure Three Modes of Fusion Pore Dynamics in Adrenal Chromaffin Cells

NOTE: The animal use procedure followed NIH guidelines and was approved by the NIH Animal Care and Use Committee.

1. Bovine chromaffin cell culture

1. Prepare Locke's solution (Table 1) and autoclave tools 1 day before chromaffin cell culture.
2. Obtain bovine adrenal glands from a local abattoir on the culture day, and keep them submerged in ice-cold Locke's solution before dissection.
   NOTE: Adrenal glands are from 21-27-month-old, healthy, black Angus of either sex (mainly male) with body weight around 1,400 pounds (~635 kg).
3. Prepare 30 mL (for 3 adrenal glands) of fresh enzyme solution containing collagenase P, trypsin inhibitor, and bovine serum albumin (Table 1) before dissection and keep it at room temperature.
4. Choose 3 intact glands without cuts or bleeds on the surface and remove the fat tissue with scissors (Figure 1A). Wash the glands by perfusion with Locke's solution until no blood comes out. To achieve this, inflate the gland through the adrenal vein (Figure 1B) using a 30 mL syringe attached with a 0.22 µm filter as many times as needed²⁵.
   NOTE: Approximately 150 mL of Locke's solution is usually needed to wash 3 glands.
5. For digestion, inject the enzyme solution through the adrenal vein using a 30 mL syringe attached with a 0.22 µm filter until the gland starts to swell. Then, leave the glands at 37 °C for 10 min. Inject once again and leave the glands at 37 °C for another 10 min.
   NOTE: Approximately 30 mL of enzyme solution is needed to digest 3 glands.
6. After digestion, cut the gland longitudinally from the vein to the other end with scissors to unfold the gland (Figure 1C). Isolate the medullae by tweezing out the white medulla into a 10 cm Petri dish containing Locke’s solution. 
   NOTE: The comparison of the interior of the gland before and after digestion is shown in Figure 1C. The details of the digestion and medullae isolation are reported previously^23,25.
7. Cut and mince the medulla into small pieces with scissors (Figure 1D). Filter the medulla suspension with an 80-100 µm nylon mesh into a beaker. Then transfer the filtrate to a 50 mL conical tube for centrifugation at 48 × g, room temperature, for 3 min with deceleration of 3.
   NOTE: The mincing of the medullae usually takes ~10 min. To obtain a good yield of cells, the minced pieces must be very small, until they cannot be tweezed up.
8. After centrifugation, remove the supernatant and resuspend the cell pellet with Locke's solution by pipetting. Filter the cell suspension with an 80-100 µm strainer, and centrifuge at 48 x g, room temperature, for 3 min with deceleration of 3.
9. Remove the supernatant and resuspend the cell pellet with 30 ml of culture medium (Table 1). Determine the cell number using hemacytometer counting chambers²⁵.

2. Transfection with electroporation

1. Transfer 2.8 × 10⁶ cells into a 15 mL tube. Pellet the cells by centrifugation at 48 × g for 2 min with deceleration of 3. Add 100 µL of transfection buffer (see the Table of Materials) provided by the manufacturer to the cell pellet, and then add 2 µg of the PH-mNG plasmid.
   NOTE: The PH-mNG plasmid was created by replacing the enhanced green fluorescent protein (EGFP) with mNG in PH-EGFP¹⁵ (see the Table of Materials).
2. Gently mix the suspension by pipetting the solution up and down, and transfer the mixture into an electroporation cuvette without delay (Figure 1E). Immediately transfer the cuvette to the electroporator (see the Table of Materials), select the O-005 program in screen list and press Enter to perform electroporation.
   NOTE: Prepare the cell suspension for transfection in a cell culture hood. Do not introduce air bubbles into the suspension during the mixing step. Proceed to the next step without delay.
3. After electroporation, add 1.8 mL of medium immediately to the cuvette and mix gently with a micropipettor equipped with a sterile tip. Then add 300 µL of the suspension of the electroporated cells on top of the coverslip (see the Table of Materials) in each dish, plating 5-6 dishes in total for one electroporation reaction.
4. Carefully transfer the dishes to a humidified incubator at 37 °C with 9% CO₂ for 30 min, and gently add 2 mL of prewarmed medium to each dish after 30 min.
5. Keep the transfected cells in the humidified incubator at 37 °C with 9% CO₂ for 2-3 days before experiment.
   ​NOTE: The cultured cells will last for a week. It is best to use cultured cells on days 2-3.

