Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

All of the experiments were performed according to the rules and regulations of animal use by the National Institutes of Health. The protocol was approved by the Animal Care and Use Committee (IACUC) at Johns Hopkins School of Medicine.

1. Isolation and Culture of Mouse Hippocampal Neurons

1. Dissect cortices from a postnatal day 0 – day 2 (P0 – 2) mouse brain¹⁸. Isolate the astrocytes from the cortices.
   NOTE: The mouse brain was isolated after decapitation. Astrocytes serve as a feeder layer for hippocampal neurons.
2. Treat the cortices with 800 µL of 0.05% Trypsin-EDTA for 15 min at 37 °C to dissociate the astrocytes.
3. Culture the astrocytes in a T-75 flask with 13 mL of DMEM containing 10% fetal bovine serum (FBS) and 0.2% penicillin-streptomycin for one week at 37 °C and 5% CO₂.
4. Place one acid-washed and sterilized 18 mm glass coverslip per well of a 12-well plate.
5. Briefly wash two 6 mm carbon-coated sapphire disks in 70% ethanol and place them on top of each glass coverslip.
6. Prepare Poly-D-Lysine (PDL) solution by mixing 3 mL of 17 nM acetic acid with 1 mL of rat tail collagen and 1 mL of PDL (1 mg/mL). Apply 200 µL of PDL solution to the sapphire disks and glass coverslips for 5 min at RT.
7. Remove the PDL solution and air dry. Prior to use, sterilize the plate as prepared in steps 1.4 – 1.6 for 30 min under ultraviolet light.
8. Seed the astrocytes from step 1.3 in 2 mL of DMEM at a density of 5×10⁴ cells/well in the plate as prepared in steps 1.4 – 1.6. Grow them at 37 °C and 5% CO₂ for one week.
9. Add 20 µL of fluoro-deoxyuridine (final concentration: 80 µM) to each well for at least a few h before culturing the neurons.
   NOTE: Fluoro-deoxyuridine stops astrocyte division.
10. Change the medium to 1.5 mL of neuronal basal medium (see the Table of Materials) containing 1% L-alanyl-L-glutamine (see the Table of Materials), 2% serum-free supplement for neuronal cells (see the Table of Materials), and 0.2% penicillin-streptomycin.
11. Prepare papain solution by adding 20 units of papain to 5 mL of enzyme solution (1.65 mM Cysteine, 1 mM CaCl₂, and 0.5 mM EDTA in DMEM). Acidify the solution by passing CO₂ gas for 20 min. Filter-sterilize with a 0.22 µm filter.
12. Harvest the brain from a P0 – 2 mouse¹⁸. Immediately transfer the brain to ice-cold Hanks'-balanced Salt Solution (HBSS). Dissect the hippocampi under a stereomicroscope, keeping the tissue submerged in HBSS.
13. Place the two hippocampi in 1 mL of the papain solution prepared in step 1.11. Incubate for 1 h on a thermomixer at 37 °C and 750 rpm.
14. Replace the papain with 1 mL of inactivating solution containing 2.5 mg of trypsin inhibitor and 0.5 mg of albumin per mL of DMEM. Incubate for 5 min at 37 °C. Aspirate off the inactivating solution.
15. Add 200 µL of neuronal basal medium (see the Table of Materials) to the isolated hippocampus. Triturate using a 200 µL pipette tip to dissociate the cells. Wait until the undissociated cells settle at the bottom. Carefully remove the medium with cells from the top.
16. Repeat step 1.15 3x. Pool all the dissociated cells in a new 1.5 mL centrifuge tube.
17. Count the number of cells using a hemocytometer. Plate neurons at a density of 6.5 x 10⁴ cells/well on top of the astrocyte layer prepared in steps 1.1 – 1.10.
18. Infect the neurons with lentivirus expressing channelrhodopsin at DIV 3 (3 d in vitro) ¹⁸.
19. Perform flash-and-freeze on DIV 14, as described in steps 2.1 – 2.5, below.

