SBEM imaging, light microscopy imaging and electron microscopy sample preparation were performed with the use of the equipment of Laboratory of Imaging Tissue and Function which serves as an imaging core facility at the Nencki Institute of Experimental Biology.
For preparation of Figure 1, the image of a mouse (Souris_02), and a vial from the https://smart.servier.com/ was used.
This work was supported by the National Science Centre (Poland) Grant Opus (UMO-2018/31/B/NZ4/01603) awarded to KR.

The research was performed in compliance with Nencki Institute guidelines and permission of the Local Ethical Committee. The studies were carried out in accordance with the European Communities Council Directive of 24 November 1986 (86/609/EEC), Animal Protection Act of Poland and approved by the first Local Ethics Committee in Warsaw. All efforts were made to minimize the number of animals used and their suffering.
CAUTION: All procedures described below must be carried out in a laboratory fume hood. Due to the hazardous nature of the reagents used. Personal safety measures such as gloves, lab coat, safety glasses and a face mask are required.
1. Preparation of the fixative for perfusion (2% wt/vol paraformaldehyde (PFA) and 0.5% vol/vol glutaraldehyde (GA) in 0.1 M phosphate buffer (PB), pH 7.4)
NOTE: Prepare the fixative solution on the same day as it will be used and do not store it longer than for 3 hours. In case of time shortage, prepare 2% PFA in 0.1 M PB the day before, store it at 4 &#176;C and add fresh GA shortly before the perfusion.
<ol>
	<li>Take 400 mL of sterile double distilled water (ddH2O) and heat it to 60 &#176;C using a stirring hot plate. Then add 20 g of PFA. Add drops of 1 M NaOH until PFA is fully dissolved and allow the mixture to cool down.</li>
	<li>Add 500 mL of 0.2 M PB (pH 7.4).</li>
	<li>Filter the solution to remove any deposit and cool it to 4 &#176;C.</li>
	<li>Just before perfusion, add 20 mL of 25% GA to the solution and then top up the volume with ddH2O to 1 L.</li>
</ol>
2. Preparation of postperfusion fixative for SBEM (2% wt/vol PFA and 2.5% vol/vol GA in 0.1 M PB, pH 7.4)
<ol>
	<li>Take 50 mL of the fixative for perfusion (2% PFA and 0.5% GA in 0.1 M PB).</li>
	<li>Add 5 mL of 25% GA.</li>
</ol>
3. Preparation of postperfusion fixative for IF staining (4% PFA in phosphate buffered saline (PBS))
<ol>
	<li>Dissolve a tablet of 1x PBS (pH 7.4) in a purified and deionized water (H2O) according to manufacturer instructions.</li>
	<li>Use a stirring hot plate to heat the solution to 60 &#176;C and add 40 g of PFA.</li>
	<li>Add drops of 1 M NaOH until the PFA is fully dissolved and allow the mixture to cool.</li>
	<li>Adjust the pH of the solution to 7.5 with 1 M HCl then top up the volume with H2O to 1 L.</li>
	<li>Filter the solution to remove any deposits.</li>
</ol>
4. Transcardial perfusion of animals
NOTE: All PFA and GA wastes must be collected and stored for disposal according to the local regulations. Anesthesia and perfusion should follow the local regulations. In the described protocol adult 3-month-old and 20&#177;1-month-old female Thy1-GFP(M) mice (Thy1-GFP +/-)12 expressing green fluorescence protein (GFP) in a sparsely distributed population of glutamatergic neurons were used but any other can be used as well. Animals were bred as heterozygotes with the C57BL/6J background in the Animal House of the Nencki Institute of Experimental Biology.
<ol>
	<li>Before perfusion anesthetize a mouse by administration of a ketamine/xylazine mixture (up to 90 mg/kg body weight ketamine and 10 mg/kg body weight xylazine) via intraperitoneal injection (27-gauge needle).
	<ol>
		<li>Assess whether the depth of anesthesia is sufficient by checking the reflex to pain stimuli (pinching) and the corneal reflex (squinting).</li>
	</ol>
	</li>
	<li>After 20 minutes perform an intraperitoneal injection (27-gauge needle) of sodium pentobarbital (50 mg/kg body weight).</li>
	<li>Perfuse a mouse according to a perfusion surgery protocol described by Gage et al.13 (see point 4; Figure 5-6). Using a perfusion pump start with a 0.1 M phosphate buffer, pH 7.4 (30 mL) for 3 minutes and continue with 2% PFA and 0.5% GA in 0.1 M PB, pH 7.4 for 6 minutes (80 mL).
	<ol>
		<li>Gently dissect the brain from the skull and divide it in half (see Figure 9-10 in Gage et al., 201213). Place one piece in a vial containing fixative for SBEM and the second into the vial with 4% PFA/PBS for IF staining.</li>
		<li>Keep the hemispheres in the fixative at 4 &#176;C overnight.</li>
	</ol>
	</li>
</ol>
5. Brain slices preparation for electron microscopy
<ol>
	<li>Choose the vibratome settings (blade travel speed: 0.075 mm/s, cutting frequency: 80 Hz).</li>
	<li>Place the slice chamber into the holder, attach it to the vibratome and surround it with ice. Then place a razor blade into the vibratome blade holder.</li>
	<li>Use a spoon, or similar object and place the chilled brain (dorsal surface up) on a tough cutting surface (e.g., a glass Petri dish lid). To prepare a coronal slice of the hippocampus make a perpendicular cut between the cerebral hemisphere and the cerebellum with a razor blade or scalpel, thus removing the cerebellum. The olfactory bulb can also be removed.</li>
	<li>Apply cyanoacrylate glue on the dry platform of the vibratome.</li>
	<li>Pick up the brain with forceps and carefully dry it on filter paper.</li>
	<li>Glue the hemisphere to the platform close to the cutting blade with the rostral tip upwards. Attach the platform to the holder and immediately fill it up with ice-cold 0.1 M PB, pH 7.4. If the perpendicular cut is properly made, the hemisphere will stand straight up providing 90&#176; angle required to make a symmetric coronal cut containing the hippocampus. Assure that the brain is covered with PB.</li>
