This work was supported by funding from the Johns Hopkins University (SW). We thank the Johns Hopkins School of Medicine Microscope Facility for their technical support. We thank Erik Jorgensen and Christian Rosenmund and members of their laboratories for the development of the technique. We thank M. Wayne Davis for the design of the initial device. We thank Paul Wurzinger, Cveta Tomova, and Delgermaa Luvsanjav for their technical assistance. We also thank Natalie R. Hamilton and Grant F. Kusick for their critical reading of the manuscript.

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
<ol>
	<li>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.</li>
	<li>Treat the cortices with 800 &#181;L of 0.05% Trypsin-EDTA for 15 min at 37 &#176;C to dissociate the astrocytes.</li>
	<li>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 &#176;C and 5% CO2.</li>
	<li>Place one acid-washed and sterilized 18 mm glass coverslip per well of a 12-well plate.</li>
	<li>Briefly wash two 6 mm carbon-coated sapphire disks in 70% ethanol and place them on top of each glass coverslip.</li>
	<li>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 &#181;L of PDL solution to the sapphire disks and glass coverslips for 5 min at RT.</li>
	<li>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.</li>
	<li>Seed the astrocytes from step 1.3 in 2 mL of DMEM at a density of 5x104 cells/well in the plate as prepared in steps 1.4 - 1.6. Grow them at 37 &#176;C and 5% CO2 for one week.</li>
	<li>Add 20 &#181;L of fluoro-deoxyuridine (final concentration: 80 &#181;M) to each well for at least a few h before culturing the neurons. 
	NOTE: Fluoro-deoxyuridine stops astrocyte division.</li>
	<li>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.</li>
	<li>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 &#181;m filter.</li>
	<li>Harvest the brain from a P0 - 2 mouse18. Immediately transfer the brain to ice-cold Hanks&#39;-balanced Salt Solution (HBSS). Dissect the hippocampi under a stereomicroscope, keeping the tissue submerged in HBSS.</li>
	<li>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 &#176;C and 750 rpm.</li>
	<li>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 &#176;C. Aspirate off the inactivating solution.</li>
	<li>Add 200 &#181;L of neuronal basal medium (see the Table of Materials) to the isolated hippocampus. Triturate using a 200 &#181;L pipette tip to dissociate the cells. Wait until the undissociated cells settle at the bottom. Carefully remove the medium with cells from the top.</li>
	<li>Repeat step 1.15 3x. Pool all the dissociated cells in a new 1.5 mL centrifuge tube.</li>
	<li>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.</li>
	<li>Infect the neurons with lentivirus expressing channelrhodopsin at DIV 3 (3 d&#160;in vitro) 18.</li>
	<li>Perform flash-and-freeze on DIV 14, as described in steps 2.1 - 2.5, below.</li>
</ol>
2. Flash-and-Freeze
<ol>
	<li>Preparation of Fixative
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Preparation of Physiological Saline
	<ol>
		<li>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 &#181;m. 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.</li>
		<li>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.</li>
		<li>Check the osmolarity using an osmometer. Ensure that it is 300 &#177; 5 mOsm.</li>
		<li>Add AMPA receptor antagonist (NBQX) to a final concentration of 3 &#181;M and GABA receptor antagonist (bicuculline) to a final concentration of 30 &#181;M. 
		NOTE: Neurotransmitter receptor antagonists are added to avoid recurrent network activity following neuronal stimulation18.</li>
		<li>Warm up the physiological saline to 37 &#176;C for use.</li>
	</ol>
	</li>
	<li>Preparation of the Specialized High-pressure Freezer and an Automated Freeze Substitution Unit
	<ol>
		<li>Prior to high-pressure freezing, cool down an automated freeze substitution unit to -90 &#176;C by filling the tank with liquid nitrogen.</li>
		<li>Cool down acetone in a small cup to -90 &#176;C by placing it inside the specimen chamber.</li>
		<li>Fill the liquid nitrogen dewar and storage dewar of the high-pressure freezer (see the Table of Materials) with liquid nitrogen.</li>