3. Preparation for patch-clamp recording and confocal imaging

NOTE: This protocol was performed with a laser scanning confocal microscope and patch-clamp amplifier with voltage-clamp recording together with a lock-in amplifier for capacitance recording. An XY plane confocal imaging at a fixed Z-plane (XY/Z_fixed scanning) was used to image all three fluorescent signals simultaneously. The Z-plane was focused at the cell bottom where the plasma membrane was adhering to the coverslips.

1. On the day of the patch-clamp and imaging experiment, observe the cells under a fluorescence microscope. Use brightfield to make sure the cell culture is not contaminated (Figure 1F) and epifluorescence to check for proper expression of the fluorescent-tagged protein.
   NOTE: For example, PH-mNG is expressed at the plasma membrane of ~20-30% of the cells.
2. Prepare patch pipettes from borosilicate glass capillaries. To do this, pull the pipettes with a pipette puller, coat their tips with liquid wax, and polish them with a microforge (see the Table of Materials).
3. Turn on the patch-clamp recording amplifier and start the patch-clamp recording software (see the Table of Materials). Set the appropriate parameters for calcium current and capacitance recording in the software (see the Table of Materials).
   1. Set a recording protocol of 60 s duration in total, where the stimulation starts at 10 s.
   2. Set the holding potential for voltage clamp recording to -80 mV. Set a 1 s depolarization from -80 mV to 10 mV as the stimulus to induce calcium influx and capacitance jump.
   3. For capacitance measurements, set the frequency of the sinusoidal stimulus to a range of 1,000-1,500 Hz with a peak-to-peak voltage of no more than 50 mV.
4. Save the protocol and create a new file for recording.
5. Turn on the confocal microscope system and set the appropriate parameters in the software (see the Table of Materials).
   1. Turn on the lasers, including 458 nm, 514 nm, and 633 nm, and set the emission collection range for each laser according to each fluorescence probe as in the following. FFN511: excitation wavelength (EX), 458 nm; emission wavelength (EM), 468-500 nm. mNG: EX, 514 nm; EM, 524-560 nm. A655: EX, 633 nm; EM, 650-700 nm.
   2. Use sequential imaging for FFN511 and mNG to avoid crosstalk between these two probes. Set a timelapse of 1 min duration for image recording (Figure 1G).

