We thank Shao-Ming Lu and Harris A. Gelbard for the use of a peristaltic pump and vibratome, as well as Karen L. Bentley and Gayle Schneider at the Electron Microscope Research Core Facility of the University of Rochester Medical Center for technical assistance. This work was funded by grants from the NIH (EY019277), Whitehall Foundation, Burroughs Wellcome Fund, and the Alfred P. Sloan Foundation to A.K.M.. M.-&Egrave;.T. is funded by a Fonds de la recherche en sant&eacute; du Qu&eacute;bec (FRSQ) postdoctoral training award.

1. Animal perfusion
<ol>
 <li>On the day before perfusion, prepare: 
 - 2 L of phosphate buffer (PB; 100 mM, pH 7.4) and 1 L of sodium phosphate-buffered saline (PBS; 0.9% NaCl in 50 mM PB, pH 7.4) in double distilled water. Store the PBS and 1L of PB at 4&deg;C. These will be used for perfusion and pre-embedding immunocytochemistry. 
 - 1L of 4.0% paraformaldehyde (PFA, pH 7.4) fixative solution, which will enable the perfusion of 6 mice. For that purpose, heat up 1 L of PB under a fume hood. Weigh 40 g of granular paraformaldehyde (PFA) and pour into the PB solution when it reaches 60&deg;C. When the solution is clear, with the PFA completely dissolved, cool to room temperature (RT) and store at 4&deg;C.</li>
 <li>On the day of perfusion, prepare 500 mL of 3.5% acrolein solution in PB under a fume hood. Then, filter the 3.5% acrolein and 4.0% PFA solutions using filter paper.</li>
 <li>For mouse anesthesia, inject sodium pentobarbital (80 mg/kg) into the peritoneum with a 27 gauge &frac12;" needle.</li>
 <li>During the perfusion, 50 mL of PBS, 75 mL of acrolein, and 150 mL of PFA will sequentially pass into the mouse circulation. To set up the peristaltic pump, fill the tubing with PBS and fix a 23 gauge &frac34;" butterfly needle at one end. Immerse the other end of the tubing into the perfusion solution (PBS, acrolein or PFA). Set the speed of the peristaltic pump to 20-25 mL/min for juvenile and adult mice. Throughout the perfusion, carefully avoid any bubbles of air forming in the tubing.</li>
 <li>Wait until the anaesthetized mouse no longer responds to painful stimuli, such as a tail pinch, before proceeding. Lay the animal in a dissection tray and fix the paws using tape. With tweezers and scissors, open up the skin and chest cavity to expose the heart. To minimize brain ischemia, the perfusion needs to be started rapidly. Cut open the right atrium with small scissors and start the peristaltic pump. While holding the heart with tweezers, insert the butterfly needle into the apex of the left ventricle.</li>
 <li>When changing solution, stop the peristaltic pump, transfer the tubing from one solution to another, and restart the pump immediately.</li>
 <li>When 150 mL of PFA has passed, stop the peristaltic pump. Using small scissors, cut off the head, open up the skin, and break the skull between the eyes. Using small tweezers, carefully chip off small pieces of skull until the brain can be easily removed. Post-fix the brain for 2 hours at 4&deg;C in PFA. Wash 3 times 10 minutes in PBS.</li>
 <li>Immediately cut the brain in transverse (50 &mu;m thick sections) in ice-cooled PBS using a vibratome. With a fine brush, transfer sections selected for immunocytochemistry into a glass vial containing PBS. Store the remaining sections at -20&deg;C in cryoprotectant (30% ethylene glycol and 30% glycerol in PBS) for up to several years.</li>
</ol>
2. Pre-embedding immunocytochemistry
<ol>
 <li>For immunocytochemistry, sections are processed freely floating in glass vials. Throughout the procedure, one needs to carefully avoid letting sections dry out.</li>
 <li>First, remove the PBS using a transfer pipette and immediately replace with a fresh solution of 0.1% sodium borohydride in PBS for 30 minutes at RT.</li>
 <li>Rinse sections with PBS 3 times 10 minutes, removing all bubbles, and incubate for 2 hours at RT in a blocking solution of PBS containing 0.5% gelatin and 5% normal serum of the animal in which the secondary antibody was generated (normal goat serum in the present example of Iba1-immunostaining).</li>
 <li>Incubate for 48 hours with primary antibody in blocking solution. For Iba1-immunostaining, use rabbit anti-Iba1 antibody (1:1000).</li>
 <li>Rinse sections with PBS 3 times 10 minutes and incubate in secondary antibody conjugated to biotin (1:1000) in blocking solution for 2 hours at RT. 
 For Iba1-immunostaining, use goat anti-rabbit IgGs.</li>
 <li>Rinse sections with PBS 3 times 10 minutes and incubate in streptavidin-horseradish peroxidase (1:1000) in blocking solution for 1 hour at RT.</li>
 <li>Rinse sections with PBS 10 minutes and with Tris/HCl-buffered saline (TBS; 50 mM, pH 7.4) 2 times 10 minutes. To reveal the labeling, incubate sections with 0.05% DAB and 0.01% hydrogen peroxide in TBS (solution freshly prepared). When the staining is dark-brown (see example in Figure 1), stop the reaction by rinsing in TBS. In any case, end the reaction after a maximum of 5 minutes. Rinse sections with TBS 10 minutes and with PB 2 times 10 minutes.</li>
</ol>
3. Processing for electron microscopy
<ol>