		<li>Set up the light stimulation protocol using the touch screen monitor.
		<ol>
			<li>Name the program by clicking on &#34;Edit&#34; next to &#34;Program name&#34; on the light stimulation window. Another window will pop up.</li>
			<li>Set up a program by typing &#34;15,000 ms&#34; in &#34;dark phase,&#34; &#34;100 ms&#34; in &#34;period,&#34; &#34;10 ms&#34; in &#34;pulse,&#34; and &#34;1&#34; in &#34;number of periods&#34; for a single stimulus of 10 ms. Freeze the cells 90 ms later (Figure 1C). 
			NOTE: The &#34;dark phase&#34; allows the cells to recover from light exposure during sample loading. &#34;Period&#34; defines the stimulation frequency. For example, if the stimulus should be applied at 20 Hz, this column should be set at 50 ms. &#34;Pulse&#34; defines the duration of the light stimulus. Finally, the &#34;number of periods&#34; defines the total number of stimuli applied. For setting up a high-frequency stimulation, please see Figure 1E.</li>
		</ol>
		</li>
		<li>For a &#34;no stimulation&#34; control, type &#34;15 s&#34; in &#34;dark phase,&#34; &#34;0 ms&#34; in &#34;period,&#34; &#34;1 ms&#34; in &#34;pulse,&#34; and &#34;0&#34; in &#34;number of periods&#34; in the light stimulation setup window, as described in step 2.3.4. 
		NOTE: By default, the &#34;pulse&#34; must be at least 1 ms.</li>
		<li>On the main screen, make sure that the box for light stimulation is checked.</li>
		<li>Set up the storage protocol by clicking &#34;Specimen Storage&#34; on the main screen. Click on &#34;Edit.&#34; In the following window, use &#34;+&#34; or &#34;-,&#34; to select &#34;2&#34; to store 2 disks in each channel (3 channels in total). Check &#34;Storage LN2 enabled.&#34;</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>Place one sapphire disk, cell-side facing up, in the well of the black, middle plate (Figure 1B).</li>
		<li>Place a 100 &#181;m spacer ring over the sapphire disk (Figure 1B).</li>
		<li>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.</li>
		<li>Place another 100 &#181;m spacer ring and a 400 &#181;m spacer ring (Figure 1B). Remove the extra liquid using filter paper.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Repeat steps 2.4.1 - 2.4.6 for each specimen.</li>
		<li>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 6th sample, step 2.5 must be performed. Once unloaded, repeat steps 2.4.1 - 2.4.6.</li>
	</ol>
	</li>
	<li>Sample Collection and Transfer to an Automated Freeze-substitution Unit
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#176;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.</li>
	</ol>
	</li>
</ol>
3. Freeze Substitution in the Automated Freeze-substitution Unit
<ol>
	<li>Using precooled tweezers (tips at ~-196 &#176;C), quickly transfer the middle plate from step 2.5.3 to the precooled acetone (-90 &#176;C).</li>
	<li>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 &#176;C) for a few minutes. Gentle tapping with tweezers also helps to dissociate the sapphire from the middle plate.</li>
	<li>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.</li>
	<li>Set up the freeze substitution program as follows: (i) -90 &#176;C for 5 - 30 h, (ii) -90 - -20 &#176;C in 14 h (5 &#176;C/h), (iii) -20 &#176;C for 12 h, and (iv) -20 &#176;C - 20 &#176;C in 4 h (10 &#176;C/h). 
	NOTE: The duration of the first step at -90 &#176;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.</li>
</ol>
4. Infiltration and Plastic Embedding with Epoxy Resin
<ol>
	<li>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.</li>
	<li>Using a pipette, add acetone (room temperature) to each cryogenic vial and wash each sapphire disk 4 - 6x for 1 - 2 h.</li>
	<li>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.</li>
	<li>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 &#181;L of benzyl dimethylamine (BDMA) while mixing. De-gas for 10 min.</li>
	<li>Prepare 30, 70, and 90% epoxy resin in acetone from the 100% epoxy resin prepared in step 4.4.</li>
	<li>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.</li>
	<li>Replace the 30% epoxy resin with 70% epoxy resin by pipetting and incubate for 3 - 4 h at RT in an orbital shaker.</li>
