This study was supported by the Natural Sciences and Engineering Research Council of Canada (NSERC, 401848-2011 to M.P.). M.P received a career award from the Fonds de recherche du Qu&#233;bec-Sant&#233; (FRQ-S). L.E. was the recipient of a doctoral fellowship from the FRQ-S (FRQ-S 14D 29441). We thank Marie-Jos&#233;e Wallman for technical assistance.

Ethics Statement: All protocols involving animals were approved by the Comit&#233; de Protection des Animaux de l&#39;Universit&#233; Laval and were made in accordance with the Canadian Council on Animal Care&#39;s Guide to the Care and Use of Experimental Animals (Ed. 2). The protocol described here was optimized for adult animals of approximately 800 g. The volumes of fixative should be adjusted according to the animal&#39;s size.
1. Preparation of Solutions for Transcardiac Perfusion
<ol>
	<li>Prepare 1 L of a 50 mM sodium Phosphate-buffered Saline (PBS) solution according to the following steps. Prepare the solution at most 24 h before perfusion and keep at 4 &#176;C until use.
	<ol>
		<li>Measure 800 mL of distilled water in a 1 L beaker. Add 5.87 g of dibasic anhydrous sodium phosphate (Na2HPO4), 1.20 g of monobasic monohydrate sodium phosphate (NaH2PO4&#183;H2O), and 9 g of sodium chloride (NaCl). Stir to dissolve.</li>
		<li>Adjust the pH to 7.4 by gradually adding 5 N NaOH. Add distilled water to reach a total volume of 1 L. 
		CAUTION: NaOH is a corrosive chemical compound. Wear the appropriate Personal Protective Equipment (PPE; laboratory coat, gloves, protective goggles, etc.).</li>
	</ol>
	</li>
	<li>Prepare 2 L of 4% paraformaldehyde (PFA) according to the following steps. This solution should be prepared at most 24 h before surgery and kept at 4 &#176;C until use.
	<ol>
		<li>Measure 1.5 L of distilled water in a 2-L beaker. Add 23.48 g of dibasic anhydrous sodium phosphate (Na2HPO4) and 4.80 g of monobasic monohydrate sodium phosphate (NaH2PO4&#183;H2O). Stir to dissolve. Add distilled water to reach a total volume of 2 L.</li>
		<li>Heat the solution under a venting hood until the temperature reaches approximately 45 - 55 &#176;C. Do not heat to over 60 &#176;C.</li>
		<li>Gradually add 80 g of PFA to the solution and stir until completely dissolved (approximately 30 - 60 min). Keep monitoring the temperature to keep it below 60 &#176;C. 
		CAUTION: PFA is highly volatile in its powder form. It is highly toxic if in contact with eyes or skin and is hazardous in case of inhalation or ingestion. Wear PPE and use with caution under a venting hood.</li>
		<li>Cool the solution down to 4 &#176;C and filter. Store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 1 L of 3% acrolein in 0.1 M Phosphate Buffer (PB) according to the following steps. The PB solution should be prepared at most 24 h prior to perfusion. 
	CAUTION: Acrolein is highly toxic if inhaled and can produce immediate damage. It is also corrosive and highly toxic if absorbed through the skin. It is carcinogen and mutagenic. Wear the appropriate PPE and use under a venting&#160;hood.
	<ol>
		<li>Measure 800 mL of distilled water. Add 8.66 g of dibasic anhydrous sodium phosphate (Na2HPO4) and 5.38 g of monobasic monohydrate sodium phosphate (NaH2PO4&#183;H2O). Stir until all the salts dissolve. Add distilled water to reach a total volume of 1 L. Keep the solution at 4 &#176;C.</li>
		<li>Before surgery, transfer 900 mL of the PB solution to a 1 L glass container and add 30.94 mL of 97% acrolein solution under a venting hood. Add PB solution to reach a total volume of 1 L and stir. Filter the solution and keep it at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Transcardiac Perfusion and Brain Dissection
<ol>
	<li>Keep solutions on ice during the entire surgical procedure. Prepare the pump by placing the tube in the first solution to be used (PBS; 50 mM) and turning the pump on until no air remains in the hose. For the transcardiac perfusion of a macaque monkey, use a 21 G needle and set the outflow at approximately 80 mL/min.</li>
	<li>Anesthetize the animal with an intramuscular injection of a mixture of ketamine (20 mg/kg), xylazine (4 mg/kg), and acepromazine (0.5 mg/kg). Maintain the animal under isoflurane (3%) sedation.</li>
	<li>Attach the animal&#39;s limbs to a venting table.</li>
	<li>With a scalpel, remove the skin up to the armpits. Cut the abdominal muscles. Use heavy-duty surgical scissors to cut the ribs laterally by carefully avoiding the vital organs.</li>
	<li>Cut the diaphragm with surgical scissors and raise the rib cage to expose the heart. Once the diaphragm is cut, proceed quickly, since the heart will stop beating within minutes.</li>
	<li>Remove the pericardium with a scalpel blade and insert the needle into the left ventricle. With a scalpel, carefully make a small excision to the right atrium. Quickly start the pump at 72 mL/min and hold the needle in place. If possible, tilt the animal in order to have its head on a lower level than the heart. 
	NOTE: Be careful not to pierce the interventricular septum when inserting the needle.
	<ol>
		<li>Rinse the blood with approximately 300 mL of PBS until the lungs are white and no blood comes out of the right atrium.</li>
	</ol>
	</li>
	<li>Stop the pump, quickly transfer the hose/tube to the 3% acrolein solution, and start the pump again. Gradually increase the pumping speed to 80 mL/min once the heart has stopped beating. Perfuse approximately 500 mL of the acrolein solution.</li>
	<li>Stop the pump, quickly transfer the hose to the 4% PFA solution, and start the pump again. This step requires approximately 1 L of PFA. 
	NOTE: The fixation is complete when the forelimbs are rigid and the neck is stiff.</li>