 <li>Transfer sections into PB in a multiwell culture plate using a fine brush. Prepare 1% osmium tetroxide solution in PB under a fuming hood. Remove PB from the wells of the culture plate and spread the sections flat with a fine brush. Immediately immerse sections in 1% osmium for 30 minutes at RT.</li>
 <li>Transfer the sections into glass vials, rinse with PB 3 times 10 minutes, and dehydrate through 2 minutes immersions into ascending concentrations of ethanol: 2 times 35%, 1 time each of 50%, 70%, 80%, 90%, and 95%, 3 times 100%, followed by 3 times propylene oxide.</li>
 <li>To prepare the Durcupan resin, combine 20 g of component A, 20 g of component B, 0.6 g of component C, and 0.4 g of component D into a disposable beaker, mix well until the color becomes uniform, and pour into an aluminum weigh dish. If needed, resin blocks can be prepared by pouring resin into embedding molds and curing in a 55&deg;C oven for 48-72 hours.</li>
 <li>Using a fine brush rinsed with propylene oxide, remove the sections from the propylene oxide and immerse into the Durcupan resin for impregnation overnight at RT.</li>
 <li>The following day, put the aluminum weigh dish into a 55&deg;C oven for 10 minutes to soften the resin. Using a brush rinsed with propylene oxide, coat an ACLAR embedding film with a thin layer of resin, lay down the sections, cover with another embedding film, and evenly distribute light weights (about 2 g; for example the plastic caps of glass vials) on top to help resin spreading. For resin polymerization, incubate in the oven for 48-72 hours (at 55&deg;C).</li>
 <li>After polymerization, remove the light weights and embedding film on the top of the sections. Under a binocular microscope, select square areas of interest (about 2x2 mm) and carefully excise them from the film using a razor blade.</li>
 <li>Glue the areas of interest at the tip of resin blocks using Superglue and cure in the 55&deg;C oven for 1 hour.</li>
 <li>In preparation for sectioning, trim the resin block to the shape of an isosceles trapezoid with a razor blade.</li>
 <li>Using an ultramicrotome and a glass knife, remove the glue and resin at the surface of the tissue. Fill the boat of the glass knife with double distilled water and cut a few semi-thin sections (0.5-1 &mu;m thick).</li>
 <li>Transfer the sections to a SuperFrost slide with a perfect loop. Dry the sections by placing the slide on a heating plate at 80&deg;C for 1 minute. Cover with a few drops of 0.1% toluidine blue stain (2 g sodium borate and 0.2 g toluidine blue in 200 mL double distilled water) for 1 minute. Rinse the excess stain with double distilled water. Examination of the sections with a light microscope will enable to distinguish the stained tissue from the resin (Figure 2).</li>
 <li>Resume cutting until the border between tissue and resin reaches the middle of the sections. Note that the following procedure requires training. Replace the glass knife with a diamond knife and fill the boat with double distilled water. Cut silver to silver-gold ultrathin sections (60-80 nm thick) and carefully collect them on copper mesh grids using fine inverted tweezers. Dry the grids on a filter paper.</li>
 <li>To enhance contrast, ultrathin sections can be stained with lead citrate (0.03 g lead citrate and 0.1 mL 10N sodium hydroxide in 10 mL double distilled water1). Using fine inverted tweezers, transfer a grid to a drop of lead citrate for 2 minutes. Remove the grid and delicately rinse in 3 successive baths of double distilled water. Lastly, dry the grids on a filter paper and store them in a grid box until electron microscopic examination (Figures 3-6).</li>
</ol>
4. Representative Results
<img src="/files/ftp_upload/2021/2021fig1.jpg" alt="Figure 1" /> 
 Figure 1. Iba1-stained section at the light microscopic level. The cellular distribution of Iba1 is restricted to microglia, which shows the specificity of the immunostaining. Scale bar=100 &mu;m.
<img src="/files/ftp_upload/2021/2021fig2.jpg" alt="Figure 2" /> 
 Figure 2. Toluidine blue-stained semi-thin section at the light microscopic level. Note the border between tissue and resin where the immunostaining will appear most intense at the electron microscope.
<img src="/files/ftp_upload/2021/2021fig3.jpg" alt="Figure 3" /> 
 Figure 3. Ultrathin section showing Iba1-immunopositive microglial perikaryon and processes at the electron microscopic level. Iba1-positive structural elements are recognized by their immunoperoxidase-DAB electron-dense precipitate. Scale bar=2 &mu;m.
<img src="/files/ftp_upload/2021/2021fig4.jpg" alt="Figure 4" /> 
 Figure 4. Ultrathin section displaying Iba1-immunopositive microglial processes of different sizes and shapes at the electron microscopic level. Scale bar=2 &mu;m.
<img src="/files/ftp_upload/2021/2021fig5.jpg" alt="Figure 5" /> 
 Figure 5. Ultrathin section showing at higher magnification an Iba1-positive microglial process from Figure 4, enabling identification of its intracellular organelles. Scale bar=1 &mu;m.
<img src="/files/ftp_upload/2021/2021fig6.jpg" alt="Figure 6" /> 
 Figure 6. Ultrathin section revealing at higher magnification the ultrastructural relationships between an Iba1-positive microglial process and nearby elements of neuropil including synapses. Scale bar=1&mu;m.