2. Flash-and-Freeze

1. Preparation of Fixative
   1. Under a chemical hood, add the following substances to a conical tube to prepare the fixative: 2.5 mL of glutaraldehyde (10% stock in acetone), 0.25 g of osmium tetroxide, 0.25 mL of water, and 22.25 mL of anhydrous acetone.
      NOTE: The final concentration of each component should be 1% in acetone. Glutaraldehyde is added to preserve the protein structures during freeze-substitution. CAUTION: The acute toxicity of osmium tetroxide is high. Exposure to vapors could damage the cornea of the eye. It should be handled only in a certified chemical hood.
   2. Aliquot 1 mL of fixative into numbered cryogenic vials (2 mL). Keep the fixative frozen in liquid nitrogen until use.
      NOTE: Osmium tetroxide and glutaraldehyde cross-react and precipitate; thus, once mixed, aliquot immediately, cap the tubes, and submerge the cryotubes in liquid nitrogen to freeze the fixative. Use a pencil to number the cryogenic vials, since acetone can wash off markers.
2. Preparation of Physiological Saline
   1. Make physiological saline solution by mixing Hepes (10 mM, pH 7.5), NaCl (140 mM), KCl (2.4 mM), and glucose (10 mM).
      NOTE: These values are final concentrations. Cryo-protectants are not used for monolayer cultures. However, a proper cryo-protectant is required for specimens thicker than 5 µm. The use of 20% BSA is typically recommended. Yeast paste and E. coli OP50 can also be used as cryo-protectants for fly larvae or C. elegans.
   2. Add CaCl₂ at 4 mM and MgCl₂ at 1 mM final concentrations.
      NOTE: The concentrations of CaCl₂ and MgCl₂ vary depending on experiments. To ensure the capture of exocytic intermediates, 4 mM calcium is used for these particular experiments to increase the release probability of vesicles¹⁸.
   3. Check the osmolarity using an osmometer. Ensure that it is 300 ± 5 mOsm.
   4. Add AMPA receptor antagonist (NBQX) to a final concentration of 3 µM and GABA receptor antagonist (bicuculline) to a final concentration of 30 µM.
      NOTE: Neurotransmitter receptor antagonists are added to avoid recurrent network activity following neuronal stimulation¹⁸.
   5. Warm up the physiological saline to 37 °C for use.
3. Preparation of the Specialized High-pressure Freezer and an Automated Freeze Substitution Unit
   1. Prior to high-pressure freezing, cool down an automated freeze substitution unit to -90 °C by filling the tank with liquid nitrogen.
   2. Cool down acetone in a small cup to -90 °C by placing it inside the specimen chamber.
   3. Fill the liquid nitrogen dewar and storage dewar of the high-pressure freezer (see the Table of Materials) with liquid nitrogen.
   4. Set up the light stimulation protocol using the touch screen monitor.
      1. Name the program by clicking on "Edit" next to "Program name" on the light stimulation window. Another window will pop up.
      2. Set up a program by typing "15,000 ms" in "dark phase," "100 ms" in "period," "10 ms" in "pulse," and "1" in "number of periods" for a single stimulus of 10 ms. Freeze the cells 90 ms later (Figure 1C).
         NOTE: The "dark phase" allows the cells to recover from light exposure during sample loading. "Period" defines the stimulation frequency. For example, if the stimulus should be applied at 20 Hz, this column should be set at 50 ms. "Pulse" defines the duration of the light stimulus. Finally, the "number of periods" defines the total number of stimuli applied. For setting up a high-frequency stimulation, please see Figure 1E.