	<li>Cut the head and carefully dissect the brain out of the skull. Take care to not damage the brain with the surgical instruments. The perfusion is optimal when the brain is pale (no trace of blood) and rigid (Figure 1A).</li>
	<li>Immerse the intact brain in 4% PFA for 1 h at 4 &#176;C.</li>
	<li>Serially cut the brain with a cooling vibratome (4 &#176;C) in the desired plane into 50 &#181;m thick sections and collect them in PBS (0.1 M) (Figure 1B). 
	NOTE: This step can be carried out in many ways according to the desired protocol. Hemispheres can be separated prior to vibratome cutting, or kept whole. If the brain is too large for the vibratome platform, it can be cut into smaller blocks. After this step, sections can be stored for a long period of time at -30 &#176;C in an antifreeze solution made of 40% PB (50 mM), 30% glycerol, and 30% ethylene glycol.</li>
</ol>
3. Pre-embedding Immunohistochemistry (Figure 1C)
<ol>
	<li>Prepare a 4 L stock solution of Tris-buffered Saline (TBS, 50 mM, pH 7.6) as follows.
	<ol>
		<li>Measure 2L of distilled water in a 4 L beaker, add 24.23 g of trihydroxymethyl aminomethane (THAM; C4H11NO3), and stir to dissolve.</li>
		<li>Adjust the pH to 7.6 with approximately 148 mL of 1 N HCl. The acid should be added with caution to avoid reaching a pH below 7.6. The total volume should be 4 L. 
		CAUTION: HCl is highly corrosive. Wear the appropriate PPE.</li>
	</ol>
	</li>
	<li>Select sections (from step 2.11) containing the region of interest to be processed for EM immunohistochemistry.</li>
	<li>Wash the free-floating sections 3x&#160;in PBS (0.1 M, pH 7.4) for 5 min at RT&#160;to rinse the antifreeze solution.</li>
	<li>Prepare a 0.5% solution of NaBH4 diluted in PBS.
	<ol>
		<li>Weigh 0.05 g of NaBH4 and dilute it in 10 mL of PBS. Do not cover. Prepare this solution just before use.</li>
	</ol>
	</li>
	<li>Incubate the sections in the freshly prepared NaBH4 solution for 30 min at RT. Rock gently. Do not cover.</li>
	<li>Wash 3x&#160;in PBS for 10 min at RT, rocking vigorously until none of the reaction gas remains.</li>
	<li>Prepare a blocking solution for EM with 2% appropriate normal serum and 0.5% cold fish gelatin diluted in PBS. 
	NOTE: The quantity should be calculated in order to have enough for the following three steps. Use a serum made from the same animal species hosting the secondary antibody.&#160;Avoid the use of antigen retrieval methods or addition of small amounts of triton (to increase the penetration of antibodies), as these methods significantly compromise the quality of the tissue.</li>
	<li>Incubate the sections in the blocking solution for 1 h at RT. Rock gently.</li>
	<li>Prepare primary antibody solution diluted in the blocking solution. 
	NOTE: The concentration of the primary antibody is usually the same as for LM immunohistochemistry, but consider conducting tests with different antibody concentrations beforehand, as some primary antibodies may not work with acrolein. If so, it is possible to use a mix of glutaraldehyde (0.1 - 2%) and 4% PFA for transcardiac perfusion and obtain similar results. Optimize the incubation time and temperature as well.</li>
	<li>Incubate the sections in primary antibody solution overnight at RT with gentle rocking. Use a previously tested incubation time and temperature.</li>
	<li>Wash the sections 3x in PBS for 5 min at RT with gentle rocking.</li>
	<li>Prepare a 1:1,000 solution of biotinylated secondary antibody diluted in the blocking solution. 
	NOTE: The secondary antibody must be raised against the host species used to generate the primary antibody.</li>
	<li>Incubate the sections in secondary antibody solution for 1.5 h at RT. Rock gently.</li>
	<li>Prepare an avidin-biotin-peroxidase (ABC) solution at least 60 min prior to the end of the secondary antibody incubation.
	<ol>
		<li>Use a calibrated pipette to measure 8.80 &#181;L/mL of solutions A and B and dilute them in PBS.</li>
		<li>Rock mildly for at least 60 min at RT to allow the complete binding between the avidin and biotin molecules.</li>
	</ol>
	</li>
	<li>After the incubation in secondary antibody solution, wash 3x in PBS for 10 min at RT.</li>
	<li>Incubate the sections in the ABC solution for 1 h at RT. Rock gently.</li>
	<li>Wash once in PBS and twice in TBS for 10 min at RT with gentle rocking.</li>
	<li>Prepare a fresh solution of 0.05% 3,3&#39;-diaminobenzidine (DAB) with 0.005% H2O2 diluted in TBS.
	<ol>
		<li>Weigh 12.5 mg of DAB and dilute it in 25 mL of cold TBS. Protect from light. 
		CAUTION: The DAB powder is highly volatile and harmful if inhaled. It is carcinogenic and teratogenic. Thus, pregnant or nursing women should not manipulate this product, even when diluted. Use a N95 mask when manipulating and wear PPE.</li>
		<li>Filter the solution and add 4.5 &#181;L of 30% H2O2 just before use.</li>
	</ol>
	</li>
	<li>Incubate sections in the DAB solution for 3 to 7 min at RT. Rock gently. 
	NOTE: The brown precipitate should not be too dark in order to avoid a high level of background staining. The incubation time should be optimized accordingly.</li>
	<li>Stop the reaction by quickly washing twice in cold TBS, then twice for 10 min in cold TBS at RT, followed by twice for 10 min in PB, with mild rocking. 
	NOTE: The use of PB (and not PBS) is critical in order to eliminate any traces of NaCl, as it would react with osmium and form crystals.</li>
</ol>
4. Osmification and Embedding for Electron Microscopic Observation
<ol>
	<li>Prepare a solution of 1% osmium tetroxide (OsO4) diluted in PB. Protect from light. 
	CAUTION: Osmium is highly toxic and should not come in contact with the skin, eyes, or mouth and should not be inhaled. It can cause death if ingested. It should only be used under a venting hood and with appropriate PPE.</li>