<ol>
	<li>Venable, J. H., Coggeshall, 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+simplified+lead+citrate+stain+for+use+in+electron+microscopy.">A simplified lead citrate stain for use in electron microscopy.</a> J Cell Biol. 25, 407-407 (1965).</li><li>Riad, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Somatodendritic+localization+of+5-HT1A+and+preterminal+axonal+localization+of+5-HT1B+serotonin+receptors+in+adult+rat+brain.">Somatodendritic localization of 5-HT1A and preterminal axonal localization of 5-HT1B serotonin receptors in adult rat brain.</a> J Comp Neurol. 417 (2), 181-181 (2000).</li><li>Tremblay, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localization+of+EphA4+in+axon+terminals+and+dendritic+spines+of+adult+rat+hippocampus.">Localization of EphA4 in axon terminals and dendritic spines of adult rat hippocampus.</a> J Comp Neurol. 501 (5), 691-691 (2007).</li><li>Tremblay, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+course+of+EphA4+cellular+and+subcellular+localization+in+the+postnatal+rat+hippocampus.">Developmental course of EphA4 cellular and subcellular localization in the postnatal rat hippocampus.</a> J Comp Neurol. 512 (6), 798-798 (2009).</li><li>Bouvier, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pre-synaptic+and+post-synaptic+localization+of+EphA4+and+EphB2+in+adult+mouse+forebrain.">Pre-synaptic and post-synaptic localization of EphA4 and EphB2 in adult mouse forebrain.</a> J Neurochem. 106 (2), 682-682 (2008).</li><li>Bouvier, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EphA4+is+localized+in+clathrin-coated+and+synaptic+vesicles+in+adult+mouse+brain.">EphA4 is localized in clathrin-coated and synaptic vesicles in adult mouse brain.</a> J Neurochem. , (2010).</li></ol>

No conflicts of interest declared.