4. Patch-clamp recording and confocal imaging

1. Choose a dish with good cell state and proper expression and add 2 µL of fluorescent false neurotransmitter FFN511 (10 mM stock, 1:1,000 working solution) into the medium. Put the dish back in the incubator for 20 min. Alternatively, load FFN511 after step 3.2 and perform steps 3.3-3.5 while waiting to save time.
2. After FFN511 loading finished, prepare the recording chamber and add 2 µL of fluorescent dye A655 into 500 µL of the bath solution (Figure 2A and Table 1). Transfer the coverslip from the dish into the recording chamber (see the Table of Materials) with tweezers, and immediately add A655-containing bath solution (Figure 2B).
   NOTE: The A655 is kept in -20 °C with concentration of 10 mM and the working concentration is 40 µM.
   CAUTION: Wear gloves to avoid direct skin contact with FFN511 or A655.
3. Place a drop of oil (refractive index: 1.518; see the Table of Materials) on the 100x oil immersion objective (Numerical Aperture = 1.4). Mount the chamber in the microscope and use the adjustment knob to make the oil just contact the bottom of the coverslip, then immerse the ground wire tip into the bath solution (Figure 2C, D).
   NOTE: Choose an immersion oil appropriate for working at room temperature. Switching between the low-and high-magnification lens is unnecessary. The room temperature is maintained around 20-22 °C during recording.
4. Bring the cells into focus and use brightfield and confocal imaging to find a good cell with mNG expression. Zoom in on the selected cell and adjust it to the center of the view to minimize blank regions.
   NOTE: The schematic drawing of fluorescence labeling is shown in Figure 3A. A good cell usually has a smooth membrane and clean edge (Figure 3B). With epifluorescence or confocal imaging, a good cell appears to have a large and flat bottom with bright mNG expression (Figure 3C).
   The pixel size is ~50 nm, within a range of ~40-80 nm according to different cell size and zoom factor.
5. Set up parameters for XY plane confocal imaging at a fixed Z-plane (XY/Z_fixed scanning) of FFN511, PH-mNG, and A655, with a minimized time interval. Adjust the focus to the bottom of the cell with the fine adjustment knob.
   NOTE: PH-mNG signals the contour of the cell plasma membrane at the confocal XY-plane crosssection (except the cell bottom). Near the cell membrane bottom, A655 spots can be observed, which reflect Ω-shape or Λ-shape membrane invaginations. If the Z-focal plane is lower than the cell bottom, the intensity of all fluorescence will become weaker.
6. Adjust the excitation laser power in the software to find a setting to get the best signal-to-noise ratio and avoid significant fluorescence bleaching. To do so, start with an initial test value of 2.5 mW power for FFN511 excited at 458 nm and 1 mW for mNG excited at 514 nm. For A655 excited at 633 nm with an HeNe laser, use high laser power, ~12-15 mW (i.e., 70%-80% of the maximum), which, upon continuous excitation, can bleach all fluorescent A655 inside an Ω-shape profile when its pore is closed.
   NOTE: If the laser power is too high, PH-mNG and FFN511 fluorescence will be bleached quickly. If the laser power is too low, signals will be too weak or noisy to be analyzed.
7. Add 9 µL of internal solution (Table 1) into a patch pipette and attach the pipette to the holder in the patch-clamp amplifier stage (see the Table of Materials).
8. Apply a small amount of positive pressure with a syringe and move the pipette tip to touch the bath solution with a micromanipulator. Ensure the amplifier shows that the pipette resistance is ~2-4 MΩ with a voltage pulse test. Press LJ/Auto to cancel the liquid junction.
9. Move the pipette toward the selected cell with the micromanipulator. To form a cell-attached mode, move the pipette tip to touch the cell (the resistance will increase); applying negative pressure gently with the syringe (resistance will increase to more than 1 GΩ), change the holding potential from 0 to -80 mV.
10. Once resistance passes 1 GΩ, wait for ~30 s for the configuration to stabilize. Press C-fast/Auto to compensate for fast capacitance.
11. To form a whole-cell mode, apply pulsed short yet powerful negative pressure with the syringe to rupture the membrane (the shape of the current pulse changes with a charged and discharged membrane capacitor). Press C-slow/Auto to compensate for slow capacitance.
12. Adjust the imaging focus slightly to focus on the cell bottom. Start confocal timelapse imaging and patch-clamp recording simultaneously by clicking the Start buttons in the two different software applications at the same time.
    NOTE: The confocal imaging software makes a movie at a rate of ~20-90 ms/frame. The patch-clamp recording includes the resting state for 10 s, the 1 s depolarization stimulation, and an additional 49 s after stimulation. The patch-clamp configuration permits recording of calcium current, capacitance jump, and decay induced by the 1 s depolarization from -80 to +10 mV (Figure 3D).
13. Once recording is finished, make sure the data are saved. Change the holding potential back to 0 mV. Move the pipette out of the bath solution and discard it.
14. Repeat steps 4.7-4.13 to record another cell in the dish. After 1 h of recording, change to another dish for recording.
    ​NOTE: After 1 h of recording, the success rate of patch-clamp recording decreases significantly. To increase the efficiency of data collection, recording for only 1 h is recommended in each dish.

5. Patch-clamp data analysis

1. Open appropriate software (see the Table of Materials) for data analysis. Click the PPT | Load PULSE file | File buttons to load the .dat file. Click Do it and four traces will be plotted in one graph automatically including the calcium current and capacitance traces.
2. Click the Windows | New Graph buttons, choose the Pulse_1_1_1, the first wave in Y Wave(s), and click Do it to plot the calcium current graph. Click the Windows | New Table buttons, choose the Pulse_1_1_1, and click Do it to show the raw data of calcium current, which could be used to plot the averaged trace of multiple cells.
3. Draw a square accordingly in the calcium current graph, right click and expand the current signal to include the baseline and the peak of calcium current. Click Graph | Show Info to show cursors A and B and move them to the baseline and peak respectively. The graph information will be shown in the bottom and parameters can be estimated.
4. Repeat step 5.2, but choose the Pulse_1_1_1_Cm, the second wave in Y Wave(s), to plot the capacitance trace. Repeat step 5.3 and place A and B cursors in the appropriate position to measure the capacitance parameters, such as baseline, amplitude, and decay rate.
5. Copy raw data in step 5.2 into a spreadsheet, calculate the mean and standard error of mean for a group of recorded cells, and plot the averaged traces of calcium current and membrane capacitance (Figure 3E) in an appropriate software.