	<li>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 &#176;C.</li>
	<li>The next day, make fresh epoxy resin medium, as in step 4.4.</li>
	<li>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.</li>
	<li>Place the samples in a 60 &#176;C oven for 48 h to polymerize.</li>
</ol>
5. Mounting Samples
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Mount the small piece of plastic (~2&#160;x 2&#160;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 &#176;C for 1 h.</li>
</ol>
6. Sectioning
<ol>
	<li>Use an ultramicrotome to section the sample.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Collect ribbons of sections on single-slot copper grids covered with 0.7% polyvinyl acetate (see the Table of Materials).</li>
</ol>
7. Imaging Using a Transmission Electron Microscope (TEM)
<ol>
	<li>Prior to TEM imaging, stain the section with 2.5% uranyl acetate in methanol for 5 min.</li>
	<li>Wash the grid 15x in 50% methanol. Then, wash in ultrapure water 15x, with each wash lasting 30 s.</li>
	<li>Briefly air-dry the section and place the grid into a specimen holder of the TEM.</li>
	<li>Image at 93,000X magnification.</li>
	<li>Acquire images. 
	NOTE: Imaging is typically done blind to the time points or genotypes, and typically ~200 images are collected/time point.</li>
</ol>
8. Image Analysis
<ol>
	<li>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.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetic+approaches+to+cell+migration+and+beyond.">Optogenetic approaches to cell migration and beyond.</a> Curr Opin Cell Biol. 30, 112-120 (2014).</li><li>Niu, J., Ben Johny, ., Dick, M., E, I., Inoue, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Following+Optogenetic+Dimerizers+and+Quantitative+Prospects.">Following Optogenetic Dimerizers and Quantitative Prospects.</a> Biophys J. 111 (6), 1132-1140 (2016).</li><li>van Bergeijk, P., Adrian, M., Hoogenraad, C. C., Kapitein, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetic+control+of+organelle+transport+and+positioning.">Optogenetic control of organelle transport and positioning.</a> Nature. 518 (7537), 111-114 (2015).</li><li>Idevall-Hagren, O., Dickson, E. J., Hille, B., Toomre, D. K., De Camilli, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetic+control+of+phosphoinositide+metabolism.">Optogenetic control of phosphoinositide metabolism.</a> Proc Natl Acad of Sci U.S.A. 109 (35), E2316-E2323 (2012).</li><li>Kramer, R. H., Fortin, D. L., Trauner, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+photochemical+tools+for+controlling+neuronal+activity.">New photochemical tools for controlling neuronal activity.</a> Curr Opin Neurobiol. 19 (5), 544-552 (2009).</li><li>Frank, J. A., Franquelim, H. G., Schwille, P., Trauner, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+Control+of+Lipid+Rafts+with+Photoswitchable+Ceramides.">Optical Control of Lipid Rafts with Photoswitchable Ceramides.</a> J Am Chem Soc. 138 (39), 12981-12986 (2016).</li><li>Rosenmund, C., Clements, J. D., Westbrook, G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonuniform+probability+of+glutamate+release+at+a+hippocampal+synapse.">Nonuniform probability of glutamate release at a hippocampal synapse.</a> Science. 262 (5134), 754-757 (1993).</li><li>Denk, W., Horstmann, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Serial+block-face+scanning+electron+microscopy+to+reconstruct+three-dimensional+tissue+nanostructure.">Serial block-face scanning electron microscopy to reconstruct three-dimensional tissue nanostructure.</a> PLoS Biology. 2 (11), e329 (2004).</li><li>Knott, G., Marchman, H., Wall, D., Lich, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Serial+section+scanning+electron+microscopy+of+adult+brain+tissue+using+focused+ion+beam+milling.">Serial section scanning electron microscopy of adult brain tissue using focused ion beam milling.</a> J Neurosci. 28 (12), 2959-2964 (2008).</li><li>Watanabe, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clathrin+regenerates+synaptic+vesicles+from+endosomes.">Clathrin regenerates synaptic vesicles from endosomes.</a> Nature. 515 (7526), 228-233 (2014).</li></ol>