	<li>Position the vibratome blade in front of the hemisphere and lower it to the coronal side of the hemisphere. Lower the blade to 400 &#181;m further in the caudal direction and start slicing. Continue slicing until the first two slices are completely separated from the tissue block.</li>
	<li>Retract the blade and lower another 100 &#181;m, then slice again.</li>
	<li>When the hippocampus becomes visible (use the mouse brain atlas Paxinos and Franklin, 200414) collect the slices with the small paintbrush or widened plastic Pasteur pipette.</li>
	<li>Transfer the slices into a 12-well plate filled with cold 0.1 M PB, pH 7.4.</li>
	<li>Dissect the hippocampus in a glass Petri dish filled with 0.1 M PB, pH 7.4 using a razor blade or scalpel, and put it into glass vials with the same phosphate buffer. 
	&#8203;NOTE: For long-term storage of the slices supplement 0.1 M PB with 0.05% sodium azide (NaN3).</li>
</ol>
6. Brain sample preparation for immunostaining
<ol>
	<li>After overnight fixation in 4% PFA/PBS solution in a fume hood, put the brain tissue into the cryopreservation solution (30% sucrose in PBS with 0.05% NaN3) and keep it at 4 &#176;C for 2 days (until sunk).</li>
	<li>Prepare an antifreeze solution (15% sucrose/30% ethylene glycol/0.05% NaN3/PBS).</li>
	<li>Set the cabinet temperature of the cryostat at -19 &#176;C and make sure that the temperature is reached before proceeding further. During sectioning, ensure that the cabinet temperature remains between -18 &#176;C and -20 &#176;C.</li>
	<li>Use a spoon, or similar object and place the chilled brain (dorsal surface up) on a tough cutting surface (e.g., a glass Petri dish lid). To prepare a coronal slice of the hippocampus make a perpendicular cut between the cerebral hemisphere and the cerebellum with a razor blade or scalpel, thus removing the cerebellum.</li>
	<li>Select a pre-cooled specimen disc, cover it with a medium for freezing on a freeing shelf and using forceps fix the hemisphere to the disc with a rostral tip upwards and wait until the specimen is completely frozen. Insert the specimen disc into a specimen head.</li>
	<li>Install a blade in a blade holder inside the cryo-chamber and cut 40 &#181;m thick slices.</li>
	<li>Transfer slices with a small paintbrush to the 96-well plate filled with cold anti-freeze solution and gently unroll the sections (collect a slice after each round of slicing to prevent them from getting lost in the slice chamber). 
	&#8203;NOTE: Brains or sections can be stored in an antifreeze solution at -20 &#176;C for a long time.</li>
</ol>
7. Immunostaining of brain slices
NOTE: All staining steps were performed in a 24-well plate on a platform shaker.
<ol>
	<li>Wash the slices with PBS three times, each time for 6 minutes.</li>
	<li>Incubate slices in 300 &#181;L of blocking solution (5% normal donkey serum (NDS)/0.3% Triton X-100) for 1 hour with gentle shaking on a rotator.</li>
	<li>Incubate the slices with the primary antibody against PSD-95 diluted 1:500 in 5% NDS/0.3% Triton X-100/PBS (300 &#181;L per well) overnight at 4 &#176;C. The final concentration of the primary antibody is 2 &#181;g/mL.</li>
	<li>Wash the slices with 0.3% Triton X-100/PBS at room temperature (RT) three times, each time for 6 minutes.</li>
	<li>Incubate slices with the secondary antibody diluted 1:500 in 300 &#181;L of 0.3% Triton X-100/PBS for 90 minutes. The final concentration of the secondary antibody is 4 &#181;g /mL.</li>
	<li>Wash the slices with PBS three times, each time for 6 minutes.</li>
	<li>Mount the slices on slides using a mounting medium and then close them with a cover slide.</li>
	<li>Examine the specimens using a confocal microscope (63 &#215; oil objective, NA 1.4, pixel size 0.13 &#181;m &#215; 0.13 &#181;m). 
	&#8203;NOTE: For long-term storage: keep the samples at 4 &#176;C, protect them from light.</li>
</ol>
8. SBEM sample preparation
CAUTION: Due to the hazardous nature of reagents used all the procedures described below must be carried out in a laboratory fume hood. Before using these chemicals read carefully the Material Safety Data Sheets provided by the manufacturers and ask the safety officer about the local rules to ensure safe handling and waste disposal.
<ol>
	<li>Sample contrasting 
	NOTE: Slices washings and room temperature incubation should be carried out with mild shaking (e.g., on a platform shaker). Autoclave degassed water was used.
	<ol>
		<li>Wash the slices with cold 0.1 M PB, pH 7.4 five times, each time for 3 minutes.</li>
		<li>Prepare a 1:1 mixture of 4% aqueous osmium tetroxide and 3% potassium ferrocyanide (1:1 by vol). The final product will turn brown. Immerse the samples in this mixture, place them on ice, and from this stage onwards it is important to protect them from light. Incubate them with gentle shaking for 1 hour.</li>
		<li>Meanwhile prepare a thiocarbohydrazide (TCH) solution. Mix 10 mL of double-distilled H2O (ddH2O) and 0.1 g of TCH and place it into an oven set at 60 &#176;C for 1 hour. It is important to swirl the solution from time to time (e.g., every 10 minutes). When ready, cool it to room temperature.</li>
		<li>Wash the slices with ddH2O five times, each time for 3 minutes.</li>
		<li>Filter TCH solution using a 0.22 &#181;m syringe filter and immerse the sample in filtered solution. The slices will turn black. Incubate them for 20 minutes at room temperature.</li>
		<li>Wash the samples with ddH2O five times, each time for 3 minutes.</li>