   5. For a "no stimulation" control, type "15 s" in "dark phase," "0 ms" in "period," "1 ms" in "pulse," and "0" in "number of periods" in the light stimulation setup window, as described in step 2.3.4.
      NOTE: By default, the "pulse" must be at least 1 ms.
   6. On the main screen, make sure that the box for light stimulation is checked.
   7. Set up the storage protocol by clicking "Specimen Storage" on the main screen. Click on "Edit." In the following window, use "+" or "-," to select "2" to store 2 disks in each channel (3 channels in total). Check "Storage LN2 enabled."
4. Sample Loading and Freezing in the High-pressure Freezer
   NOTE: All the sample assembly and loading steps are done under a stereomicroscope with a 7.5-60X magnification range. A tweezer is used in steps 2.4.1 – 2.4.4 to manipulate the specimens. Experiments must be performed at physiological temperature.
   1. Place one sapphire disk, cell-side facing up, in the well of the black, middle plate (Figure 1B).
   2. Place a 100 µm spacer ring over the sapphire disk (Figure 1B).
   3. Place a blank sapphire disk over the spacer ring (Figure 1B) after dipping one side of the disk in the pre-warmed saline solution from step 2.2. Make sure that no air bubbles are trapped between the two sapphire disks.
   4. Place another 100 µm spacer ring and a 400 µm spacer ring (Figure 1B). Remove the extra liquid using filter paper.
   5. Place the assembly from step 2.4.3 between the two transparent half-cylinders (Figure 1A). Close the top red cover to initiate the freezing process.
      NOTE: The preset protocol runs automatically once the cover is closed. The sample stays in the same orientation in the freezing chamber, and a flash of light is applied from the top of the sample assembly. The red cover pops back up automatically once the freezing process is completed.
   6. Store the specimen in the storage dewar.
      NOTE: After freezing, the specimen is automatically dropped into a storage dewar filled with liquid nitrogen and stored there until further processing. The storage dewar has three chambers, and each chamber can hold up to 3 samples at most. Typically, two specimens are frozen under the same stimulation conditions, and the high-pressure freezer is programmed to store both in the same chamber.
   7. Repeat steps 2.4.1 – 2.4.6 for each specimen.
   8. Proceed to step 2.5 once all the chambers are full.
      NOTE: The device was set to store up to 6 specimens in the storage dewar at a time. Therefore, after every 6^th sample, step 2.5 must be performed. Once unloaded, repeat steps 2.4.1 – 2.4.6.
5. Sample Collection and Transfer to an Automated Freeze-substitution Unit
   1. Open the door to the storage dewar, which is located under the table of the high-pressure freezer. Remove the storage dewar and place it on the benchtop.
   2. Using hands, remove the specimen chamber from the dewar and transfer it in the specialized sample tray filled with liquid nitrogen. Unlock the knob to release the specimen cup.
   3. Remove the transparent half-cylinders from the specimen cup using a pair of tweezers after pre-cooling the tips of the tweezers with liquid nitrogen (~-196 °C). Carefully transfer the middle, black plate to a small cup containing liquid nitrogen.
      NOTE: The tips of the tweezers must be precooled to the temperature of liquid nitrogen. The sample must be kept under liquid nitrogen at all times to prevent ice crystal formation.