Open Access publication of&#160;this article was sponsored&#160;by Leica Mikrosysteme GmbH.

The &#34;flash-and-freeze&#34; approach visualizes membrane dynamics by inducing a particular cellular event with optogenetics and by freezing cells at defined time points after stimulation19. In this demonstration, we used channelrhodopsin, a light-sensitive cation channel, to stimulate neurons and captured the fusion and recovery of synaptic vesicles at the synaptic terminals. In recent years, many optogenetic tools have been developed22,23, all of which are compatible with flash-and-freeze. For example, organelle trafficking can be induced using light-induced heterodimerization of cryptochrome and CIB124. Similarly, the lipid composition of the plasma membrane can be altered by the light-induced translocation of phosphoinositide phosphatases to the plasma membrane25. Furthermore, small, light-sensitive compounds like azobenzene change conformation depending on the illumination wavelengths. This conformational change can be used to activate ligand-gated channels or to change lipid composition in the membrane26,27. Caged compounds can also be used to induce cellular activity. However, the LED used in the current setup may not produce sufficient energy for uncaging; thus, further optimizations of the system are likely necessary. Nevertheless, the applications of these light-activatable tools are flexible-many cellular events can be induced by a flash of light. &#34;Flash-and-freeze&#34; can capture the resulting membrane dynamics.
There are two main limitations to the &#34;flash-and-freeze&#34; method. First, it captures &#34;snapshots&#34; of a particular event from different cells. In other words, it is not possible to follow membrane dynamics in one cell over a period of time. Thus, for the reconstruction of any cellular event, one must acquire and analyze a large number of images from each sample and at each time point. Furthermore, in neurons, an even larger number of images is necessary, since the fusion of synaptic vesicles only takes places in 20 - 30% of the synapses in mouse hippocampal neurons18,28. The analysis of such a large dataset requires tremendous amounts of time. In the future, image acquisition and analysis need to be automated to make the approach more efficient29,30.
The second limitation is imposed by the nature of the high-pressure freezing technique. When cells freeze, cellular water rearranges to form ice crystals if the freezing rate is below 100 K/s21. These ice crystals can penetrate membranes or concentrate solutes to alter local osmotic pressure, resulting in the rupture of membranes. To avoid ice crystals, high pressure (~2,000 atm) is applied to the specimens. Due to the super-cooling effect, a freezing rate of 100 K/s is sufficient to prevent water from forming ice crystals at this pressure21. In theory, specimens as thick as 500 &#181;m can be frozen without ice crystals, but approximately 200 &#181;m is likely the practical limit, as cuboidal forms of ice tend to form in thick tissue, compromising morphology. When processing specimens thicker than 5 &#181;m, the use of a proper cryo-protectant, such as BSA, is necessary. However, BSA will change the osmolarity of the solution and may affect the physiological response of cells. Therefore, extensive control experiments are required to validate the use of BSA in particular systems. Ice crystals can also form after high-pressure freezing if the specimens are accidently removed from the liquid nitrogen bath. Thus, it is critical to keep the specimens in the liquid nitrogen at all times and to use pre-cooled forceps to manipulate them.
When planning experiments, the following three points should be considered. First, the maximal intensity of light (the 460 nm line) is 5.5 - 8.0 mW/mm2. Whether this intensity is sufficient to induce activity must be verified with live-cell imaging on a fluorescence microscope prior to flash-and-freeze experiments. Second, experiments must be performed at physiological temperature. The stage of the high-pressure freezer is warmed up to 37 &#176;C for the experiments with mouse hippocampal neurons31. Finally, the time points must be carefully chosen to capture the membrane dynamics. Initial studies indicated that endocytosis is complete after 100 ms of stimulation. Thus, three additional time points (15 , 30, &#38; 50 ms) were also examined to follow the membrane dynamics17,18. These time points were necessary to visualize membrane trafficking events during synaptic transmission. However, the requirement for the number of time points are different in each cellular event. Therefore, a few time points should be sampled before initiating large dataset collection.