		<li>Incubate the samples with a 2% aqueous solution of OsO4 for 30 minutes at room temperature.</li>
		<li>Wash the samples with ddH2O five times, each time for 3 minutes.</li>
		<li>Place the samples in filtered 1% aqueous uranyl acetate and incubate them at 4 &#176;C overnight. Use 0.22 &#181;m syringe filter for filtration.</li>
		<li>Prepare L-aspartic acid solution by mixing 0.4 g of L-aspartic acid and 80 mL of ddH2O, adjust the pH to 3.8 for easier dissolving, and then top up with water to 100 mL.</li>
		<li>Next day, begin with Walton&#39;s lead aspartate15 preparation. Mix 0.066 g of lead nitrate with 10 mL of L-aspartic acid solution (point 8.1.10) pre-warmed to 60 &#176;C and adjust pH to 5.5 (measured at 60 &#176;C) with 1 M NaOH. Close the vial with lead aspartate and leave it at 60 &#176;C for 30 minutes in a water bath. The solution should be clear. If it turns cloudy, it must be discarded and a new one needs to be prepared.</li>
		<li>In the meantime, wash the samples with degassed ddH2O five times, each time for 3 minutes. Then keep them in the oven set at 60 &#176;C for 30 minutes.</li>
		<li>Immerse the samples in the freshly prepared lead aspartate solution and incubate them in the oven set at 60 &#176;C for 20 minutes.</li>
		<li>Wash the samples with degassed ddH2O five times, each time for 3 minutes.</li>
	</ol>
	</li>
	<li>Dehydration and resin embedding
	<ol>
		<li>Prepare epoxy resin. Weigh the ingredients (33.3 g of component A/M, 33.3 g of component B, and 1 g of component D) and mix the resin well (e.g., shake it on a rotary shaker in a 15 mL tube) for at least 30 minutes before adding 16 drops of accelerator DMP 30. Stir again for another 10 minutes. 
		NOTE: Weigh the ingredients under the fume hood. This amount of components gives approximately 60 mL of resin what is enough for 15 vials with samples. You can prepare less or store the rest of the resin in a syringe at 4 &#176;C and utilize it the next day. Remember to seal the tip of the syringe.</li>
		<li>Prepare vials with graded dilutions of ethanol (30%, 50%, 70%, 90%, 100%, 100% ethanol in water and 100% dried on a molecular sieve).</li>
		<li>Mix the resin with 100% ethanol in 1:1 proportion to obtain 50% resin. Mix it well.</li>
		<li>Dehydrate the samples for 5 minutes in each dilution of ethanol starting from 30% and ending with 100% anhydrous ethanol dried on molecular sieve. Remember, the samples must never dry completely.</li>
		<li>Infiltrate the samples first in 50% resin for 30 minutes, next in 100% resin for one hour, and then again in 100% resin overnight. Perform all infiltration steps with constant slow shaking.</li>
		<li>The next day, place the samples in fresh 100% resin for one hour and then embed them between fluoropolymer embedding sheets. Degrease pieces of embedding sheet with ethanol, and flat embed the samples between two layers of it, using two glass slides as support. Try to avoid air bubbles in resin on or close to the sample.</li>
		<li>Cure the samples in the oven at 70 &#176;C for at least 48 hours.</li>
	</ol>
	</li>
	<li>Trimming and mounting
	<ol>
		<li>Separate embedding sheets and cut a piece of the embedded sample (approximately 1 mm x 1 mm) with a razor blade. Transfer it to a parafilm. This will minimize the danger of losing the sample due to electrostatics.</li>
		<li>Take an aluminum pin that has been degreased with ethanol. Mix a conductive epoxy well and use a little amount of it to mount the sample to the pin. Cure conductive epoxy at 70 &#176;C for 10 minutes.</li>
		<li>Trim each side of the sample block with the diamond knife and then polish the face of the block until the tissue is exposed. 
		NOTE: At this step, confirm the presence of a region of interest by collecting some sections and performing toluidine blue staining16 or checking under the electron microscope.</li>
		<li>Reduce the size of the sample as much as possible. Then ground the sample to the pin with conductive paint and cure it (for 24 hours at room temperature or in the oven at 65 &#176;C for 40 minutes).</li>
		<li>To minimize charging artifacts, sputter coat the samples with a thin layer of gold or gold/palladium.</li>
	</ol>
	</li>
</ol>
9. SBEM imaging
<ol>
	<li>Place the pin with a sample into the chamber of the serial block-face scanning electron microscope. Align the sample to the knife. Close the chamber and set the parameters.</li>
	<li>Collect a stack of images at desired magnification, pixel size, slice thickness, accelerated voltage (EHT), aperture, pressure, etc. 
	&#8203;NOTE: The parameters depend on the sample and the goal of the experiment. Exemplary initial imaging settings are included in the table of materials.</li>
</ol>
10. 3D reconstructions
NOTE: For the steps mentioned below we use open-access software e.g. FijiJ17 (ImageJ version 1.49b), Microscopy Image Browser (MIB)18 and Reconstruct19 but various other software can be employed.
<ol>
	<li>Convert digital micrograph (dm) files into TIFF format. First import the image sequence (in FijiJ: File &#62; Import &#62; Image sequence &#62; Choose 8 Bit Format) and then save as TIFF (in FijiJ: File &#62; Save As &#62; Image Sequence &#62; Choose Tiff).</li>
	<li>Adjust brightness and contrast of the stack of images (in FijiJ: Image &#62; Adjust &#62; Brightness/Contrast) and, if necessary, denoise it (use DenoisEM plugin for FijJ20 : Plugins &#62; DenoiseEM &#62; Denoise).</li>
	<li>Align the stack&#160;(in FijiJ: Plugins &#62; StagReg or MIB: Dataset &#62; Alignment Tools).</li>
	<li>Segment dendritic spines and synapses&#160;(In MIB or Reconstruct software, comprehensive tutorials are available (See Table of Materials for details).</li>
</ol>