	<li>Incubate the sections in OsO4 solution for 30 min at RT under the venting hood (without agitation) and cover them with aluminum foil to protect them from light. Completely flatten the sections prior to adding the OsO4 solution. 
	NOTE: The sections become very dark and rigid and should be manipulated with care following this step.</li>
	<li>Prepare water-repellent epoxy resin during the osmification.
	<ol>
		<li>Add appropriate amounts of each component of the epoxy resin mix (20 g of epoxy resin, 20 g of hardener, 0.6 g of accelerator and 0.4 g of plasticizer) to a large plastic cup. Stir with a wooden stick or plastic pipette until a homogeneous brown color is obtained. 
		NOTE: It is critical to use the exact proportion of each component.</li>
		<li>Transfer equal quantities to aluminum cups of appropriate sizes, depending on the number of sections to be processed. Allow it to rest.</li>
	</ol>
	</li>
	<li>Wash the osmificated sections 3x in PB for 10 min at RT with low-speed rocking.</li>
	<li>Dehydrate the sections in the following series of graded ethanol for 2 min each: 2 times in 35% ethanol; 1 time each in 50, 70, 80, 90, and 95% ethanol; and 3x&#160;in 100% ethanol.</li>
	<li>Transfer the sections to glass vials to complete the dehydration process by incubating the sections 3 times for 2 min in propylene oxide. 
	CAUTION: Propylene oxide is a highly volatile and toxic organic solvent. It can cause serious damage in the eyes or skin in case of contact or if inhaled or ingested. It has been classified as a grade 2 carcinogenic substance. It should only be used under a venting hood and with PPE. It is also highly flammable and should be kept away from any heat source. 
	NOTE: Prior to this step, sections should be carefully transferred in glass vials, since propylene oxide is an organic solvent and is incompatible with plastic. At this point, sections are very fragile and should be manipulated with care. Sections can be alternatively transferred to glass vials prior to step 4.5.</li>
	<li>Transfer sections carefully, one by one, in the aluminum cups and avoid contact with air as much as possible. Flat-embed the sections in previously mixed water-repellent epoxy resin and incubate them overnight under the venting hood at RT. 
	NOTE: At this step, the sections are completely dehydrated and very fragile and should be manipulated with care.</li>
	<li>Using mineral oil, prepare grease-coated glass slides along with greased plastic coverslips.</li>
	<li>Soften the resin by incubating aluminum cups at 60 &#176;C for 12-15 min at most. Carefully flatten the sections on the greased side of the glass slide. Place the greased coverslip and carefully push out any remaining air.</li>
	<li>Incubate the slides at 60 &#176;C for 48 h. 
	NOTE: It is critical to not exceed 48 h incubation time, as the resin will become too hard.</li>
	<li>Remove the plastic coverslip.</li>
</ol>
5. Sample Preparation for Ultrathin Sectioning and Observation Using a Transmission Electron Microscope
<ol>
	<li>Use binoculars to find the region of interest and cut a small quadrangular piece of approximately 1 mm2 with a scalpel.</li>
	<li>File the tip of a resin block and glue the quadrangular piece onto it (Figure 1C). Allow the glue to dry for at least 1 h or overnight prior to sectioning.</li>
	<li>Using an ultramicrotome, cut the quadrangular piece into 80 &#181;m-thick sections (Figure 1D).
	<ol>
		<li>Place the resin block in an ultramicrotome deck in the vertical position and, using a sharp razor blade, gradually cut each side of the resin block to form a trapezoid with smooth sides.</li>
		<li>Put the deck in its horizontal position and rotate the block until the longest side of the trapezoid is facing downward.</li>
		<li>Use a diamond trimming tool or a glass knife to trim the surface of the quadrangular piece. Set the ultramicrotome to cut 300 &#181;m thick sections at 1 mm/s. Adjust the knife to be vertically parallel to the quadrangular piece and to display a very small horizontal angle of approximately 1&#176;. 
		NOTE: This angle will allow the user to approach the tissue almost parallel to the surface of the block, where immunolabeled elements are more likely to be found. When using these parameters with the diamond trimming tool, the resin displays a shiny white color. When the tissue is being cut, it changes to a purple or greenish color.&#160;</li>
		<li>Use an ultra 45&#176; diamond knife equipped with a boat filled with distilled water to cut 80 &#181;m thick sections,&#160;smoothen sections by passing over them with a piece of absorbing paper tipped in xylene, and collect serial sections on formvar-coated nickel slot grids or bare 150 mesh copper grids (Figure 1E).</li>
	</ol>
	</li>
	<li>Place the grids in a grid storage box.</li>
	<li>Stain the grids with lead citrate.
	<ol>
		<li>Use a 5 mL syringe and a 0.2 &#181;m syringe filter to prepare a 1:1 solution of filtered lead citrate stock solution and filtered distilled water. Protect it from light. 
		NOTE: The stock solution of lead citrate should be made fresh every month to avoid the formation of solid deposits. See the material data sheet for the stock recipe. In addition, if many series of grids need to be stained, change the diluted solution when it becomes milky.</li>
		<li>Place each grid onto a drop of the diluted solution, with the section in contact with the solution. Incubate for 3 min.</li>
		<li>Use small tweezers to hold the grid, and thoroughly rinse it in two beakers containing distilled water.</li>
		<li>Remove excess water by gently using absorbing paper. Store the grids in a grid box. Wait 30 min before examining the sections by Transmission Electron Microscopy (TEM; Figure 1F).</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

In this article, we present a reliable protocol for transcardiac perfusion of non-human primates and pre-embedding immunohistochemistry suitable for EM sample examination. Although typical cryo-EM, such as CEMOVIS, provides a good preservation of brain ultrastructure, it also limits the use of immunohistochemistry12. Other techniques, including cryo-substitution and Tokuyaso technique, allow post-embedding immunohistochemistry, but these techniques are expensive due to additional devices needed du...

Light microscopy, including confocal and two-photon microscopy, has proven to be an efficient tool for studying in vivo neuronal processes, among other things1,2. Although the typical spatial resolution at the Light Microscopic (LM) level is approximately 200 nm, recent technological advances using different light sources, such as extreme ultraviolet and soft X-ray microscopy, have notably increased this resolution to a nearly 10 nm spatial resolution3,4,5. Other technological advances in imaging include combined magnetic resonance imaging with histology and provide a novel method for measuring the thickness of the myelin sheath in vivo, a parameter that was traditionally measurable only at the Electron Microscopic (EM) level6,7. Although these advances at the LM level provide an excellent tool for studying living processes, a detailed view and characterization of structures, such as synaptic contacts, can only be achieved with EM, which offers a resolution that can reach 0.5 nm. However, observation at the EM level requires the specimens to be dead and altered in some ways, with chemical fixatives and dehydration processes, in order to preserve the cytoarchitecture. Thus, examining biological samples at high resolution can be challenging due to radiation damage from the electron beam, low contrast, structural deviations of membranes, or even the presence of artifacts that can occur following dehydration and epoxy embedding8,9,10.
Preserving specimens in their native form for structural analysis can be achieved by using &#34;Cryo EM of Vitrified Sections&#34; or CEMOVIS, a sectioning approach that involves rapidly freezing and embedding the sample in vitreous ice and examining the sections under the EM at a cryogenic temperature11,12. This procedure allows for the examination of samples while they are still solid and fully hydrated, thus eliminating artifacts caused by dehydration processes13. However, this method involves additional devices for cryo-ultramicrotomy, as well as additional devices on the standard EM, in order to allow this observation at very low temperatures, which generate significant additional costs. In addition, the CEMOVIS approach precludes the use of immunolabeling techniques, since antibodies usually have to be incubated at RT. Alternatively, it is possible to combine ultrastructural analysis with immunohistochemical procedures by using a freeze-substitution approach, during which cryo-fixed specimens are slowly thawed while immerged in cryo-protective chemicals and are then embedded in specialized resins, such as Lowicryls. Post-embedding immunolabeling can then be performed on such material12. However, freeze-substitution and cryo-fixation techniques are time consuming. They require the installation of additional equipment and still require samples to be exposed to organic solvent and chemical fixative that can alter the cytoarchitecture, despite the use of a low temperature14,15. Hence, despite all the technological advances both at the LM and EM level, chemical fixation of brain tissue, particularly with acrolein, remains a low-cost and time-efficient method to combine immunohistochemistry with EM16.
In the last decades, many attempts were made to find a mixture of aldehydes that provide the best tissue preservation. Before the 1960s, the only chemical fixative that gave acceptable results for EM was osmium tetroxide. However, osmium tetroxide is highly toxic and expensive, precluding its use through the vascular system to fix organs such as the brain. Acrolein was introduced in the late 1950s as a reliable method for animal tissue preservation suitable for EM observation of cellular structures17. It penetrates the tissue more deeply and reacts more quickly than other aldehydes when used for fixation by immersion and allows good preservation of cytoplasmic components, with minimal shrinkage of the tissue17. Such a feature gives acrolein fixation a clear advantage over other aldehydes when used in fresh tissue, by allowing a more accurate localization of living molecular compounds, such as enzymes and other proteins18. Indeed, it has been validated through the years as an easy, efficient and low-cost method of fixation for visualization at the EM level in many species, including amphibians and rodents, as it efficiently stabilizes peptides and proteins, retains antigenicity and provides relatively intact ultrastructure when used in combination with another aldehyde fixative16,18,19,20,21. Protocols for acrolein fixation in rodents have since then been standardized and used extensively, particularly by the Pickel group, to implement dual immunolabeling for EM16,22. A few groups have used acrolein fixation in non-human primate brain tissue23. However, to our knowledge, there is only one published protocol efficiently describing chemical fixation with acrolein in non-human primates that is compatible with EM immunolabeling24.
In this article, we provide an easy and reliable method to efficiently chemically fix non-human primate brains with acrolein, allowing for a potentially long-term preservation along with pre-embedding immunolabeling and transmission EM examination.

<table><tbody><tr><td>Dibasic anhydrous sodium phosphate (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Fisher scientific</td><td>S374-500</td><td></td></tr><tr><td>Monobasic monohydrate sodium phosphate (NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;&amp;#183;H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>EM Science</td><td>SX0710-1</td><td></td></tr><tr><td>Sodium chloride (NaCl)</td><td>Fisher scientific</td><td>S271-3</td><td></td></tr><tr><td>Hydroxymethyl aminomethane (THAM)</td><td>Fisher scientific</td><td>T370-500</td><td></td></tr><tr><td>HCl&amp;nbsp;</td><td>EMD</td><td>HX0603-3</td><td>1 N dilution. Product is corrosive. Use with appropriate protection.</td></tr><tr><td>NaOH</td><td>EMD</td><td>SX0590-1</td><td>5 N dilution. Product is corrosive. Use with appropriate protection.</td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Sigma</td><td>P6148</td><td>4% dilution. Product is highly volatile in its powder form and highly toxic. Use with caution under a venting hood with appropriate protection.</td></tr><tr><td>Acrolein (90%)</td><td>Sigma</td><td>110221</td><td>3% dilution. Product is highly toxic. Use under a venting hood with appropriate protection.</td></tr><tr><td>Autopsy venting table</td><td>Mopec</td><td>CE400</td><td></td></tr><tr><td>Electronic perfusion pump</td><td>cole parmer</td><td>masterflex L/S 7523-90</td><td></td></tr><tr><td>Needle (perfusion)</td><td>terumo&amp;nbsp;</td><td>NN-1838R</td><td>18 G 11/2</td></tr><tr><td>Needle</td><td>terumo&amp;nbsp;</td><td>NN-2713R</td><td>21 G 1/2</td></tr><tr><td>Ketamine</td><td></td><td></td><td>20 mg/kg</td></tr><tr><td>Xylazine</td><td></td><td></td><td>4 mg/kg</td></tr><tr><td>Acepromazine</td><td></td><td></td><td>0.5 mg/kg</td></tr><tr><td>Scalpel</td><td></td><td></td><td></td></tr><tr><td>Scalpel blades</td><td>Feather&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; lance</td><td>201011&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; J9913</td><td>No.22 for surgery and No. 11 for EM</td></tr><tr><td>Surgical scissors</td><td></td><td></td><td></td></tr><tr><td>Rongeurs</td><td></td><td></td><td></td></tr><tr><td>Vibratome</td><td>Leica</td><td>VT 1200S</td><td>Calibrate blade before each use, when the device allows it</td></tr><tr><td>Vibratome razor blade</td><td>Gillette</td><td>GIN 642107</td><td></td></tr><tr><td>Glycerol</td><td>Fisher scientific</td><td>G33-4</td><td>30% dilution</td></tr><tr><td>Ethylene glycol</td><td>Fisher scientific</td><td>E178-4</td><td>30% dilution</td></tr><tr><td>Sodium borohydride (NaBH&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma</td><td>S-9125</td><td></td></tr><tr><td>Normal horse serum (NHS)</td><td>Jackson immunoResearch Laboratories</td><td>008-000-121</td><td>2% dilution</td></tr><tr><td>Cold-fish gelatin</td><td>Aurion</td><td>900.033</td><td>0.5% dilution. Original product is concentrated at 40%</td></tr><tr><td>Primary antibody, SERT</td><td>Santa Cruz biotechnology</td><td>SC-1458</td><td>1/500 dilution</td></tr><tr><td>Primary antibody, ChAT</td><td>Chemicon (Millipore)</td><td>AB144P</td><td>1/25 dilution</td></tr><tr><td>Primary antibody, TH</td><td>ImmunoStar</td><td>22941</td><td>1/1,000 dilution</td></tr><tr><td>Biotinylated secondary antibody, goat</td><td>Vector laboratories</td><td>BA-9500</td><td>1/1,000 dilution</td></tr><tr><td>Biotinylated secondary antibody, mouse</td><td>Vector laboratories</td><td>BA-2000</td><td>1/1,000 dilution</td></tr><tr><td>Vectastain elite ABC kit</td><td>Vector laboratories</td><td>PK6100</td><td>8.8 &amp;micro;L/mL of A and B each</td></tr><tr><td>3,3&#039;-diaminobenzidine (DAB)</td><td>Sigma</td><td>D5637</td><td>0.05% dilution. Product is highly volatile in its powder form and toxic. Do not throw waste in the sink.</td></tr><tr><td>Peroxide (H&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;2&lt;/sub&gt;) 30%</td><td>Fisher scientific</td><td>H-323</td><td>0.005% dilution</td></tr><tr><td>Osmium tetroxide (OsO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Electron microscopic science</td><td>2% 19152&lt;br /&gt;4% 19150&lt;br /&gt;</td><td>Original solution can be either 2 or 4%. Keep attention to which one is used to calculate the final 1% dilution. Product is very sensitive to light. Osmium is highly toxic. Use only under a venting hood with appropriate protection.</td></tr><tr><td>Durcupan water-repellent epoxy resin</td><td>Sigma</td><td>A: M epoxy resin (44611)&lt;br /&gt;B: hardener 964 (44612)&lt;br /&gt;C: accelerator 960 (DY 060) (44613)&lt;br /&gt;D: plasticizer (44614)&amp;nbsp;</td><td>Polymerize 48 h at 58 &amp;deg;C before throwing in waste.</td></tr><tr><td>Alumium cups</td><td>Electron microscopic science</td><td>70048-01</td><td></td></tr><tr><td>Ethanol</td><td>commercial alcohols</td><td>1019C</td><td>Dilute in distilled water with appropriate concentration</td></tr><tr><td>Propylene oxide</td><td>Electron microscopic science</td><td>20401</td><td>Organic solvent. Highly volatile and toxic. Use under a venting hood.</td></tr><tr><td>Non-coated medium glass slides</td><td>brain research laboratories</td><td>3875-FR</td><td>Grease surface with mineral oil</td></tr><tr><td>Plastic film (Aclar embedding film)</td><td>Electron microscopic science</td><td>50425-25</td><td>Grease surface with mineral oil</td></tr><tr><td>Ultramicrotome</td><td>Leica UC7</td><td>EM UC7</td><td></td></tr><tr><td>Diamond trimming tool (ultratrim)</td><td>Diatome&amp;nbsp;</td><td>UT 1081</td><td>Can use glass knife alternatively</td></tr><tr><td>Ultra 45&amp;deg; Diatome Diamond knife</td><td>Diatome&amp;nbsp;</td><td>MC13437</td><td>equipped with a boat</td></tr><tr><td>Xylenes</td><td>Fisher scientific</td><td>X5SK-4</td><td></td></tr><tr><td>150-mesh copper grids</td><td>Electron microscopic science</td><td>G150-cu</td><td></td></tr><tr><td>grid-box</td><td>Electron microscopic science</td><td>71138</td><td>Can store up to 100 grids</td></tr><tr><td>Sodium citrate</td><td>Anachemia</td><td>81983</td><td></td></tr><tr><td>Lead nitrate</td><td>Sigma</td><td>L-6258</td><td>Make a stock solution of lead citrate made of 1.33&amp;nbsp;g of lead nitrate and 1.76&amp;nbsp;g of sodium citrate diluted in 42&amp;nbsp;mL of preboiled and cooled distilled water to which 8&amp;nbsp;mL of 1&amp;nbsp;N NaOH are added after the conversion from lead nitrate to lead citrate is complete. pH should be approximately 12. Store solution in a hermetic plastic bottle and protect from light.</td></tr><tr><td>Syringe</td><td>terumo&amp;nbsp;</td><td>SS-05L</td><td>5 mL</td></tr><tr><td>Syringe filter</td><td>Corning</td><td>431222</td><td>0.2 &amp;micro;m</td></tr><tr><td>Absorbing paper (bibulous paper)</td><td>Electron microscopic science</td><td>70086-1</td><td></td></tr><tr><td>Parafilm</td><td>Laboratory film</td><td>PM-999</td><td></td></tr><tr><td>Mineral oil</td><td>Sigma</td><td>M5904</td><td></td></tr></tbody></table>