Visualizing membrane and protein dynamics within a cell is a key step towards understanding the cell biology of particular processes. Dynamic trafficking events can be captured using light or fluorescence microscopy. However, the subcellular context is largely missing in such images because subcellular structures cannot be completely &#34;painted&#34; by dyes or fluorescent probes and resolved spatially and spectrally1,2. On the other hand, while electron microscopy can delineate subcellular architecture in exquisite detail, it cannot capture cellular dynamics, because specimens must be fixed prior to imaging. Thus, it is typically not sufficient to completely understand cellular dynamics using only one imaging modality.
To overcome the limitations of light and electron microscopy, correlative microscopy techniques have been developed. Correlative Light and Electron Microscopy (CLEM) visualizes intracellular dynamics using light microscopy and underlying subcellular structures with electron microscopy. In CLEM, cells engaged in various processes, such as cytokinesis and endocytosis3,4,5,6, are live-imaged and then processed for electron microscopy. Although CLEM captures certain aspects of intracellular dynamics, there are four factors that limit the utility of this approach. First, the temporal resolution is limited by how quickly the cells can be immobilized, which typically takes s - min due to the slow diffusion and reaction of fixatives7. Second, the subcellular architecture is observed post facto8; thus, the dynamic morphological changes cannot be captured using this approach. Third, the fluorescence and electron micrographs cannot be precisely aligned due to tissue shrinkage caused by dehydration during the sample preparation for electron microscopy9,10. Fourth, events like cytokinesis and endocytosis do not take place at the same time in every cell5,11, and thus, a particular cell that is engaged in the event must be identified from a large population of cells. This process is often laborious. Thus, a new method is necessary to induce particular events in every cell and to capture the resulting cellular dynamics by the rapid immobilization of cells at defined time points.
Recently, several tools have been developed to induce particular cellular dynamics using light (optogenetics). Channelrhodopsin is a light-sensitive, non-selective cation channel isolated from Chlamydomonas reinhardtii12,13. When channelrhodopsin is expressed in neuronal membranes, a brief flash of light induces an influx of sodium ions into neurons and triggers an action potential14,15. The action potential then propagates into the synaptic terminals, where synaptic vesicles fuse within milliseconds16,17,18 Therefore, channelrhodopsin induces neuronal activity. To follow membrane dynamics at synaptic terminals, neurons must be immobilized at defined time points after stimulation with ms precision.
To capture membrane dynamics after inducing neuronal activity, we coupled light stimulation with high-pressure freezing17,18,19. High-pressure freezing allows for the near-instantaneous immobilization of cells with reduced ice crystal formation20. Ice crystals can rupture membranes and disrupt the subcellular architecture21. By varying the time intervals between stimulation and freezing, membrane trafficking within synaptic terminals was captured following the induction of an action potential.
Here, we demonstrate experimental procedures using a commercialized high-pressure freezer that couples a ms temporal control of light stimulation with high-pressure freezing. Unlike other instruments that require an external device to control light stimulation and freezing, light stimulation is fully integrated in this system and can be applied with ms precision19. This process involves multiple steps. 1) Mouse hippocampal neurons are cultured on sapphire disks and infected with lentivirus carrying an expression vector for channelrhodopsin18. 2) Neurons are stimulated and frozen at defined time points after stimulation. 3) The vitrified water is substituted with an organic solvent, while lipids and proteins are cross-linked by fixatives to preserve the intracellular architecture. 4) The samples are infiltrated and embedded in epoxy resin. 5) Ultrathin sections are collected using an ultramicrotome. 6) Thin sections are imaged on a transmission electron microscope. 7) Image acquisition and analysis are performed blindly with respect to time points or genotypes. Cellular dynamics can be determined through the reconstruction of time-resolved images17,18. Sample preparation (steps 2 - 5 above) requires a week, but the subsequent image analysis requires months to a year.