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R. G., 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=3D+correlative+light+and+electron+microscopy+of+cultured+cells+using+serial+blockface+scanning+electron+microscopy.">3D correlative light and electron microscopy of cultured cells using serial blockface scanning electron microscopy.</a> Journal of Cell Science. 130 (1), 278-291 (2017).</li><li>P&#322;achno, B. J., &#346;wi&#261;tek, P., Jobson, R. W., Ma&#322;ota, K., Brutkowski, W. <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+SEM+visualization+of+unusual+plant+nuclear+tubular+extensions+in+a+carnivorous+plant+(Utricularia,+Lentibulariaceae).">Serial block face SEM visualization of unusual plant nuclear tubular extensions in a carnivorous plant (Utricularia, Lentibulariaceae).</a> Annals of Botany. 120 (5), 673-680 (2017).</li><li>Genoud, C., Titze, B., Graff-Meyer, A., Friedrich, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+Homogeneous+En+Bloc+Staining+of+Large+Tissue+Samples+for+Volume+Electron+Microscopy.">Fast Homogeneous En Bloc Staining of Large Tissue Samples for Volume Electron Microscopy.</a> Frontiers in Neuroanatomy. 12, (2018).</li><li>Puhka, M., Joensuu, M., Vihinen, H., Belevich, I., Jokitalo, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progressive+sheet-to-tubule+transformation+is+a+general+mechanism+for+endoplasmic+reticulum+partitioning+in+dividing+mammalian+cells.">Progressive sheet-to-tubule transformation is a general mechanism for endoplasmic reticulum partitioning in dividing mammalian cells.</a> Molecular Biology of the Cell. 23 (13), 2424-2432 (2012).</li><li>Gluenz, E., Wheeler, R. J., Hughes, L., Vaughan, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+and+three-dimensional+electron+microscopy+methods+for+the+study+of+Trypanosoma+brucei+and+Leishmania+mexicana+flagella.">Scanning and three-dimensional electron microscopy methods for the study of Trypanosoma brucei and Leishmania mexicana flagella.</a> Methods in Cell Biology. 127, 509-542 (2015).</li><li>Starborg, T., 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=Using+transmission+electron+microscopy+and+3View+to+determine+collagen+fibril+size+and+three-dimensional+organization.">Using transmission electron microscopy and 3View to determine collagen fibril size and three-dimensional organization.</a> Nature Protocols. 8 (7), 1433-1448 (2013).</li><li>Hughes, L., Borrett, S., Towers, K., Starborg, T., Vaughan, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterns+of+organelle+ontogeny+through+a+cell+cycle+revealed+by+whole-cell+reconstructions+using+3D+electron+microscopy.">Patterns of organelle ontogeny through a cell cycle revealed by whole-cell reconstructions using 3D electron microscopy.</a> Journal of Cell Science. 130 (3), 637-647 (2017).</li><li>Bojko, A., 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=Improved+Autophagic+Flux+in+Escapers+from+Doxorubicin-Induced+Senescence/Polyploidy+of+Breast+Cancer+Cells.">Improved Autophagic Flux in Escapers from Doxorubicin-Induced Senescence/Polyploidy of Breast Cancer Cells.</a> International Journal of Molecular Sciences. 21 (17), 6084 (2020).</li><li>Knott, G. W., Holtmaat, A., Trachtenberg, J. T., Svoboda, K., Welker, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+preparing+GFP-labeled+neurons+previously+imaged+in+vivo+and+in+slice+preparations+for+light+and+electron+microscopic+analysis.">A protocol for preparing GFP-labeled neurons previously imaged in vivo and in slice preparations for light and electron microscopic analysis.</a> Nature Protocols. 4 (8), 1145-1156 (2009).</li><li>Glauert, A. M., Lewis, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biological specimen preparation for transmission electron microscopy. , (2014).</li><li>Genoud, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Altered+Synapse+Formation+in+the+Adult+Somatosensory+Cortex+of+Brain-Derived+Neurotrophic+Factor+Heterozygote+Mice.">Altered Synapse Formation in the Adult Somatosensory Cortex of Brain-Derived Neurotrophic Factor Heterozygote Mice.</a> Journal of Neuroscience. 24 (10), 2394-2400 (2004).</li></ol>