6. Confocal imaging data analysis

1. Open the raw imaging files with any manufacturer-supplied software (see the Table of Materials).
   NOTE: Some other free programs, such as ImageJ or Fiji, could be utilized for data processing and quantifying the images obtained in section 4.
2. Click Process | ProcessTools and use the tools to generate rolling average (e.g., rolling average for every 4 images) files for each timelapse image and save those files.
   NOTE: After rolling average, appropriate adjustment of the brightness and background could be done to show the fluorescence changes of three channels better, which may help to identify fusion events. Check the fluorescent intensity in some regions without fusion event may help to distinguish the fluorescent signal of fusion events from background signals.
3. Click the buttons Quantify | Tools | Stack Profile, check timepoints before and after stimulation to identify fluorescence changes in each channel. Click the Draw ellipse button to circle the regions of interest (ROIs) for fusion events. Right click on the image and click Save ROIs to save the ROIs.
   NOTE: The size of ROI, which covers all the three fluorescent signals, was similar with vesicle size in chromaffin cells²⁴. Raw data were used for the analysis, while the rolling averaging data which increase the signal-to-noise ratio were provided to show clearly the three signals.
4. Click Open projects to locate the raw file, right click on the image and click Load ROIs to load the ROI file in the raw imaging files to measure the fluorescence intensities.
   NOTE: The raw fluorescent signal will be used to plot traces of different channels. For each ROI, the baseline is defined as the intensity before stimulation, and the intensity traces can be normalized to baseline.
5. Click Tools | Sort ROIs in the software and plot the traces for all three channels of each ROI. Click Report to save the ROI data, including both digital data and imaging traces for each ROI, into a file folder.
   1. Identify a fusion event by the simultaneous increase in the intensity of PH-mNG (F_PH) and A655 (F₆₅₅) spot fluorescence (within a single frame), accompanied by a decrease in FFN511 spot fluorescence (F_FFN).
   2. Look for the appearance of F_PH and F₆₅₅, which indicates PH-mNG/A655 spot formation due to an Ω-profile generated by fusion, allowing the diffusion of PH-mNG from the plasma membrane and persistent diffusion of A655 from the bath.
   3. Look for F_FFN decrease, which indicates its release from a vesicle due to fusion pore opening and excludes the possibility that F_PH and F₆₅₅ appearance may be from membrane invagination caused by endocytosis.
   4. Inspect the timelapse XY/Z_fixed images that show fusion events happening on the cell bottom. Analyze three modes of fusion events: 1) close-fusion, 2) stay-fusion, and 3) shrink-fusion.
      NOTE: Fusion events were rarely observed before depolarization, whereas tens of fusion events could be observed after depolarization. Following fusion, the Ω-profile may close its pore, maintain its open pore, or shrink to merge into the plasma membrane.
      1. To identify close-fusion, look for dimming F₆₅₅ because fusion pore closure prevents the exchange of bath A655, while F_PH sustains, reflecting the continuing conversion of PtdIns(4,5)P₂ into PtdIns(4)P, or F_PH decays with a delay, reflecting vesicle pinch-off (close-fusion, Figure 4A,B).
      2. Look for sustained F_PH and F₆₅₅ to identify stay-fusion (Figure 4C).
      3. Look for parallel decreases of F_PH and F₆₅₅, indicating Ω-profile shrinkage to identify shrink fusion (Figure 4D).
         NOTE: Check the original imaging files to inspect the imaging traces if uncertain of the event type. Checking the fluorescent intensity in some regions without fusion event may help to distinguish the fluorescent signal of fusion events from background signals.
6. Plot representative traces of intensity changes for these three channels: FFN511, PH-mNG, and A655. Quantify the percentage of different fusion modes in each cell and plot figures.