<table><tbody><tr><td>Freeze-substitution and Low-temperature Embedding System&amp;nbsp; for Light and Electron Microscopy -AFS II</td><td>Leica&amp;nbsp;</td><td></td><td></td></tr><tr><td>High Pressure Freezer - EM-ICE</td><td>Leica&amp;nbsp;</td><td></td><td>EM ICE is a specialized high pressure freezer that allows precise control of light stimulation and freezing.</td></tr><tr><td>Osmometer</td><td>Gonotec</td><td></td><td></td></tr><tr><td>Ultramicrotome UC7</td><td>Leica&amp;nbsp;</td><td></td><td></td></tr><tr><td>Oven</td><td>Blue M</td><td></td><td></td></tr><tr><td>Razor blade</td><td>Personna</td><td></td><td></td></tr><tr><td>Glutaraldehyde</td><td>EMS</td><td>16530</td><td></td></tr><tr><td>Osmium tetroxide</td><td>EMS</td><td>RT19132</td><td>Toxic, open only under certified chemical hood</td></tr><tr><td>Acetone</td><td>EMS</td><td>RT10016</td><td></td></tr><tr><td>HEPES</td><td>Emdmillipore</td><td>391340-250GM</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>49159-1KG</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>746436-1KG</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653-1KG</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>21115-250ML</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>J.T.Baker</td><td>2444-01</td><td></td></tr><tr><td>Liquid epoxy resin Eponate 12</td><td>Ted Pella</td><td>18028</td><td></td></tr><tr><td>Bisphenol A epoxy resin Araldite 502</td><td>Ted Pella</td><td>18028</td><td></td></tr><tr><td>Dodecenyl succinic anhydride (DDSA)</td><td>Ted Pella</td><td>18028</td><td></td></tr><tr><td>Benzyl dimethyl amine (BDMA)</td><td>Ted Pella</td><td>18241</td><td></td></tr><tr><td>Special embedding (BEEM) capsule</td><td>EMS</td><td>70021</td><td></td></tr><tr><td>Copper Grid</td><td>Ted Pella</td><td>1GC12H</td><td></td></tr><tr><td>Polyvinyl acetate (Pioloform F)</td><td>Ted Pella</td><td>19244</td><td></td></tr><tr><td>Uranyl acetate</td><td>Polysciences</td><td>21447-25</td><td></td></tr><tr><td>Ethyl cyanoacrylate (Super glue)</td><td>Scotch</td><td>170497</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Themo scientific</td><td>25300-120</td><td></td></tr><tr><td>DMEM (Dulbecco&#039;s Modified Eagle&#039;s Medium)</td><td>Thermo scientific</td><td>10569-044</td><td>Should be warmed to 37 &amp;deg;C before use</td></tr><tr><td>FBS (Fetal Bovine Serum)</td><td>Thermo scientific</td><td>26140-079</td><td></td></tr><tr><td>Pen-Strep</td><td>Thermo scientific</td><td>15140-122</td><td></td></tr><tr><td>Fluoro-deoxyuridine (FUDR)</td><td>Sigma</td><td>F0503</td><td></td></tr><tr><td>Glass cover slip</td><td>Fisher</td><td>S175223</td><td>Should be acid-washed</td></tr><tr><td>Sapphire disc</td><td>Technotrade</td><td>616-100</td><td></td></tr><tr><td>Acetic acid</td><td>Emdmillipore</td><td>1000631011</td><td></td></tr><tr><td>Poly-D-Lysine (PDL)</td><td>Sigma</td><td>P6407</td><td></td></tr><tr><td>Rat tail collagen</td><td>Thermo scientific</td><td>A10483-01</td><td></td></tr><tr><td>Neurobasal A</td><td>Fisher</td><td>10888022</td><td>Should be warmed to 37 &amp;deg;C before use</td></tr><tr><td>L-alanyl-L-glutamine (Glutamax)</td><td>Fisher</td><td>35050-079</td><td></td></tr><tr><td>Seraum free supplement&amp;nbsp; (B-27)&amp;nbsp;</td><td>Fisher</td><td>17504044</td><td></td></tr><tr><td>Hanks&#039;-balanced Salt Solution (HBSS)</td><td>Thermo scientific</td><td>24020117</td><td></td></tr><tr><td>Papain</td><td>Worthington</td><td>LS003126</td><td>Active Unit should be calculated for each batch</td></tr><tr><td>Thermomixer</td><td>Eppendorf&amp;nbsp;</td><td>Thermomixer C</td><td></td></tr><tr><td>Trypsin inhibitor</td><td>Sigma</td><td>T9253</td><td></td></tr><tr><td>NBQX</td><td>Tocris</td><td>03-731-0</td><td></td></tr><tr><td>Bicculine</td><td>Tocris</td><td>01-091-0</td><td></td></tr><tr><td>Whatman I filter paper</td><td>GE</td><td></td><td></td></tr><tr><td>Transmission Electron Microscope</td><td>Philips CM20</td><td></td><td></td></tr></tbody></table>

flash-and-freeze: a novel technique to capture membrane dynamics with electron microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

Using the protocol described above, we performed &#34;flash-and-freeze&#34; experiments in mouse hippocampal neurons expressing channelrhodopsin. These neurons were frozen either 15 ms or 100 ms after light onset. We have previously shown that the exocytosis and endocytosis of synaptic vesicles occur in the synaptic terminals at the 15 ms and 100 ms time points, respectively18. These events were successfully captured at the appropriate times (Figure 2), suggesting that flash-and-freeze experiments can be successfully performed on the chosen specialized high-pressure freezer (see the Table of Materials).
<img alt="Figure 1" src="/files/ftp_upload/55664/55664fig1.jpg" /> 
Figure 1. Sample Loading and Programming in the High-pressure Freezer. A) Sample loading table of a high-pressure freezer. The middle plate, shown in the inset for structural detail, is placed in a CLEM holder for sample loading. One of the half-cylinders is placed at the bottom part of the sample loading table, and the other is attached with a clip to the top cover. Once the sample is loaded, the middle plate is pushed forward to the bottom half-cylinder and the cover is closed to initiate the freezing. B) Specimen assembly. The sapphire disk containing neurons is placed in the well of the middle plate, with the cell-side facing up. A 100 &#181;m ring is placed directly above the sapphire disk inside the well. Then, an empty sapphire disk dipped in physiological saline is placed with the solution-side down. Air bubbles must be avoided. Finally, a 100 &#181;m ring and a 400 &#181;m ring are snugly placed above. Any extra liquid is removed with filter paper. C) A cross-section of an embedding capsule with the sapphire disk submerged in epoxy resin. The sapphire disk is placed at the bottom of the capsule, with the cell-side facing up and covered with epoxy resin for infiltration and embedding. D) Programming the high-pressure freezer for a single, 10 ms stimulus. The specimens are frozen 90 ms after the light pulse. E) Programming the high-pressure freezer for 10 stimuli at 20 Hz. The specimens are frozen 5 ms after the last light pulse. <a href="http://cloudflare.jove.com/files/ftp_upload/55664/55664fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55664/55664fig2.jpg" /> 
Figure 2. Visualization of Exocytosis and Endocytosis in Mouse Hippocampal Neurons. Hippocampal neurons are stimulated once and frozen at the indicated times. Electron micrographs show the exocytosis of a synaptic vesicle A) and ultrafast endocytosis B). PSD, post-synaptic Density. <a href="http://cloudflare.jove.com/files/ftp_upload/55664/55664fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy at JoVE.com

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

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

flash-freeze-novel-technique-to-capture-membrane-dynamics-with

Research

JoVE Journal

Biology

13.9K Views.  Johns Hopkins School of Medicine. We developed a novel technique in electron microscopy, "flash-and-freeze," that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

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

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Biochemistry

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

Bioengineering

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

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

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

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

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular communication. EVs consist of three populations of vesicles, namely microvesicles (MVs), exosomes, and apoptotic bodies. The limiting membrane of EVs is crucially involved in the interactions with the recipient cells, which could lead to the transfer of biologically active molecules to the recipient cells and, consequently, affect their behavior. The freeze-fracture electron microscopy technique is used to study the internal organization of biological membranes. Here, we present a protocol for MV isolation from cultured cancerous urothelial cells and the freeze-fracture of MVs in the steps of rapid freezing, fracturing, making and cleaning the replicas, and analyzing them with transmission electron microscopy. The results show that the protocol for isolation yields a homogenous population of EVs, which correspond to the shape and size of MVs. Intramembrane particles are found mainly in the protoplasmic face of the limiting membrane. Hence, freeze-fracture is the technique of choice to characterize the MVs' diameter, shape, and distribution of membrane proteins. The presented protocol is applicable to other populations of EVs.

We present a protocol for the isolation and freeze-fracturing of extracellular vesicles (EVs) originating from cancerous urothelial cells. The freeze-fracture technique revealed the EVs' diameter and shape and-as a unique feature-the internal organization of the EV membranes. These are of immense importance in understanding how EVs interact with the recipient membranes.

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular ...

Cancer Research

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

Immunology and Infection

Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated to generate a carbon/platinum &#8220;cast&#8221; intended for examination by transmission electron microscopy. Specimens are subjected to ultrarapid freezing rates, often in the presence of cryoprotective agents to limit ice crystal formation, with subsequent fracturing of the specimen at liquid nitrogen cooled temperatures under high vacuum. The resultant fractured surface is replicated and stabilized by evaporation of carbon and platinum from an angle that confers surface three-dimensional detail to the cast. This technique has proved particularly enlightening for the investigation of cell membranes and their specializations and has contributed considerably to the understanding of cellular form to related cell function. In this report, we survey the instrument requirements and technical protocol for performing freeze-fracture, the associated nomenclature and characteristics of fracture planes, variations on the conventional procedure, and criteria for interpretation of freeze-fracture images. This technique has been widely used for ultrastructural investigation in many areas of cell biology and holds promise as an emerging imaging technique for molecular, nanotechnology, and materials science studies.

Basic techniques and refinements of freeze-fracture processing of biological specimens and nanomaterials for examination by transmission electron microscopy are described. This technique is a preferred method for revealing ultrastructural features and specializations of biological membranes and for obtaining ultrastructural level dimensional and spatial data in materials sciences and nanotechnology products.

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated ...

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize macromolecular complexes at very high resolution. However, to get as close as possible to the native state, perfect sample preservation is required. Conventional electron microscopy (EM) fixation with aldehydes, for instance, does not provide good ultrastructural preservation. The slow penetration of fixatives induces cell reorganization and loss of various cell components. Therefore, conventional EM fixation does not allow for an instantaneous stabilization and preservation of structures and antigenicity. The best choice for examining intracellular events is to use cryofixation followed by the freeze-substitution fixation method that keeps cells in their native state. High-pressure freezing/freeze-substitution, which preserves the integrity of cellular ultrastructure, is the most commonly used method, but requires expensive equipment. Here, an easy-to-use and low-cost freeze fixation method followed by freeze-substitution for suspension cell cultures is presented.

This manuscript describes an easy-to-use and low-cost cryofixation method for visualizing suspension cells by transmission electron microscopy.

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...