The authors have nothing to disclose.

There are many variations of the primary NCMIR method described by Deerinck in 201010. The basic principles remain the same but, depending on the type of material studied, slight changes are implemented. It was described previously that different resins can be used to embed specimens for SBEM and for example in the case of plants, Spurr&#39;s is the resin of choice due to its low viscosity that allows better infiltration through the cell walls22,<sup class="xref"...

Most of the excitatory synapses in the central nervous system are located on dendritic spines - small protrusions of a neuronal membrane. These protrusions form confined biochemical compartments that control intracellular signal transduction. Structural plasticity of dendritic spines and synapses is closely related to the functional changes in synaptic efficacy that underlie such important processes as learning and memory1,2. It is important to note that electron microscopy (EM) is the only technique that allows to determine if a dendritic spine has a presynaptic input. EM resolution is also needed to study ultrastructural details such as shape of a postsynaptic density (PSD), representing a postsynaptic part of a synapse, or dimensions of a dendritic spine, as well as the size and shape of an axonal bouton. Additionally, with EM it is possible to visualize synapses and their surroundings.
Thanks to advances in imaging and computing technologies it is possible to reconstruct entire neural circuits. Volume electron microscopy techniques, such as serial section transmission electron microscopy (ssTEM), serial block-face scanning electron microscopy (SBEM), and focused ion beam scanning electron microscopy (FIB-SEM) are commonly used for neuronal circuit reconstructions3.
In our studies, the SBEM method is successfully employed to investigate the structural plasticity of dendritic spines and PSDs in samples of the mouse hippocampus and organotypic brain slices 4,5. The SBEM is based on the installation of a miniature ultramicrotome inside the scanning electron microscope chamber6,7,8,9. The top of the sample block is imaged, and then the sample is cut at a specified depth by the ultramicrotome, revealing a new block-face, which is again imaged and then the process is repeated8. As a result, only the image of a block-face is left while the slice which has been cut is lost as debris. That is why SBEM is called a destructive technique, meaning it is not possible to image the same place again. However, the advantage of the destructive on-block methods is that they do not suffer from warping problems and section loss that can significantly affect the data quality and the data analysis3. Moreover, SBEM gives the possibility to image a relatively large field of view ( &#62; 0.5 mm &#215; 0.5 mm) at high resolution3.
To employ SBEM, samples have to be prepared according to a dedicated, highly contrasting protocol due to the backscattered electron detector used for acquiring images. We show here how to perform sample preparation according to the protocol based on a procedure developed by Deerinck10 (National Center for Microscopy and Imaging Research (NCMIR)&#160;method), using reduced osmium-thiocarbohydrazide-osmium (rOTO) stains developed in the 1980s8,11. In addition, we introduce a two-step fixation approach, with mild fixation perfusion that allows to use the same brain both for immunofluorescence studies with light microscopy and SBEM.
In the protocol a mouse brain is primarily fixed with a mild fixative, and then cut into halves, and one hemisphere is postfixed and prepared for immunofluorescence (IF), while the other for EM studies (Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62712/62712fig01.jpg" /> 
Figure 1. Schematic representation of the workflow for the dendritic spines preparation for the analysis with SBEM. Mice were sacrificed and perfused with a mild primary fixative. The brain was cut into halves, and one hemisphere was postfixed with immunofluorescence (IF)-dedicated fixative, cryoprotected, sliced using a cryostat and processed for IF studies, while the other hemisphere was postfixed with EM fixative, sliced with the vibratome and prepared for EM studies. Brain slices for SBEM studies were contrasted, flat embedded in resin, then a CA1 region of the hippocampus was mounted to the pin, and imaged with SBEM (Figure 1). The part of the protocol highlighted in a yellow box was featured in the video. <a href="https://www.jove.com/files/ftp_upload/62712/62712fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Anesthetic:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine/xylazine mixture (Ketamina/Sedazin)</td>
			<td>Biowet Pulawy, Pulawy, Poland</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium pentobarbital (Morbital)</td>
			<td>Biowet Pulawy, Pulawy, Poland</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fixatives:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde (GA)</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>G5882</td>
			<td>Grade I, 25% in H2O, specially purified for use as an electron microscopy fixative</td>
		</tr>
		<tr>
			<td>Hydrochloric acid (HCl)</td>
			<td>POCH, Gliwice, Poland</td>
			<td>575283115</td>
			<td>pure p.a.</td>
		</tr>
		<tr>
			<td>Paraformaldehyde (PFA)</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>441244</td>
			<td>prilled, 95%</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS), pH 7.4</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>P4417-50TAB</td>
			<td>tablets</td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>S5881</td>
			<td>reagent grade, <img alt="Equation" src="/files/ftp_upload/62712/62712symbol.png" />98%, pellets (anhydrous)</td>