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.

In this section, we present representative results that were obtained following the observation, at the transmission EM&#160;level, of immunostained primate brain tissue chemically fixed with a mixture of 3% acrolein and 4% PFA. We achieved good preservation of the ultrastructure, as indicated by the relatively intact myelin sheath and the neat visualization of double membranes (Figure 2A). Synaptic contacts, along with neuronal elements from the microenvironment, can eas...

Watch this Scientific Journal Video about Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy at JoVE.com

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

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-non-human-primate-brain-tissue-for-pre-embedding

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

14.0K Views.  Université Laval. 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.

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

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.

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 ...

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; Gan et al., 2000). Here we provide a detailed description of each step required together with exemplary images of good and bad outcomes that will help when setting up the technique. In our experience, a few steps proved critical for the successful application of DiOLISTICS. These considerations include the quality of the DiI-coated bullets, the extent of fixative exposure, and the concentration of detergent used in the incubation solutions. Tips and solutions for common problems are provided. This is a versatile labeling technique that can be applied to multiple animal species at a wide range of ages. Unlike other fluorescent labeling techniques that are limited to preparations from young animals or restricted to mice because they rely on the expression of a fluorescent transgene, DiOLISTIC labeling can be applied to animals of all ages, species and genotypes and it can be used in combination with immunostaining to identify a specific subpopulation of cells. Here we demonstrate the use of DiOLISTICS to label neurons in brain slices from adult mice and adult non-human primates with the purpose of quantifying dendrite branching and dendritic spine morphology.

We demonstrate the use of the gene gun to introduce fluorescent dyes, such as DiI, into neurons in brain slices from rodents and non-human primates of different ages. In this particular case, we use adult mice (3-6 months old) and adult cynomologus monkeys (9-15 years old). This technique, originally described by the laboratory of Dr. Lichtman (Gan et al., 2000), is well suited for the study of dendritic branching and dendritic spine morphology and can be combined with traditional immunostaining, if detergents are kept at a low concentration.

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; ...

Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy

Major advances in high-throughput, high-resolution, 3D microscopy techniques have enabled the acquisition of large volumes of neuroanatomical data at submicrometer resolution. One of the first such instruments producing whole-brain-scale data is the Knife-Edge Scanning Microscope (KESM)7, 5, 9, developed and hosted in the authors' lab. KESM has been used to section and image whole mouse brains at submicrometer resolution, revealing the intricate details of the neuronal networks (Golgi)1, 4, 8, vascular networks (India ink)1, 4, and cell body distribution (Nissl)3. The use of KESM is not restricted to the mouse nor the brain. We have successfully imaged the octopus brain6, mouse lung, and rat brain. We are currently working on whole zebra fish embryos. Data like these can greatly contribute to connectomics research10; to microcirculation and hemodynamic research; and to stereology research by providing an exact ground-truth. 
In this article, we will describe the pipeline, including specimen preparation (fixing, staining, and embedding), KESM configuration and setup, sectioning and imaging with the KESM, image processing, data preparation, and data visualization and analysis. The emphasis will be on specimen preparation and visualization/analysis of obtained KESM data. We expect the detailed protocol presented in this article to help broaden the access to KESM and increase its utilization.

The full process from brain specimen preparation to serial sectioning imaging using the Knife-Edge Scanning Microscope, to data visualization and analysis is described. This technique is currently used to acquire mouse brain data, but it is applicable to other organs, other species.

Major advances in high-throughput, high-resolution, 3D microscopy techniques have enabled the acquisition of large volumes of neuroanatomical data at ...

Bioengineering

Brain Banking: Making the Most of your Research Specimens

Unbiased stereology is a method for accurately and efficiently estimating the total neuron number (or other cell type) in a given area of interest1. To achieve this goal 6-10 systematic sections should be probed covering the entire structure. Typically this involves processing 1/5 sections which leaves a significant amount of material unprocessed. In order to maximize the material, we propose an inexpensive method for preserving fixed tissue as part of a long-term storage research plan. As tissue is sliced and processed for the desired stain or antibody, alternate sections should be systematically placed in antigen preserve at -20&deg;C for future processing. Using 24-well plates, sections can be placed in order for future retrieval. Using this method, tissue can be stored and processed for immunohistochemistry over the course of years.

Brain banking and systematic sampling of biological material provides the basis for unbiased stereology and maximizes the potential data obtained from each specimen.

Unbiased stereology is a method for accurately and efficiently estimating the total neuron number (or other cell type) in a given area of interest1.  To ...

Biology

Batch Immunostaining for Large-Scale Protein Detection in the Whole Monkey Brain

Immunohistochemistry (IHC) is one of the most widely used laboratory techniques for the detection of target proteins in situ. Questions concerning the expression pattern of a target protein across the entire brain are relatively easy to answer when using IHC in small brains, such as those of rodents. However, answering the same questions in large and convoluted brains, such as those of primates presents a number of challenges. Here we present a systematic approach for immunodetection of target proteins in an adult monkey brain. This approach relies on the tissue embedding and sectioning methodology of NeuroScience Associates (NSA) as well as tools developed specifically for batch-staining of free-floating sections. It results in uniform staining of a set of sections which, at a particular interval, represents the entire brain. The resulting stained sections can be subjected to a wide variety of analytical procedures in order to measure protein levels, the population of neurons expressing a certain protein.

Large-scale immunodetection of target proteins across the entire primate brain is possible by employing novel tissue embedding and sectioning methods combined with the use of creative apparatus for batch staining of multiple free-floating sections at a given time.

Immunohistochemistry (IHC) is one of the most widely used laboratory techniques for the detection of target proteins in situ. Questions concerning the ...

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and simple protocol to detect protein expression during craniofacial morphogenesis/pathogenesis using mouse craniofacial tissues as examples. The protocol consists of preparation and cryosectioning of tissues, indirect immunofluorescence, image acquisition, and quantification. In addition, a method for preparation and cryosectioning of undecalcified hard tissues for immunostaining is described, using craniofacial tissues and long bones as examples. Those methods are key to determine the protein expression and morphological/anatomical changes in various tissues during craniofacial morphogenesis/pathogenesis. They are also applicable to other tissues with appropriate modifications. Knowledge of the histology and high quality of sections are critical to draw scientific conclusions from experimental outcomes. Potential limitations of this methodology include but are not limited to specificity of antibodies and difficulties of quantification, which are also discussed here.

Here, we present a detailed protocol to detect and quantify protein levels during craniofacial morphogenesis/pathogenesis by immunostaining using mouse craniofacial tissues as examples. In addition, we describe a method for preparation and cryosectioning of undecalcified hard tissues from young mice for immunostaining.

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and ...

Developmental Biology

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brain tissue, using markers for microglia and neuronal elements as an example. Specifically, this paper is a step-by-step protocol for fluorescent visualization of microglia and neurons via immunohistochemistry for Iba1 and Pan-neuronal, respectively. Fluorescence double-labeling is particularly useful for the localization of multiple proteins within the same sample, providing the opportunity to accurately observe interactions between cell types, receptors, ligands, and/or the extracellular matrix in relation to one another as well as protein co-localization within a single cell. Unlike other visualization techniques, fluorescence immunohistochemistry staining intensity may decrease in the weeks to months following staining, unless appropriate precautions are taken. Despite this limitation, in many applications fluorescence double-labeling is preferred over alternatives such as 3,3&#39;-diaminobenzidine tetrahydrochloride (DAB) or alkaline phosphatase (AP), as fluorescence is more time efficient and allows for more precise differentiation between two or more markers. The discussion includes troubleshooting tips and advice to promote success.

This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brains, using markers for microglia and neuronal elements as an example.

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level ...

Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the hematogenous route, which involves infection of the endothelial cells and pericytes of the brain. The second is the intracerebroventricular (ICV) route. Once within the central nervous system (CNS), viruses may spread to the subarachnoid space, meninges, and choroid plexus via the cerebrospinal fluid. In experimental models, the earliest stages of CNS viral distribution are not well characterized, and it is unclear whether only certain cells are initially infected. Here, we have analyzed the distribution of cytomegalovirus (CMV) particles during the acute phase of infection, termed primary viremia, following ICV or intravascular (IV) injection into the neonatal mouse brain. In the ICV injection model, 5 &#181;l of murine CMV (MCMV) or fluorescent microbeads were injected into the lateral ventricle at the midpoint between the ear and eye using a 10-&#181;l syringe with a 27 G needle. In the IV injection model, a 1-ml syringe with a 35 G needle was used. A transilluminator was used to visualize the superficial temporal (facial) vein of the neonatal mouse. We infused 50 &#181;l of MCMV or fluorescent microbeads into the superficial temporal vein. Brains were harvested at different time points post-injection. MCMV genomes were detected using the in situ hybridization method. Fluorescent microbeads or green fluorescent protein expressing recombinant MCMV particles were observed by fluorescent microscopy. These techniques can be applied to many other pathogens to investigate the pathogenesis of encephalitis.

Here, we describe a simple method of intracerebroventricular and intravascular injection of viral particles or fluorescent microbeads into the neonatal mouse brain. The localization pattern of the virus and nanoparticles could be detected by microscopic evaluation or by in situ hybridization.

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the ...

Short-Term Free-Floating Slice Cultures from the Adult Human Brain

Organotypic, or slice cultures, have been widely employed to model aspects of the central nervous system functioning in vitro. Despite the potential of slice cultures in neuroscience, studies using adult nervous tissue to prepare such cultures are still scarce, particularly those from human subjects. The use of adult human tissue to prepare slice cultures is particularly attractive to enhance the understanding of human neuropathologies, as they hold unique properties typical of the mature human brain lacking in slices produced from rodent (usually neonatal) nervous tissue. This protocol describes how to use brain tissue collected from living human donors submitted to resective brain surgery to prepare short-term, free-floating slice cultures. Procedures to maintain and perform biochemical and cell biology assays using these cultures are also presented. Representative results demonstrate that the typical human cortical lamination is preserved in slices after 4 days in vitro (DIV4), with expected presence of the main neural cell types. Moreover, slices at DIV4 undergo robust cell death when challenged with a toxic stimulus (H2O2), indicating the potential of this model to serve as a platform in cell death assays. This method, a simpler and cost-effective alternative to the widely used protocol using membrane inserts, is mainly recommended for running short-term assays aimed to unravel mechanisms of neurodegeneration behind age-associated brain diseases. Finally, although the protocol is devoted to using cortical tissue collected from patients submitted to surgical treatment of pharmacoresistant temporal lobe epilepsy, it is argued that tissue collected from other brain regions/conditions should also be considered as sources to produce similar free-floating slice cultures.

A protocol to prepare free-floating slice cultures from adult human brain is presented. The protocol is a variation of the widely used slice culture method using membrane inserts. It is simple, cost-effective, and recommended for running short-term assays aimed to unravel mechanisms of neurodegeneration behind age-associated brain diseases.

Organotypic, or slice cultures, have been widely employed to model aspects of the central nervous system functioning in vitro. Despite the potential of ...

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within cells. Neurochemical phenotyping of functionally identified neurons usually requires concurrent labelling with multiple antibodies (targeting protein) using immunohistochemistry (IHC) and optimization of in situ hybridization (targeting RNA), in tandem. A &#34;neurochemical signature&#34; to characterize particular neurons may be achieved however complicating factors include the need to verify FISH and IHC targets before combining the methods, and the limited number of RNAs and proteins that may be targeted simultaneously within the same tissue section.
Here we describe a protocol, using both fresh frozen and fixed mouse brain preparations, which detects multiple mRNAs and proteins in the same brain section using RNAscope FISH followed by fluorescence immunostaining, respectively. We use the combined method to describe the expression pattern of low abundance mRNAs (e.g., galanin receptor 1) and high abundance mRNAs (e.g., glycine transporter 2), in immunohistochemically identified brainstem nuclei.
Key considerations for protein labelling downstream of the FISH assay extend beyond tissue preparation and optimization of FISH probe labelling. For example, we found that antibody binding and labelling specificity can be detrimentally affected by the protease step within the FISH probe assay. Proteases catalyze hydrolytic cleavage of peptide bonds, facilitating FISH probe entry into cells, however they may also digest the protein targeted by the subsequent IHC assay, producing off target binding. The subcellular location of the targeted protein is another factor contributing to IHC success following FISH probe assay. We observed IHC specificity to be retained when the targeted protein is membrane bound, whereas IHC targeting cytoplasmic protein required extensive troubleshooting. Finally, we found handling of slide-mounted fixed frozen tissue more challenging than fresh frozen tissue, however IHC quality was overall better with fixed frozen tissue, when combined with RNAscope.

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

This protocol describes a method for combining fluorescence in situ hybridization (FISH) and fluorescence immunohistochemistry (IHC) in both fresh frozen and fixed mouse brain sections, with the goal of achieving multilabel FISH and fluorescence IHC signal. IHC targeted cytoplasmic and membrane attached proteins.

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within ...

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

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Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy

Brain Banking: Making the Most of your Research Specimens

Batch Immunostaining for Large-Scale Protein Detection in the Whole Monkey Brain

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain

Short-Term Free-Floating Slice Cultures from the Adult Human Brain

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections