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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Freeze Substitution and Low Temperature Embedding System&#160; for Light and Electron Microscopy -AFS II</td>
			<td>Leica&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>High Pressure Freezer - EM-ICE</td>
			<td>Leica&#160;</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&#160;</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>CaCl2</td>
			<td>Sigma</td>
			<td>21115-250ML</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</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</td>
			<td>Thermo scientific</td>
			<td>10569-044</td>
			<td>Should be warmed at 37&#176;C before use</td>
		</tr>
		<tr>
			<td>FBS</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</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 at 37&#176;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&#160; (B-27)&#160;</td>
			<td>Fisher</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks 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&#160;</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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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

Nanyang Technological University

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

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Capture Compound Mass Spectrometry - A Powerful Tool to Identify Novel c-di-GMP Effector Proteins

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis (diguanylate cyclases) and degradation (phosphodiesterases) of the second messenger c-di-GMP. In contrast, little information is available regarding the molecular mechanisms and cellular components through which this signaling molecule regulates a diverse range of cellular processes. Most of the known effector proteins belong to the PilZ family or are degenerated diguanylate cyclases or phosphodiesterases that have given up on catalysis and have adopted effector function. Thus, to better define the cellular c-di-GMP network in a wide range of bacteria experimental methods are required to identify and validate novel effectors for which reliable in silico predictions fail.
We have recently developed a novel Capture Compound Mass Spectrometry (CCMS) based technology as a powerful tool to biochemically identify and characterize c-di-GMP binding proteins. This technique has previously been reported to be applicable to a wide range of organisms1. Here we give a detailed description of the protocol that we utilize to probe such signaling components. As an example, we use Pseudomonas aeruginosa, an opportunistic pathogen in which c-di-GMP plays a critical role in virulence and biofilm control. CCMS identified 74% (38/51) of the known or predicted components of the c-di-GMP network. This study explains the CCMS procedure in detail, and establishes it as a powerful and versatile tool to identify novel components involved in small molecule signaling.

The ubiquitous second messenger c-di-GMP controls growth and behavior of many bacteria. We have developed a novel Capture Compound Mass Spectrometry based technology to biochemically identify and characterize c-di-GMP binding proteins in virtually any bacterial species.

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis ...

Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused throughout tissues made of diverse cell types, such as enteroendocrine cells of the intestinal epithelium. These gastrointestinal sensors of food and bacteria have been difficult to study because they are dispersed among other epithelial cells at a ratio of 1:1,000. Recently, transgenic reporter mice have been generated to identify enteroendocrine cells by means of fluorescence. One of those is the peptide YY-GFP mouse. Using this mouse, we developed a method to correlate confocal and serial block-face scanning electron microscopy. We named the method cocem3D and applied it to identify a specific enteroendocrine cell in tissue and unveil the cell&#8217;s ultrastructure in 3D. The resolution of cocem3D is sufficient to identify organelles as small as secretory vesicles and to distinguish cell membranes for volume rendering. Cocem3D can be easily adapted to study the 3D ultrastructure of other specific cell types in their native tissue.

Here, we introduce a method, cocem3D, to unveil the ultrastructure of a specific cell in its native tissue by bridging confocal and serial block-face scanning electron microscopy.

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

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 ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...