		</tr>
		<tr>
			<td>Sodium phosphate dibasic (Na2HPO4)</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>S3264</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium phosphate monobasic (NaH2PO4)</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>S3139</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusion:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Large blunt/blunt curved scissors (~14.5 cm)</td>
			<td>Fine Science Tools, Foster City, CA, USA</td>
			<td>14519-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-spatula (double 2&#34; flat ends, one rounded, one tapered to 1/8&#34;)</td>
			<td>Fine Science Tools, Foster City, CA, USA</td>
			<td>10091-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle tip, 15 GA, blunt (perfusion needle)</td>
			<td>KD Medical GmbH Hospital Products, Berlin, Germany</td>
			<td>KD-FINE 900413</td>
			<td>1.80 x 40 mm</td>
		</tr>
		<tr>
			<td>Pair of fine (Graefe) tweezers</td>
			<td>Fine Science Tools, Foster City, CA, USA</td>
			<td>11050-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusion pump</td>
			<td>Lead Fluid</td>
			<td>BQ80S</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic vials</td>
			<td>Profilab, Warsaw, Poland</td>
			<td>534.02</td>
			<td>plastic vials with blue cap for tissue storage, 20 ml, 31 x 48 mm</td>
		</tr>
		<tr>
			<td>Straight iris scissors (~9 cm)</td>
			<td>Fine Science Tools, Foster City, CA, USA</td>
			<td>14058-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Brain slices preparation for EM:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>12-well plate</td>
			<td>NEST, Rahway, NJ, USA</td>
			<td>712001</td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrylic glue</td>
			<td>Fenedur, Montevideo, Uruguay</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass vials</td>
			<td>Electron Microscopy Sciences, Hatfield, PA, USA</td>
			<td>72632</td>
			<td>20 ml Scintillation Vial, a pack of 100</td>
		</tr>
		<tr>
			<td>Pasteur pipette</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>612-4545</td>
			<td>LDPE, disposable, 7.5 ml</td>
		</tr>
		<tr>
			<td>Razor blade</td>
			<td>Wilkinson Sword, London, UK</td>
			<td></td>
			<td>Classic double edge safety razor blades</td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td>Swann-Morton, Sheffield, UK</td>
			<td>No. 20</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica Microsystems, Vienna, Austria</td>
			<td>Leica VT1000 S</td>
			<td></td>
		</tr>
		<tr>
			<td>Brain slices preparation for IF:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>96-well plate</td>
			<td>NEST, Rahway, NJ, USA</td>
			<td>701101</td>
			<td></td>
		</tr>
		<tr>
			<td>Criostat</td>
			<td>Leica Microsystems, Vienna, Austria</td>
			<td>Leica CM 1950</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylene glycol</td>
			<td>Bioshop, Burlington, Canada</td>
			<td>ETH001</td>
			<td></td>
		</tr>
		<tr>
			<td>Low-profile disposable blade 819</td>
			<td>Leica Biosystems Inc., USA</td>
			<td>14035838925</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td>Swann-Morton, Sheffield, UK</td>
			<td>No. 20</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide (NaN3)</td>
			<td>POCH, Gliwice, Poland</td>
			<td>792770426</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>POCH, Gliwice, Poland</td>
			<td>772090110</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue freezing medium for cryosectioning, OCT-Compound</td>
			<td>Leica Biosystems, Switzerland</td>
			<td>14020108926</td>
			<td></td>
		</tr>
		<tr>
			<td>Immunostaining:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>24-well plate</td>
			<td>NEST, Rahway, NJ, USA</td>
			<td>702001</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Post Synaptic Density Protein 95 Antibody</td>
			<td>Merck-Millipore, Burlington, MA, USA</td>
			<td>MAB1598</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Zeiss, G&#246;ttingen, Germany</td>
			<td></td>
			<td>Zeiss Spinning Disc microscope (63 &#215; oil objective, NA 1.4, pixel size 0.13 &#181;m &#215; 0.13 &#181;m)</td>
		</tr>
		<tr>
			<td>Cover slide</td>
			<td>Menzel Glaser, Braunschweig, Germany</td>
			<td>B-1231</td>
			<td>24 x 60 mm</td>
		</tr>
		<tr>
			<td>Donkey anti-Mouse IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 555</td>
			<td>Invitrogen, Carlsbad, CA, USA</td>
			<td>A31570</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoromount-G Mounting Medium, with DAPI</td>
			<td>Invitrogen, Carlsbad, CA, USA</td>
			<td>00-4959-52</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slide</td>
			<td>Thermo Scientific, Waltham, MA, USA</td>
			<td>AGAA00008</td>
			<td>SuperFrost</td>
		</tr>
		<tr>
			<td>Normal donkey serum (NDS)</td>
			<td>Jackson ImmunoResearch Laboratories, West Grove, PA, USA</td>
			<td>017-000-121</td>
			<td></td>
		</tr>
		<tr>
			<td>Shaker</td>
			<td>JWElectronic, Warsaw, Poland</td>
			<td>KL-942</td>
			<td></td>
		</tr>
		<tr>
			<td>TritonT X-100 Reagent Grade</td>
			<td>Bioshop, Burlington, Canada</td>
			<td>TRX506</td>
			<td></td>
		</tr>
		<tr>
			<td>Electron microsocpy sample preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hexacyanoferrate(II) trihydrate</td>
			<td>POCH, Gliwice, Poland</td>
			<td>746980113</td>
			<td></td>
		</tr>
		<tr>
			<td>Aclar 33C Film</td>
			<td>Electron Microscopy Sciences, Hatfield, PA, USA</td>
			<td>50425</td>
			<td>Fluoropolymer Film embedding sheet</td>
		</tr>
		<tr>
			<td>DMP-30, 2,4,6-Tris(dimethylaminomethyl)phenol</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>T58203</td>
			<td>Epoxy embedding medium accelerator</td>
		</tr>
		<tr>