3. Freeze Substitution in the Automated Freeze-substitution Unit

1. Using precooled tweezers (tips at ~-196 °C), quickly transfer the middle plate from step 2.5.3 to the precooled acetone (-90 °C).
2. Separate the sapphire disk from the middle plate by gently shaking or tapping.
   NOTE: Occasionally, it may be difficult to separate the sapphire disk from the middle plate. In such a situation, leave the middle plate in precooled acetone (-90 °C) for a few minutes. Gentle tapping with tweezers also helps to dissociate the sapphire from the middle plate.
3. Place a cryogenic vial containing fixatives (step 2.1) inside a specimen chamber of the substitution unit. Transfer the sapphire disk to the cryogenic vial and place a cap on the vial.
4. Set up the freeze substitution program as follows: (i) -90 °C for 5 – 30 h, (ii) -90 – -20 °C in 14 h (5 °C/h), (iii) -20 °C for 12 h, and (iv) -20 °C – 20 °C in 4 h (10 °C/h).
   NOTE: The duration of the first step at -90 °C could be varied. The total duration of freeze substitution is set in such a way that the program ends in the morning (~1.5 d postexperiment) around 8 am so that the subsequent steps can be performed during the daytime.

4. Infiltration and Plastic Embedding with Epoxy Resin

1. Once the program ends, use gloved hands to transfer the cryogenic vials containing the sapphire disks from the specimen chamber of the substitution unit to a chemical hood.
2. Using a pipette, add acetone (room temperature) to each cryogenic vial and wash each sapphire disk 4 – 6x for 1 – 2 h.
3. Optionally, incubate the specimens in 0.1% uranyl acetate for 1 h if extra contrast is needed. Wash 4 – 6x with acetone over 1 – 2 h.
   NOTE: CAUTION: There is risk associated with internal exposure following the inhalation of uranyl acetate, which causes irritation of the upper respiratory tract. High exposure can damage blood cells. Work with uranyl acetate should be done under exhaust ventilation. Protective clothing is recommended.
4. Prepare liquid epoxy resin medium by weighing 6.2 g of glycerol polyglycidyl ether, 4.4 g of bisphenol-A epoxy resin, and 12.2 g of dodecenyl succinic anhydride (DDSA). Mix thoroughly and add 800 µL of benzyl dimethylamine (BDMA) while mixing. De-gas for 10 min.
5. Prepare 30, 70, and 90% epoxy resin in acetone from the 100% epoxy resin prepared in step 4.4.
6. Add 30% epoxy resin to the cryogenic vials containing sapphire disks and incubate for 2 – 3 h at RT in an orbital shaker at 120 rpm.
7. Replace the 30% epoxy resin with 70% epoxy resin by pipetting and incubate for 3 – 4 h at RT in an orbital shaker.
8. Using tweezers, transfer each sapphire disk, cell-side-up, to the cap of an embedding capsule. Add 90% epoxy resin to the cap. Incubate O/N at 4 °C.
9. The next day, make fresh epoxy resin medium, as in step 4.4.
10. Transfer each sapphire disk, cell-side facing up, to the cap of an embedding capsule. Fill the cap with freshly prepared 100% epoxy resin. Change to fresh 100% epoxy resin every 2 h, repeating three times.
11. Place the samples in a 60 °C oven for 48 h to polymerize.

5. Mounting Samples

1. Place the sample upside-down on the stereomicroscope so that the sapphire disk is located at the top of the resin block. Remove the thin layer of epoxy resin from the top by scratching it with a razor blade.
2. Using a razor blade, cut a shallow line along the edge of the sapphire disk; this line helps to separate the sapphire disk from the epoxy resin in step 5.3.
3. Separate the sapphire disk from the epoxy resin by dipping it into liquid nitrogen for approximately 10 s.
   NOTE: The cells will stay in the epoxy resin.
4. After removing the sapphire disk, use a dissecting scope to find an area with cells (4-10X magnification). Cut around the region of interest (~2 x 2 mm) using a razor blade (double-edged). To keep the specimen in place, perform this step while the specimen is taped to the surface of the microscope.
5. Mount the small piece of plastic (~2 x 2 x 5 mm) containing the cells using glue containing ethyl cyanoacrylate on a cylindrical dummy block made of epoxy resin. Incubate the block at 60 °C for 1 h.

6. Sectioning

1. Use an ultramicrotome to section the sample.
   1. Trim the surface of the plastic-embedded specimen with a glass knife at a speed of 3 mm/s and a thickness of 200 nm/section. Cut 4 – 5 sections from the surface where the astrocytes are located.
   2. Switch to a diamond knife and section at a speed of 0.8 mm/s and a thickness of 40 nm/section. Cut 20 – 25 sections.
2. Collect ribbons of sections on single-slot copper grids covered with 0.7% polyvinyl acetate (see the Table of Materials).

7. Imaging Using a Transmission Electron Microscope (TEM)

1. Prior to TEM imaging, stain the section with 2.5% uranyl acetate in methanol for 5 min.
2. Wash the grid 15x in 50% methanol. Then, wash in ultrapure water 15x, with each wash lasting 30 s.
3. Briefly air-dry the section and place the grid into a specimen holder of the TEM.
4. Image at 93,000X magnification.
5. Acquire images.
   NOTE: Imaging is typically done blind to the time points or genotypes, and typically ~200 images are collected/time point.

8. Image Analysis

1. Analyze the images using a software (e.g., ImageJ) with a custom macro (Watanabe, Davis, and Jorgensen, unpublished).
   NOTE: The x/y coordinates are recorded from vesicles, the plasma membrane, the active zone membrane, and all other membrane-bound organelles at synapses. The text files containing the information are exported to another software (e.g., Matlab). The data are further analyzed using custom programs.