Here we have described a protocol for preparation of mouse brain tissue for immunoelectron microscopy that provides an excellent compromise between optimal ultrastructural preservation and immunocytochemical detection, when the procedures are rigorously performed.
Combined with TEM, this method enables to distinguish cellular elements with few distinctive features and identification criteria. In particular, the immunoperoxidase-DAB staining of Iba1 enables to identify microglial perikarya and processes, within the neuropil, as well as to analyze their morphology, intracellular organelles, and ultrastructural relationships with other cellular elements (Figures 3-6). Additionally, this protocol allows analyzing the ultrastructural locations of soluble or membrane-bound proteins in glial or neuronal elements, as well as their association with intracellular organelles. Indeed, using this protocol with specific antibodies, we have previously revealed the ultrastructural localization of 5-HT1A and 5-HT1B serotonergic receptors in the neuronal perikarya, dendrites, dendritic spines, unmyelinated axons, and endothelial cells of adult rat raphe dorsalis nucleus, substantia nigra, globus pallidus, and hippocampus2. We have also shown the ultrastructural localization of EphA4 and EphB2 tyrosine kinase receptors in clathrin-coated vesicles and different astrocytic and neuronal elements in mouse and rat hippocampus and cerebral cortex, during postnatal development and adulthood3,4,5,6. Lastly, this protocol can be combined with pre-embedding immunogold to analyze the ultrastructural relationships between two simultaneously labeled proteins, for example the co-localization of Vglut1 glutamatergic transporter and EphA4 receptor in axon terminals of adult mouse cerebral cortex6. Therefore, this technique can be successfully applied, alone or in combination with immunogold labeling, for the ultrastructural analysis of various proteins within different regions and cell types of the central nervous system, throughout postnatal life, in health and disease.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Sodium pentobarbital (Nembutal)</td>
<td>&nbsp;</td>
<td>Ovation Pharmaceuticals</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Filter paper 315 24 cm</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>28331-081</td>
<td>&nbsp;</td>
<tr>
<td>*Acrolein purum (&ge;95%)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich (Fluka)</td>
<td>01680</td>
<td>&nbsp;</td>
<tr>
<td>Sodium borohydride (&ge;98%)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich (Aldrich)</td>
<td>452173</td>
<td>&nbsp;</td>
<tr>
<td>*Prilled paraformaldehyde (PFA) (95%)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>441244</td>
<td>&nbsp;</td>
<tr>
<td>Normal goat serum</td>
<td>&nbsp;</td>
<td>Jackson ImmunoResearch</td>
<td>005-000-121</td>
<td>&nbsp;</td>
<tr>
<td>Rabbit anti-Iba1 primary antibody</td>
<td>&nbsp;</td>
<td>Wako</td>
<td>019-19741</td>
<td>Stored at -20&deg;C, 1:1 in glycerol</td>
</tr>
<tr>
<td>Biotin-SP-conjugated AffiniPure F(ab&#x2019;)2 fragment goat anti-rabbit IgG, Fc fragment specific</td>
<td>&nbsp;</td>
<td>Jackson ImmunoResearch</td>
<td>111-066-046</td>
<td>Stored at -20&deg;C, 1:1 in glycerol</td>
</tr>
<tr>
<td>Peroxidase-conjugated streptavidin</td>
<td>&nbsp;</td>
<td>Jackson ImmunoResearch</td>
<td>016-030-084</td>
<td>Stored at -20&deg;C, 1:1 in glycerol</td>
</tr>
<tr>
<td>*Osmium tetroxide 4% solution</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>19150</td>
<td>Light sensitive</td>
</tr>
<tr>
<td>Propylene oxide (&ge;99%)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich (Fluka)</td>
<td>82325</td>
<td>&nbsp;</td>
<tr>
<td>Polypropylene disposable beakers (50 mL)</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>01-291-10</td>
<td>&nbsp;</td>
<tr>
<td>Aluminium weigh dishes (70 mm)</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>NC9261784</td>
<td>&nbsp;</td>
<tr>
<td>Durcupan epoxy resin</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>14040</td>
<td>&nbsp;</td>
<tr>
<td>Embedding mold</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>70907</td>
<td>&nbsp;</td>
<tr>
<td>DPX mountant for histology</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich (BioChemika)</td>
<td>44581</td>
<td>&nbsp;</td>
<tr>
<td>Gelatin subbed slides</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>100241-864</td>
<td>&nbsp;</td>
<tr>
<td>ACLAR embedding films (7.8 mm)</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>50425</td>
<td>&nbsp;</td>
<tr>
<td>Capsule mold</td>
<td>&nbsp;</td>
<td>Electron microscopy Sciences</td>
<td>70150</td>
<td>&nbsp;</td>
<tr>
<td>High-performance super glue</td>
<td>&nbsp;</td>
<td>Corporate Express</td>
<td>LOC30379</td>
<td>&nbsp;</td>
<tr>
<td>Perfect loop for ultra thin sections</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>70944</td>
<td>&nbsp;</td>
<tr>
<td>Superfrost slides</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>22-178-277</td>
<td>&nbsp;</td>
<tr>
<td>Diamond knife ultra 45&deg; (2.5-3.5 mm)</td>
<td>&nbsp;</td>
<td>Diatome</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Gelatin from cold water fish skin (~45%)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich (Sigma)</td>
<td>G7765</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>
			<div>
			*One should always work under a fume hood and wear nitrile gloves when handling acrolein, paraformaldehyde, and osmium and should also dispose of their waste properly.		</div>

preparation of mouse brain tissue for immunoelectron microscopy

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments (dendrite, dendritic spine, axon, axon terminal, perikaryon), as well as their intracellular organelles and cytoskeleton, in the central nervous system at high spatial resolution. Combined with TEM, pre-embedding immunocytochemistry allows the discrimination of cellular elements with few distinctive features and identification criteria (e.g., microglial perikarya and processes, when using an antibody against the microglia-specific marker Iba1 (ionized calcium binding adaptor molecule 1; as presented here)), identifying the neurotransmitter contents of cellular elements (e.g., serotonergic) and their ultrastructural localization of soluble or membrane-bound proteins (e.g., 5 HT1A and EphA4 receptors). Here, we describe a protocol for transcardiac perfusion of mice with acrolein fixative, removal and sectioning of the brain, as well as immunoperoxidase-diaminobenzidine (DAB) staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with TEM. When rigorously performed, this technique provides an excellent compromise between optimal ultrastructural preservation and immunocytochemical detection.

Watch this Scientific Journal Video about Preparation of Mouse Brain Tissue for Immunoelectron Microscopy at JoVE.com

Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

We describe a protocol for transcardiac perfusion of mice, removal and sectioning of the brain, as well as immunoperoxidase staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with transmission electron microscopy.

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments ...

preparation-of-mouse-brain-tissue-for-immunoelectron-microscopy

Nanyang Technological University

Research

JoVE Journal

Neuroscience

36.8K Views.  University of Rochester. We describe a protocol for transcardiac perfusion of mice, removal and sectioning of the brain, as well as immunoperoxidase staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with transmission electron microscopy.

Video: Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

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 ...

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the organ before imaging.1,2 However, for organs that contain fluorescent proteins such as GFP and YFP, optical clearing protocols that use methanol dehydration and clear using benzyl alcohol:benzyl benzoate (BABB) while unprotected from light3 do not preserve the fluorescent signal. The protocol presented here is a novel way in which to perform whole organ optical clearing on mouse brain while preserving the fluorescence signal of YFP expressed in neurons. Altering the optical clearing protocol such that the organ is dehydrated using an ethanol graded series has been found to reduce the damage to the fluorescent proteins and preserve their fluorescent signal for multiphoton imaging.4 Using an optimized method of optical clearing with ethanol-based dehydration and clearing by BABB while shielded from light, we show high-resolution multiphoton images of yellow fluorescent protein (YFP) expression in the neurons of a mouse brain more than 2 mm beneath the tissue surface.

Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the ...

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

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 ...

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 ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.

Here, we provide an easy, low-cost, and time-efficient protocol to chemically fix primate brain tissue with acrolein fixative, allowing for long-term preservation that is compatible with pre-embedding immunohistochemistry for transmission electron microscopy.

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and ...

Preparation of Newborn Rat Brain Tissue for Ultrastructural Morphometric Analysis of Synaptic Vesicle Distribution at Nerve Terminals

Our laboratory and many others have exploited the high resolving power of transmission electron microscopy to study the morphology and spatial organization of synaptic vesicles. In order to obtain high-quality electron micrographs that can yield the degree of morphological detail necessary for quantitative analysis of pre-synaptic vesicle distribution, optimal specimen preparation is critical. Chemical fixation is the first step in the process of specimen preparation, and of utmost importance to preserve fine ultrastructure. Vascular fixation with a glutaraldehyde-formaldehyde solution, followed by treatment of vibratome-sectioned specimens with osmium tetroxide, stabilizes the maximum number of molecules, especially proteins and lipids, and results in superior conservation of ultrastructure. Tissue is then processed with counterstaining, sequential dehydration and resin-embedding. En bloc staining with uranyl acetate (i.e., staining of vibratome-sectioned tissue before resin embedding) enhances endogenous contrast and stabilizes cell components against extraction during specimen processing. Contrast can be further increased by applying uranyl acetate as a post-stain on ultrathin sections. Double-staining of ultrathin sections with lead citrate after uranyl acetate treatment also improves image resolution, by intensifying electron-opacity of nucleic acid-containing structures through selective binding of lead to uranyl acetate. Transmission electron microscopy is a powerful tool for characterization of the morphological details of synaptic vesicles and quantification of their size and spatial organization in the terminal bouton. However, because it uses fixed tissue, transmission electron microscopy can only provide indirect information regarding living or evolving processes. Therefore, other techniques should be considered when the main objective is to study dynamic or functional aspects of synaptic vesicle trafficking and exocytosis.

We describe procedures for processing newborn rat brain tissue to obtain high-resolution electron micrographs for morphometric analysis of synaptic vesicle distribution at nerve terminals. The micrographs obtained with these methods can also be used to study the morphology of a number of other cellular components and their dimensional structural relationships.

Our laboratory and many others have exploited the high resolving power of transmission electron microscopy to study the morphology and spatial ...

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered tissue samples. Traditional immunolabeling procedures require cutting the sample into thin sections, which restricts the ability to label and examine intact structures. However, if brain tissue can remain intact during processing, structures and circuits can remain intact for the analysis. Previously established clearing methods take significant time to completely clear the tissue, and the harsh chemicals can often damage sensitive antibodies. The iDISCO method quickly and completely clears tissue, is compatible with many antibodies, and requires no special lab equipment. This technique was initially validated for the use in mice tissue, but the current protocol adapts this method to image hemispheres of control and transgenic rat brains. In addition to this, the present protocol also makes several adjustments to preexisting protocol to provide clearer images with less background staining. Antibodies for Iba-1 and tyrosine hydroxylase were validated in the HIV-1 transgenic rat and in F344/N control rats using the present hydrophobic tissue clearing method. The brain is an interwoven network, where structures work together more often than separately of one another. Analyzing the brain as a whole system as opposed to a combination of individual pieces is the greatest benefit of this whole brain clearing method.

Here we present a hydrophobic tissue clearing method that allows for the viewing of target molecules as part of intact brain structures. This technique has now been validated for F344/N control and HIV-1 transgenic rats of both sexes.

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered ...

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.

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.

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

Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

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Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Preparation of Newborn Rat Brain Tissue for Ultrastructural Morphometric Analysis of Synaptic Vesicle Distribution at Nerve Terminals

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

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