			<td>Durcupan ACM single component A, M</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>44611</td>
			<td>Durcupan ACM single component A, M epoxy resin</td>
		</tr>
		<tr>
			<td>Durcupan ACM single component B</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>44612</td>
			<td>Durcupan ACM single component B, hardener 964</td>
		</tr>
		<tr>
			<td>Durcupan ACM single component D</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>44614</td>
			<td>Durcupan ACM single component D , plasticizer</td>
		</tr>
		<tr>
			<td>Ethyl alcohol absolute</td>
			<td>POCH, Gliwice, Poland</td>
			<td>64-17-5</td>
			<td>Ethyl alcohol absolute 99.8 % pure P.A.-BASIC</td>
		</tr>
		<tr>
			<td>Genlab laboratory oven</td>
			<td>Wolflabs, York, UK</td>
			<td>Mino/18/SS</td>
			<td>Oven Genlab MINO/18/SS 18l volume, no fan circulation, no digital display, standard temperature gradient, standard recovery rate, no timer, 250&#176;C maximum temperature, 240V electrical supply</td>
		</tr>
		<tr>
			<td>L-Aspartic acid</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>A-9256</td>
			<td>reagent grade, <img alt="Equation" src="/files/ftp_upload/62712/62712symbol.png" />98% (HPLC)</td>
		</tr>
		<tr>
			<td>Lead (II) nitrate</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>467790</td>
			<td><img alt="Equation" src="/files/ftp_upload/62712/62712symbol.png" />99.95% trace metals basis</td>
		</tr>
		<tr>
			<td>Osmium tetroxide</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>75632</td>
			<td>for electron microscopy, 4% in H2O</td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Elmetron, Zabrze, Poland</td>
			<td>CP-5-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Rotator</td>
			<td>BioSan, J&#243;zef&#243;w, Poland</td>
			<td>Multi Bio RS-24</td>
			<td>rotator Multi Bio RS-24</td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>S5881</td>
			<td>reagent grade, <img alt="Equation" src="/files/ftp_upload/62712/62712symbol.png" />98%, pellets (anhydrous)</td>
		</tr>
		<tr>
			<td>Sunflower mini shaker</td>
			<td>Grant bio, Shepreth,UK</td>
			<td>PD-3D</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter</td>
			<td>Millipore, Burlington, MA, USA</td>
			<td>SLGP033NB</td>
			<td>0,22 &#181;m pore size</td>
		</tr>
		<tr>
			<td>Thiocarbohydrazide</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>88535</td>
			<td>purum p.a., for electron microscopy, <img alt="Equation" src="/files/ftp_upload/62712/62712symbol.png" />99.0% (N)</td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Serva, Heidelberg, Germany</td>
			<td>77870</td>
			<td>Uranyl acetate&#183;2H2O, research grade</td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>WSL, Swietochlowice, Poland</td>
			<td>LWT</td>
			<td></td>
		</tr>
		<tr>
			<td>Specimen mounting for SBEM</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>96-well culture plate</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>734-2782</td>
			<td>96-well plates, round bottom, non treated</td>
		</tr>
		<tr>
			<td>AM Gatan 3View stub handling tweezers</td>
			<td>Micro to Nano, Haarlem, Netherlands 
			Netherlands</td>
			<td>50-001521</td>
			<td></td>
		</tr>
		<tr>
			<td>Binocular</td>
			<td>OPTA-TECH, Warsaw, Poland</td>
			<td>X2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Conductive glue</td>
			<td>Chemtronics, Georgia, USA</td>
			<td>CW2400</td>
			<td>conductive eopxy</td>
		</tr>
		<tr>
			<td>Gatan 3View sample pin stubs</td>
			<td>Micro to Nano, Haarlem, Netherlands 
			Netherlands</td>
			<td>10-006003</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Sigma-Aldrich,St. Louis, MI, USA</td>
			<td>P7793</td>
			<td>roll size 20&#160;in. &#215; 50&#160;ft</td>
		</tr>
		<tr>
			<td>Pelco conductive silver paint</td>
			<td>Ted Pella, Redding, CA, USA</td>
			<td>16062-15</td>
			<td>PELCO&#174;&#160;Conductive Silver Paint, 15g</td>
		</tr>
		<tr>
			<td>Razor blades double edge</td>
			<td>Electron Microscopy Sciences, Hatfield, PA, USA</td>
			<td>72000</td>
			<td>Stainless Steel &#34;PTFE&#34; coated. PERSONNA brand .004&#34; thick, wrapped individually, 250 blades in a box.</td>
		</tr>
		<tr>
			<td>Scanning Electron Microscope</td>
			<td>Zeiss, Oberkochen, Germany</td>
			<td></td>
			<td>Sigma VP with Gatan 3View2 chamber, acceleration voltage 2.5 kV, variable pressure 5 Pa, aperture 20 &#181;m, dwell time 6 &#181;s, slice thickness 60 nm, magnification 15 000 x, image resolution 2048 x 2048 pixels, pixel size 7.3 nm</td>
		</tr>
		<tr>
			<td>trim 90&#176; diamond knife</td>
			<td>Diatome Ltd., Nidau, Switzerland</td>
			<td>DTB90</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultramicrotome</td>
			<td>Leica Microsystems, Vienna, Austria</td>
			<td>Leica ultracutR</td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td>webpage</td>
			<td></td>
			<td>tutorials</td>
		</tr>
		<tr>
			<td>FijiJ</td>
			<td>https://fiji.sc/</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscopy Image Browser</td>
			<td>http://mib.helsinki.fi/</td>
			<td></td>
			<td>http://mib.helsinki.fi/tutorials.html</td>
		</tr>
		<tr>
			<td>Reconstruct</td>
			<td>https://synapseweb.clm.utexas.edu/software-0</td>
			<td></td>
			<td>https://synapseweb.clm.utexas.edu/software-0)</td>
		</tr>
		<tr>
			<td>Animals</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mice</td>
			<td>Adult 3-month old and&#160;20&#177;1&#160;month old&#160;female&#160;Thy1-GFP(M) mice (Thy1-GFP +/-) (Feng et al.,2000) which express GFP in a sparsely distributed population of glutamatergic neurons. Animals were bred as heterozygotes with the C57BL/6J background in the Animal House of the Nencki Institute of Experimental Biology.</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

serial block-face scanning electron microscopy (sbem) for the study of dendritic spines

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In addition, some features of dendritic spines, such as volume of the spine and post-synaptic density (PSD) (representing post-synaptic part of the synapse), presence of presynaptic terminal, and smooth endoplasmic reticulum or atypical form of PSD (e.g., multi-innervated spines), can be observed only with 3D EM. By employing serial block-face scanning electron microscopy (SBEM) it is possible to obtain 3D EM data easier and in a more reproducible manner than when performing traditional serial sectioning. Here we show how to prepare mouse hippocampal samples for SBEM analysis and how this protocol can be combined with immunofluorescence study of dendritic spines. Mild fixation perfusion allows us to perform immunofluorescence studies with light microscopy on one half of the brain, while the other half was prepared for SBEM. This approach reduces the number of animals to be used for the study.

Using the method described above high contrast, good resolution images of the mouse brain tissue can be obtained. A large field of view provided by the SBEM technique facilitates precise selection of the region of interest. The large image of the CA1 region of the hippocampus was taken to measure the length of stratum radiatum (SR) (Figure 2A) and to set the imaging precisely in the center (Figure 2B). Next, the stack of images was acquired and the obje...

Watch this Scientific Journal Video about Serial Block-Face Scanning Electron Microscopy SBEM for the Study of Dendritic Spines at JoVE.com

Serial Block-Face Scanning Electron Microscopy SBEM for the Study of Dendritic Spines

Serial Block-Face Scanning Electron Microscopy (SBEM) is applied to image and analyze dendritic spines in the murine hippocampus.

Serial Block-Face Scanning Electron Microscopy (SBEM) for the Study of Dendritic Spines

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In ...

serial-block-face-scanning-electron-microscopy-sbem-for-study

Research

JoVE Journal

Neuroscience

3.6K Views.  Nencki Institute of Experimental Biology of Polish Academy of Sciences. Serial Block-Face Scanning Electron Microscopy (SBEM) is applied to image and analyze dendritic spines in the murine hippocampus. 

Video: Serial Block-Face Scanning Electron Microscopy SBEM for the Study of Dendritic Spines

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Analysis of Brain Mitochondria Using Serial Block-Face Scanning Electron Microscopy

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, tri-carboxylic acid (TCA) cycle and oxidative phosphorylation (OXPHOS) pathways to produce cellular energy in the form of adenosine triphosphate (ATP). Impairment of mitochondrial ATP production causes mitochondrial disorders, which present clinically with prominent neurological and myopathic symptoms. Mitochondrial defects are also present in neurodevelopmental disorders (e.g. autism spectrum disorder) and neurodegenerative disorders (e.g. amyotrophic lateral sclerosis, Alzheimer&#39;s and Parkinson&#39;s diseases). Thus, there is an increased interest in the field for performing 3D analysis of mitochondrial morphology, structure and distribution under both healthy and disease states. The brain mitochondrial morphology is extremely diverse, with some mitochondria especially those in the synaptic region being in the range of &#60;200 nm diameter, which is below the resolution limit of traditional light microscopy. Expressing a mitochondrially-targeted green fluorescent protein (GFP) in the brain significantly enhances the organellar detection by confocal microscopy. However, it does not overcome the constraints on the sensitivity of detection of relatively small sized mitochondria without oversaturating the images of large sized mitochondria. While serial transmission electron microscopy has been successfully used to characterize mitochondria at the neuronal synapse, this technique is extremely time-consuming especially when comparing multiple samples. The serial block-face scanning electron microscopy (SBFSEM) technique involves an automated process of sectioning, imaging blocks of tissue and data acquisition. Here, we provide a protocol to perform SBFSEM of a defined region from rodent brain to rapidly reconstruct and visualize mitochondrial morphology. This technique could also be used to provide accurate information on mitochondrial number, volume, size and distribution in a defined brain region. Since the obtained image resolution is high (typically under 10 nm) any gross mitochondrial morphological defects may also be detected.

Mitochondrial visualization and analysis from mammalian brain tissue is a challenging task. Here, we describe how three dimensional (3D) reconstruction analysis from the serial block-face scanning electron microscopy (SBFSEM) can be used to gain insights on the morphological and volumetric analysis of this critical energy generating organelle.

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, ...

3D Modeling of Dendritic Spines with Synaptic Plasticity

Computational modeling of diffusion and reaction of chemical species in a three-dimensional (3D) geometry is a fundamental method to understand the mechanisms of synaptic plasticity in dendritic spines. In this protocol, the detailed 3D structure of the dendrites and dendritic spines is modeled with meshes on the software Blender with CellBlender. The synaptic and extrasynaptic regions are defined on the mesh. Next, the synaptic receptor and synaptic anchor molecules are defined with their diffusion constants. Finally, the chemical reactions between synaptic receptors and synaptic anchors are included and the computational model is solved numerically with the software MCell. This method describes the spatiotemporal path of every single molecule in a 3D geometrical structure. Thus, it is very useful to study the trafficking of synaptic receptors in and out of the dendritic spines during the occurrence of synaptic plasticity. A limitation of this method is that the high number of molecules slows the speed of the simulations. Modeling of dendritic spines with this method allows the study of homosynaptic potentiation and depression within single spines and heterosynaptic plasticity between neighbor dendritic spines.

The protocol develops a three-dimensional (3D) model of a dendritic segment with dendritic spines for modeling synaptic plasticity. The constructed mesh can be used for computational modeling of AMPA receptor trafficking in the long-term synaptic plasticity using the software program Blender with CellBlender and MCell.

Computational modeling of diffusion and reaction of chemical species in a three-dimensional (3D) geometry is a fundamental method to understand the ...

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic spines have been described for DD GABAergic neurons as the input sites from presynaptic cholinergic neurons in the motor circuit of Caenorhabditis elegans. This synaptic circuit can now serve as a powerful new in vivo model of spine morphogenesis and function that exploits the facile genetics and ready accessibility of C. elegans to live-cell imaging. 
This protocol describes experimental strategies for assessing DD spine structure and function. In this approach, a super-resolution imaging strategy is used to visualize the intricate shapes of actin-rich dendritic spines. To evaluate the DD spine function, the light-activated opsin, Chrimson, stimulates the presynaptic cholinergic neurons, and the calcium indicator, GCaMP, reports the evoked calcium transients in postsynaptic DD spines. Together, these methods comprise powerful approaches for identifying genetic determinants of dendritic spines in C. elegans that could also direct spine morphogenesis and function in the brain.

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are important cellular features of the nervous system. Here live imaging methods are described for assessing the structure and function of dendritic spines in C. elegans.&#160;These approaches support the development of mutant screens for genes that define dendritic spine shape or function.

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic ...