This work was supported by a Discovery Grant to CDCB from the Natural Sciences and Engineering Research Council of Canada (NSERC), a John R. Evans Leaders Fund research infrastructure grant to CDCB from Canada Foundation for Innovation (CFI project number 30381), and by a Discovery Grant to NJM from NSERC. ELL was supported by an Ontario Graduate Scholarship and by an OVC Scholarship from the Ontario Veterinary College at the University of Guelph. CDS was supported by an Undergraduate Student Research Assistantship from NSERC. ALM was supported by an Alexander Graham Bell Scholarship from NSERC and by an OVC Scholarship from the Ontario Veterinary College at the University of Guelph.

Adult female CD1-strain mice were used in this study. Similar staining can be accomplished using both sexes at various ages. Experimental animals were cared for according to the principles and guidelines of the Canadian Council on Animal Care, and the experimental protocol was approved by the University of Guelph Animal Care Committee.
1. Golgi-Cox Staining
<ol>
	<li>Golgi-Cox Impregnation of Brains
	<ol>
		<li>Make the Golgi-Cox solution of 1% (w/v) potassium dichromate, 0.8% (w/v) potassium chromate and 1% (w/v) mercuric chloride by dissolving the potassium dichromate and potassium chromate into high-quality water separately. Mix the solutions, add the mercuric chloride and filter the final solution using grade 1 filter paper. Store the solution in the dark for up to one month.</li>
		<li>Anesthetize the mouse using 5% isoflurane.</li>
		<li>Euthanize the mouse by decapitation and quickly remove the brain.</li>
		<li>Place the brain into a 20 mL glass scintillation vial containing 17 mL of Golgi-Cox solution and incubate in the dark at RT for 25 d.</li>
		<li>Cryoprotect the brain by placing it into a 50 mL conical tube containing 40 mL of sucrose cryoprotectant (30% (w/v) sucrose in 0.1 M phosphate buffer, pH 7.4) in the dark at 4 &#176;C for 24 h. 
		NOTE: The following optional three steps may be employed as an alternative to the main protocol, in order to freeze the brain at this stage for long-term storage.</li>
		<li>Freeze the whole brain by immersing it into 200 mL isopentane that has been precooled on dry ice.</li>
		<li>Place the frozen brain into a 50 mL conical tube and store in the dark at -80 &#176;C.</li>
		<li>When ready to proceed, thaw the brain by placing it into a 50 mL conical tube containing 40 mL of sucrose cryoprotectant in the dark at 4 &#176;C for 24 h.</li>
	</ol>
	</li>
	<li>Brain Sectioning
	<ol>
		<li>Remove the brain from the sucrose and block it for sectioning by cutting off the cerebellum using a razor blade and leaving a flat edge at the remaining caudal end of the brain.</li>
		<li>Heat agar (3% (w/v) in water) until it is melted and let cool until it is slightly above its melting point.</li>
		<li>Place the brain in a small disposable weigh dish with its caudal end face-down and add a sufficient amount of melted agar to cover the brain.</li>
		<li>Once the agar has solidified, trim excess agar leaving approximately 2 - 4 mm surrounding the brain and glue the brain to the stage of a vibratome with its caudal end face-down using a small amount of ethyl cyanoacrylate glue.</li>
		<li>&#160;Fill the stage area of the vibratome with a sufficient amount of sucrose cryoprotectant to cover the brain, and section the brain at a slice thickness of 400 - 500 &#181;m (depending on the brain region to be examined) using a vibration frequency of 86 Hz and a blade advancement speed of 0.125 mm/s.</li>
		<li>Using a small paint brush, place brain sections into a well of a 6-well tissue culture plate containing commercially-available mesh-bottom inserts and pre-filled with 10 mL of 6% (w/v) sucrose in 0.1 M phosphate buffer, pH 7.4.</li>
		<li>Incubate sections in the dark at 4 &#176;C O/N.</li>
	</ol>
	</li>
	<li>Developing Brain Sections
	<ol>
		<li>Using the mesh-bottom inserts, transfer brain sections into a new well containing 5 mL of 2% (w/v) paraformaldehyde (PBD) in 0.1 M phosphate buffer, pH 7.4. Incubate on a rocker moving at slow speed in the dark at RT for 15 min.</li>
		<li>Wash sections twice by transferring them to new wells containing 5 mL of water. Wash on a rocker moving at a moderate speed in the dark at RT for 5 min.</li>
		<li>Transfer sections into a new well containing 5 mL of 2.7% (v/v) ammonium hydroxide. Incubate on a rocker moving at a slow speed in the dark at RT for 15 min.</li>
		<li>Wash sections twice by transferring them to new wells containing 5 mL of water. Wash on a rocker moving at a moderate speed in the dark at RT for 5 min.</li>
		<li>Transfer sections into a new well containing 5 mL of Fixative A (see Materials Table) that has been diluted in water 10x from its original purchased concentration. Incubate on a rocker moving at a slow speed in the dark at RT for 25 min.</li>
		<li>Wash sections twice by transferring them to new wells containing 5 mL of water. Wash on a rocker moving at a moderate speed in the dark at RT for 5 min.</li>
	</ol>
	</li>
	<li>Mounting Brain Sections
	<ol>
		<li>Using a small paint brush, mount sections onto microscope slides. Remove excess water and agar using tweezers and a small tissue. Ensure that all agar is removed before proceeding with dehydration.</li>
		<li>Allow sections to air dry at RT for approximately 45 min (400 &#956;m sections) or 90 min (500 &#956;m sections). 
		NOTE: The timing and proper level of dryness is critical, and may need to be determined in each laboratory depending on ambient temperature and humidity level. Too short a drying time leads to sections falling off of slides during subsequent dehydration steps, and too long a drying time leads to sections cracking. Sections will still appear to be shiny at the appropriate level of dryness.</li>
		<li>Stain sections with cresyl violet by placing slides into Coplin staining jars as indicated. 
		NOTE: This optional step may be employed for sections that had never been frozen, in order to stain neuronal nuclei with cresyl violet. We have found that clearing and rehydrating sections before incubation in cresyl violet leads to even staining and low background across sections.
		<ol>
			<li>Place in clearing agent for 5 min. Repeat once.</li>
			<li>Place in 100% ethanol for 5 min. Repeat once.</li>
			<li>Place in 95% ethanol in water, then 75% ethanol in water, and then 50% ethanol in water for 2 min each.</li>
			<li>Place in water for 5 min.</li>
			<li>Place in 0.5% (w/v) cresyl violet in water for 7 min.</li>
			<li>Place in water for 2 min. Repeat once.</li>
		</ol>
		</li>
		<li>Dehydrate sections by placing slides into Coplin staining jars as indicated.
		<ol>
			<li>Place in 50% ethanol in water, then 75% ethanol in water, and then 95% ethanol in water for 2 min each.</li>
			<li>Place in 100% ethanol for 5 min. Repeat once.</li>
			<li>Place in clearing agent for 5 min. Repeat once. 
			NOTE: These dehydration and clearing times are sufficient to process mouse brains as described in this manuscript. However, we have observed for other species including rat and cowbird, that the final clearing step may need to be extended up to 15 min total.</li>
		</ol>
		</li>
		<li>Coverslip sections using an anhydrous mounting medium.</li>
		<li>Allow slides to dry horizontally in the dark at RT for at least 5 d.</li>
	</ol>
	</li>
</ol>
2. Imaging Stained Neurons within Thick Brain Sections
<ol>
	<li>Capturing Image Stacks
	<ol>
		<li>Turn on the microscope light bulb, camera, and stage controller.</li>
		<li>Open the microscope software (e.g., Neurolucida).</li>
		<li>Place a slide on the microscope stage.</li>
		<li>Capture a 2D wide-view image of the brain section using a low magnification objective such as 1.25X 0.4 N.A. PlanAPO or 4X 0.16 N.A.
		<ol>
			<li>Focus image and adjust camera settings including the exposure time and white balance.</li>
			<li>Create a reference point by left-clicking anywhere on the section</li>
			<li>Capture the image by selecting &#34;acquire single image&#34; within the image acquisition window.</li>
		</ol>
		</li>
		<li>Capture mid-resolution image stacks of the area containing the neurons(s) of interest, using a 10X 0.3 N.A. UPlan FL N objective.
		<ol>
			<li>Focus the image and adjust camera settings including the exposure and white balance.</li>
			<li>Set the upper and lower boundaries for the image stack by focusing to the top of the section and selecting &#34;set&#34; next to &#34;top of stack&#34; within the image acquisition window, and then focusing to the bottom of the section and selecting &#34;set&#34; next to &#34;bottom of stack&#34; within the image acquisition window.</li>
			<li>Set the step distance to 5 &#956;m by entering &#34;5 &#956;m&#34; next to &#34;distance between images&#34; within the image acquisition window.</li>
			<li>Capture the image stack by selecting &#34;acquire image stack&#34; within the image acquisition window.</li>
			<li>Repeat the above steps to capture the entire area of interest, making sure that all image stacks overlap by at least 10% in the X and Y axes.</li>
		</ol>
		</li>
		<li>Capture high-resolution image stacks of the area containing the neuron of interest, using a 30X 1.05 N.A. silicone oil-immersion objective.
		<ol>
			<li>Apply 3 - 4 drops of silicone immersion oil to the slide and place the objective over the slide, ensuring to make contact between the objective and oil.</li>
			<li>Focus the image and adjust camera settings including the exposure and white balance.</li>
			<li>Set the upper and lower boundaries for the image stack by focusing to the top of the section and selecting &#34;set&#34; next to &#34;top of stack&#34; within the image acquisition window, and then focusing to the bottom of the section and selecting &#34;set&#34; next to &#34;bottom of stack&#34; within the image acquisition window.</li>
			<li>Set the step distance to 1 &#956;m by entering &#34;1 &#956;m&#34; next to &#34;distance between images&#34; within the image acquisition window.</li>
			<li>Capture the image stack by selecting &#34;acquire image stack&#34; within the image acquisition window.</li>
			<li>Repeat the above steps to capture the entire area of interest, making sure that all image stacks overlap by at least 10% in the X and Y axes.</li>
		</ol>
		</li>
		<li>Save the data file and save all image files in TIFF format for external processing.</li>
	</ol>
	</li>
	<li>Creating Z-projection Images and Image Montages
	<ol>
		<li>Create Z-projection images in ImageJ
		<ol>
			<li>Open ImageJ software that has the Bio-Formats plugin installed.</li>
			<li>Select &#34;Plugins&#34; -&#62; &#34;Bio-Formats&#34; -&#62; &#34;Bio-Formats Importer&#34;.</li>
			<li>Select the image stack file to be opened.</li>
			<li>Once the file is opened, change the format to RGB by selecting &#34;Image&#34; -&#62; &#34;Type&#34; -&#62; &#34;RGB Color&#34;.</li>
			<li>Create the Z-projection by selecting &#34;Image&#34; -&#62; &#34;Stacks&#34; -&#62; &#34;Z Project&#8230;&#34;.</li>
			<li>Save two-dimensional Z-projection image as a TIFF file.</li>
		</ol>
		</li>
		<li>Create a two-dimensional image montage of entire area of interest.
		<ol>
			<li>Open the software (e.g., Adobe Photoshop).</li>
			<li>Select &#34;File&#34; -&#62; &#34;Automate&#34; -&#62; &#34;Photomerge&#34;.</li>
			<li>Select &#34;Browse&#34; and then add all images files to be merged.</li>
			<li>Ensure that &#34;Blend Images Together&#34; is selected and then select &#34;OK&#34;.</li>
			<li>Save the resulting montage image of the entire area of interest as a TIFF file.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Golgi, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sulla+struttura+della+sostanza+grigia+del+cervello.">Sulla struttura della sostanza grigia del cervello.</a> Gazzetta Medica Italiana. Lombardia. 33, 244-246 (1873).</li><li>Cox, W. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impr&#228;gnation+des+centralen+Nervensystems+mit+Quecksilbersalzen.">Impr&#228;gnation des centralen Nervensystems mit Quecksilbersalzen.</a> Archiv f. mikrosk. Anat. 37, 16-21 (1891).</li><li>Zaqout, S., Kaindl, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Golgi-Cox+Staining+Step+by+Step.">Golgi-Cox Staining Step by Step.</a> Front Neuroanat. 10, 38 (2016).</li><li>Levine, N. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+thin+tissue+Golgi-Cox+impregnation:+fast,+reliable+methods+for+multi-assay+analyses+in+rodent+and+non-human+primate+brain.">Advances in thin tissue Golgi-Cox impregnation: fast, reliable methods for multi-assay analyses in rodent and non-human primate brain.</a> J Neurosci Methods. 213 (2), 214-227 (2013).</li><li>Ranjan, A., Mallick, B. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modified+method+for+consistent+and+reliable+Golgi-Cox+staining+in+significantly+reduced+time.">A modified method for consistent and reliable Golgi-Cox staining in significantly reduced time.</a> Front Neurosci. 1, 157 (2010).</li><li>Gibb, R., Kolb, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+vibratome+sectioning+of+Golgi-Cox+stained+whole+rat+brain.">A method for vibratome sectioning of Golgi-Cox stained whole rat brain.</a> J Neurosci Methods. 79 (1), 1-4 (1998).</li><li>Friedland, D. R., Los, J. G., Ryugo, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modified+Golgi+staining+protocol+for+use+in+the+human+brain+stem+and+cerebellum.">A modified Golgi staining protocol for use in the human brain stem and cerebellum.</a> J Neurosci Methods. 150 (1), 90-95 (2006).</li><li>Narayanan, S. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Appraisal+of+the+effect+of+brain+impregnation+duration+on+neuronal+staining+and+morphology+in+a+modified+Golgi-Cox+method.">Appraisal of the effect of brain impregnation duration on neuronal staining and morphology in a modified Golgi-Cox method.</a> J Neurosci Methods. 235, 193-207 (2014).</li><li>Das, G., Reuhl, K., Zhou, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Golgi-Cox+method.">The Golgi-Cox method.</a> Methods Mol Biol. 1018, 313-321 (2013).</li><li>Sutherland, R., Gibb, R., Kolb, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+hippocampus+makes+a+significant+contribution+to+experience-dependent+neocortical+plasticity.">The hippocampus makes a significant contribution to experience-dependent neocortical plasticity.</a> Behav Brain Res. 214 (1), 121-124 (2010).</li><li>Hamilton, G. F., Whitcher, L. T., Klintsova, A. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postnatal+binge-like+alcohol+exposure+decreases+dendritic+complexity+while+increasing+the+density+of+mature+spines+in+mPFC+Layer+II/III+pyramidal+neurons.">Postnatal binge-like alcohol exposure decreases dendritic complexity while increasing the density of mature spines in mPFC Layer II/III pyramidal neurons.</a> Synapse. 64 (2), 127-135 (2010).</li><li>Nashed, M. G., Seidlitz, E. P., Frey, B. N., Singh, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Depressive-like+behaviours+and+decreased+dendritic+branching+in+the+medial+prefrontal+cortex+of+mice+with+tumors:+A+novel+validated+model+of+cancer-induced+depression.">Depressive-like behaviours and decreased dendritic branching in the medial prefrontal cortex of mice with tumors: A novel validated model of cancer-induced depression.</a> Behav Brain Res. 294, 25-35 (2015).</li><li>Frauenknecht, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mice+with+experimental+antiphospholipid+syndrome+display+hippocampal+dysfunction+and+a+reduction+of+dendritic+complexity+in+hippocampal+CA1+neurones.">Mice with experimental antiphospholipid syndrome display hippocampal dysfunction and a reduction of dendritic complexity in hippocampal CA1 neurones.</a> Neuropathol Appl Neurobiol. 41 (5), 657-671 (2015).</li><li>Camacho-Abrego, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearrangement+of+the+dendritic+morphology+of+the+neurons+from+prefrontal+cortex+and+hippocampus+after+subthalamic+lesion+in+Sprague-Dawley+rats.">Rearrangement of the dendritic morphology of the neurons from prefrontal cortex and hippocampus after subthalamic lesion in Sprague-Dawley rats.</a> Synapse. 68 (3), 114-126 (2014).</li><li>Redila, V. A., Christie, B. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exercise-induced+changes+in+dendritic+structure+and+complexity+in+the+adult+hippocampal+dentate+gyrus.">Exercise-induced changes in dendritic structure and complexity in the adult hippocampal dentate gyrus.</a> Neuroscience. 137 (4), 1299-1307 (2006).</li><li>Bailey, C. D., Alves, N. C., Nashmi, R., De Biasi, M., Lambe, E. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nicotinic+alpha5+subunits+drive+developmental+changes+in+the+activation+and+morphology+of+prefrontal+cortex+layer+VI+neurons.">Nicotinic alpha5 subunits drive developmental changes in the activation and morphology of prefrontal cortex layer VI neurons.</a> Biol Psychiatry. 71 (2), 120-128 (2012).</li><li>Lytton, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> From computer to brain : foundations of computational neuroscience. , (2002).</li><li>Paxinos, G., Franklin, K. B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The mouse brain in stereotaxic coordinates. , (2001).</li><li>Ha, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modified+Golgi-Cox+method+with+counterstain+for+the+study+of+synapses.">A modified Golgi-Cox method with counterstain for the study of synapses.</a> Anat Rec. 155 (1), 59-64 (1966).</li><li>Swanson, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+locus+coeruleus:+a+cytoarchitectonic,+Golgi+and+immunohistochemical+study+in+the+albino+rat.">The locus coeruleus: a cytoarchitectonic, Golgi and immunohistochemical study in the albino rat.</a> Brain Res. 110 (1), 39-56 (1976).</li><li>Pilati, N., Barker, M., Panteleimonitis, S., Donga, R., Hamann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+method+combining+Golgi+and+Nissl+staining+to+study+neuronal+morphology+and+cytoarchitecture.">A rapid method combining Golgi and Nissl staining to study neuronal morphology and cytoarchitecture.</a> J Histochem Cytochem. 56 (6), 539-550 (2008).</li></ol>

The authors declare that they have no competing financial interests.

We describe here a Golgi-Cox staining protocol along with two useful variants for visualizing neurons within thick brain sections. As shown in the Representative Results, the use of a high-resolution objective that has a long 800 &#956;m working distance allows for the reliable visualization of entire neurons throughout the depth of brain sections cut at 500 &#956;m. This study of relatively thick brain sections increases the probability that stained neurons of any type are fully contained within the slice, which is espe...

The visualization of individual neurons within tissue samples allows for the in situ analysis of neuron morphological characteristics, which has significantly advanced our understanding of the brain and how it may be influenced by endogenous disease or exogenous environmental factors. The Golgi-Cox staining method is a cost-effective, relatively simple means of staining a random sample of neurons within the brain. First developed by Golgi1 and modified by Cox2 in the 1800s, researchers have further refined this technique over the years to produce clear, well-stained neurons that can be used to visualize and quantify both dendritic tree morphology and spine density3,4,5,6,7,8,9.
A major technical consideration for the visualization of stained neurons within brain sections is the maximum slice thickness, which is limited by the working distance of available high-magnification/high-resolution microscope objectives. Common oil-immersion objectives in the 60 - 100X range provide excellent resolution, but are limited by their working distances that are typically no greater than 200 &#956;m. Brain sections cut at the 200 &#956;m range may be adequate to visualize certain neuron types that can be contained within this slice thickness, for example pyramidal neurons in shallow layers of the cerebral cortex10,11,12, pyramidal neurons in the CA1 region of the hippocampus13,14, and granule cells in the dentate gyrus of the hippocampus15. Neurons with relatively longer dendritic&#160;trees, such as pyramidal neurons within deep layers of the cerebral cortex that for mouse can extend more than 800 &#956;m from the cell body16, provide a greater challenge because brains would need to be sectioned at a perfect angle to contain the entire dendrite tree within 200 &#956;m slices. This may not even be feasible if a dendrite or any of its branches extend in the rostral or caudal direction. While it is possible to address this limitation by tracing a neuron across multiple adjacent brain sections, this approach introduces a significant technical challenge in aligning the sections accurately for tracing17. A more practical approach would be to visualize entire neurons contained within brain sections that are cut at a greater thickness.
We report here a technique to stain neurons within 400 - 500 &#956;m thick brain sections of mice using the Golgi-Cox method, and to visualize their morphology using a high-resolution silicon oil-immersion objective that has an 800 &#956;m working distance. The Golgi-Cox impregnation and processing protocol that we describe is modified from one of the most-cited modern protocols in the literature6. Our approach with thick brain sections provides the advantage of increasing the probability of identifying neurons of any type that are fully contained within the section. In addition to the main protocol, we also present two variations that provide unique advantages: (1) Golgi-Cox staining with the cresyl violet counterstain on the surface of mounted sections, in order to define boundaries of brain regions and to identify layers of the cerebral cortex, and (2) Golgi-Cox staining with an intermediate freezing step for the long-term storage of impregnated whole brains prior to sectioning and final processing.

<table><tbody><tr><td>potassium dichromate</td><td>Fisher Scientific</td><td>P188-100</td><td>Hazardous</td></tr><tr><td>potassium chromate</td><td>Fisher Scientific</td><td>P220-100</td><td>Hazardous</td></tr><tr><td>mercuric chloride</td><td>Fisher Scientific</td><td>S25423</td><td>Hazardous</td></tr><tr><td>Whatman grade 1 filter paper</td><td>Fisher Scientific</td><td>1001-185</td><td></td></tr><tr><td>isoflurane</td><td>Pharmaceutical Partners of Canada</td><td>CP0406V2</td><td></td></tr><tr><td>20 mL scintillation vial</td><td>Fisher Scientific</td><td>03-337-4</td><td></td></tr><tr><td>sucrose</td><td>Bioshop Canada</td><td>SUC700.1</td><td></td></tr><tr><td>sodium phosphate monobasic</td><td>Sigma Aldrich</td><td>S5011-500G</td><td></td></tr><tr><td>sodium phosphate dibasic</td><td>Sigma Aldrich</td><td>S9390-500G</td><td></td></tr><tr><td>50 mL conical tube</td><td>Fisher Scientific</td><td>12-565-271</td><td></td></tr><tr><td>isopentane</td><td>Fisher Scientific</td><td>AC126470010</td><td>Also known as 2-methylbutane; hazardous</td></tr><tr><td>agar</td><td>Sigma Aldrich</td><td>A1296-100G</td><td></td></tr><tr><td>small weigh dish</td><td>Fisher Scientific</td><td>02-202-100</td><td></td></tr><tr><td>vibratome</td><td>Leica</td><td>VT1000 S</td><td></td></tr><tr><td>6-well tissue culture plates</td><td>Fisher Scientific</td><td>08-772-1b</td><td></td></tr><tr><td>mesh bottom tissue culture inserts</td><td>Fisher Scientific</td><td>07-200-214</td><td></td></tr><tr><td>paraformadelhyde (PFA), 16%</td><td>Electron Microscope Sciences</td><td>15710-S</td><td>Hazardous</td></tr><tr><td>ammonium hydroxide</td><td>Fisher Scientific</td><td>A669S-500</td><td>Hazardous</td></tr><tr><td>Kodak Fixative A</td><td>Sigma Aldrich</td><td>P7542</td><td></td></tr><tr><td>superfrost plus slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>CitroSolv clearing agent</td><td>Fisher Scientific</td><td>22-143-975</td><td></td></tr><tr><td>anhydrous ethyl alcohol</td><td>Commercial Alcohols</td><td>N/A</td><td></td></tr><tr><td>cresyl violet</td><td>Sigma Aldrich</td><td>C1791</td><td></td></tr><tr><td>permount</td><td>Fisher Scientific</td><td>SP15-100</td><td></td></tr><tr><td>upright microscope</td><td>Olympus</td><td>BX53 model</td><td></td></tr><tr><td>colour camera, 12 bit</td><td>MBF Biosciences</td><td>DV-47d</td><td>QImaging part 01-MBF-2000R-F-CLR-12</td></tr><tr><td>3D motorized microscope stage, controller and enoders</td><td>MBF Biosciences</td><td>N/A</td><td>Supplied and integrated with microscope by MBF Biosciences</td></tr><tr><td>4X microscope objective</td><td>Olympus</td><td>4x 0.16 N.A. UplanSApo</td><td></td></tr><tr><td>10X microscope objective</td><td>Olympus</td><td>10x 0.3 N.A. UPlan FL N&amp;nbsp;</td><td></td></tr><tr><td>30X microscope objective</td><td>Olympus</td><td>30x 1.05 N.A. UPlanSApo&amp;nbsp;</td><td></td></tr><tr><td>60X microscope objective</td><td>Olympus</td><td>60x 1.42 N.A. PlanAPO N</td><td></td></tr><tr><td>silicone immersion oil</td><td>Olympus</td><td>Z-81114</td><td></td></tr><tr><td>Neurolucida software</td><td>MBF Biosciences</td><td>Version 10</td><td></td></tr><tr><td>ImageJ software</td><td>U. S. National Institutes of Health</td><td>Current version</td><td>With the OME Bio-Formats plugin installed</td></tr><tr><td>Photoshop software</td><td>Adobe</td><td>version CS6</td><td></td></tr></tbody></table>

imaging neurons within thick brain sections using the golgi-cox method

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within histological brain samples. While it is preferable from a practical perspective to prepare brain sections at the greatest thickness possible, in order to increase the probability of identifying stained neurons that are fully contained within single sections, this approach is limited from a technical perspective by the working distance of high-magnification microscope objectives. We report here a protocol to stain neurons using the Golgi-Cox method in mouse brain sections that are cut at 500 &#956;m thickness, and to visualize neurons throughout the depth of these sections using an upright microscope fitted with a high-resolution 30X 1.05 N.A. silicone oil-immersion objective that has an 800 &#956;m working distance. We also report two useful variants of this protocol that may be employed to counterstain the surface of mounted brain sections with the cresyl violet Nissl stain, or to freeze whole brains for long-term storage prior to sectioning and final processing. The main protocol and its two variants produce stained thick brain sections, throughout which full neuron dendritic&#160;trees and dendrite spines may be reliably visualized and quantified.

This Golgi-Cox staining protocol and its two described optional variants may be employed to visualize individual neurons within 400 - 500 &#956;m thick brain sections. Representative image montages of two-dimensional Z-projections captured using a 10X&#160;objective and 5 &#956;m steps in the Z axis are shown in Figure 1: A1 - C1&#160;for a large area of coronal brain sections that includes the anterior cingulate cortex area 1 and the secondary motor cortex<sup class="xre...

Watch this Scientific Journal Video about Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method at JoVE.com

Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method

We present a protocol for using the Golgi-Cox staining method in thick brain sections, in order to visualize neurons with long dendritic&#160;trees contained within single tissue samples. Two variants of this protocol are also presented that involve cresyl violet counterstaining, and the freezing of unprocessed brains for long-term storage.

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within ...

imaging-neurons-within-thick-brain-sections-using-the-golgi-cox-method

Research

JoVE Journal

Neuroscience

18.3K Views.  University of Guelph.  We presenteren een protocol voor het gebruik van Cox Golgi-kleuring werkwijze dikke hersenpreparaten, teneinde neuronen met lange dendritische bomen opgenomen in één weefselmonsters zichtbaar. Twee varianten van dit protocol worden ook getoond dat cresyl violet tegenkleuring, en het bevriezen van onbewerkte hersenen voor langdurige opslag omvatten. 

Video: Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method

A Rapid Approach to High-Resolution Fluorescence Imaging in Semi-Thick Brain Slices

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are characteristic to neurons in both normal and diseased brain. Towards this, a great deal of effort has been placed on building high-resolution neuroanatomical maps1-3. With the expansion of molecular genetics and advances in light microscopy has come the ability to query not only neuronal morphologies, but also the molecular and cellular makeup of individual neurons and their associated networks4. Major advances in the ability to mark and manipulate neurons through transgenic and gene targeting technologies in the rodent now allow investigators to 'program' neuronal subsets at will5-6. Arguably, one of the most influential contributions to contemporary neuroscience has been the discovery and cloning of genes encoding fluorescent proteins (FPs) in marine invertebrates7-8, alongside their subsequent engineering to yield an ever-expanding toolbox of vital reporters9. Exploiting cell type-specific promoter activity to drive targeted FP expression in discrete neuronal populations now affords neuroanatomical investigation with genetic precision.
Engineering FP expression in neurons has vastly improved our understanding of brain structure and function. However, imaging individual neurons and their associated networks in deep brain tissues, or in three dimensions, has remained a challenge. Due to high lipid content, nervous tissue is rather opaque and exhibits auto fluorescence. These inherent biophysical properties make it difficult to visualize and image fluorescently labelled neurons at high resolution using standard epifluorescent or confocal microscopy beyond depths of tens of microns. To circumvent this challenge investigators often employ serial thin-section imaging and reconstruction methods10, or 2-photon laser scanning microscopy11. Current drawbacks to these approaches are the associated labor-intensive tissue preparation, or cost-prohibitive instrumentation respectively.
Here, we present a relatively rapid and simple method to visualize fluorescently labelled cells in fixed semi-thick mouse brain slices by optical clearing and imaging. In the attached protocol we describe the methods of: 1) fixing brain tissue in situ via intracardial perfusion, 2) dissection and removal of whole brain, 3) stationary brain embedding in agarose, 4) precision semi-thick slice preparation using new vibratome instrumentation, 5) clearing brain tissue through a glycerol gradient, and 6) mounting on glass slides for light microscopy and z-stack reconstruction (Figure 1).
For preparing brain slices we implemented a relatively new piece of instrumentation called the 'Compresstome' VF-200 (http://www.precisionary.com/products_vf200.html). This instrument is a semi-automated microtome equipped with a motorized advance and blade vibration system with features similar in function to other vibratomes. Unlike other vibratomes, the tissue to be sliced is mounted in an agarose plug within a stainless steel cylinder. The tissue is extruded at desired thicknesses from the cylinder, and cut by the forward advancing vibrating blade. The agarose plug/cylinder system allows for reproducible tissue mounting, alignment, and precision cutting. In our hands, the 'Compresstome' yields high quality tissue slices for electrophysiology, immunohistochemistry, and direct fixed-tissue mounting and imaging. Combined with optical clearing, here we demonstrate the preparation of semi-thick fixed brain slices for high-resolution fluorescent imaging.

Here we describe a rapid and simple method to image fluorescently labeled cells in semi-thick brain slices. By fixing, slicing, and optically clearing brain tissue we describe how standard epifluorescent or confocal imaging can be used to visualize individual cells and neuronal networks within intact nervous tissue.

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are ...

Visualizing the Effects of a Positive Early Experience, Tactile Stimulation, on Dendritic Morphology and Synaptic Connectivity with Golgi-Cox Staining

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile stimulation is a positive early experience that mimics maternal licking and grooming in the rat. Exposing rat pups to this positive experience can be completed easily and cost-effectively by using highly accessible materials such as a household duster. Using a cross-litter design, pups are either stroked or left undisturbed, for 15 min, three times per day throughout the perinatal period. To measure the neuroplastic changes related to this positive early experience, Golgi-Cox staining of brain tissue is utilized. Owing to the fact that Golgi-Cox impregnation stains a discrete number of neurons rather than all of the cells, staining of the rodent brain with Golgi-Cox solution permits the visualization of entire neuronal elements, including the cell body, dendrites, axons, and dendritic spines. The staining procedure is carried out over several days and requires that the researcher pay close attention to detail. However, once staining is completed, the entire brain has been impregnated and can be preserved indefinitely for ongoing analysis. Therefore, Golgi-Cox staining is a valuable resource for studying experience-dependent plasticity.

This paper describes the procedures for tactile stimulation of rat pups and subsequent Golgi-Cox staining of neuronal morphology. Tactile stimulation is a positive experience that is administered in the perinatal period by stroking pups with a household duster. Golgi-cox staining is a reliable procedure permitting the visualization of entire neurons.

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile ...

Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity and extent of dendritic arborization correlates well with the synaptic plasticity in these cells. A reliable method for assessment of dendritic arborization is needed to characterize the deleterious effects of neurological disorders on these structures and to determine the effects of therapeutic interventions. However, quantification of these structures has proven to be a formidable task given their complex and dynamic nature. Fortunately, sophisticated imaging techniques can be paired with conventional staining methods to assess the state of dendritic arborization, providing a more reliable and expeditious means of assessment. Below is an example of how these imaging techniques were paired with staining methods to characterize the dendritic arborization in wild type mice. These complementary imaging methods can be used to qualitatively and quantitatively assess dendritic arborization that span a rather wide area within the hippocampal region.

We describe two methods for visualization and quantification of dendritic arborization in the hippocampus of mouse models: real-time and extended depth of field imaging. While the former method allows sophisticated topographical tracing and quantification of the extent of branching, the latter allows speedy visualization of the dendritic tree.

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity ...

Immunostaining of Biocytin-filled and Processed Sections for Neurochemical Markers

Electrophysiological recordings of cells using the patch clamp technique have allowed for the identification of different neuronal types based on firing patterns. The inclusion of biocytin/neurobiotin in the recording electrode permits post-hoc recovery of morphological details, which are necessary to determine the dendritic arborization and the regions targeted by the axons of the recorded neurons. However, given the presence of morphologically similar neurons with distinct neurochemical identities and functions, immunohistochemical staining for cell-type-specific proteins is essential to definitively identify neurons. To maintain network connectivity, brain sections for physiological recordings are prepared at a thickness of 300 &#181;m or greater. However, this thickness often hinders immunohistological postprocessing due to issues with antibody penetration, necessitating the resectioning of the tissue. Resectioning of slices is a challenging art, often resulting in the loss of tissue and morphology of the cells from which electrophysiological data was obtained, rendering the data unusable. Since recovery of morphology would limit data loss and guide in the selection of neuronal markers, we have adopted a strategy of recovering cell morphology first, followed by secondary immunostaining. We introduce a practical approach to biocytin filling during physiological recordings and subsequent serial immunostaining for the recovery of morphology, followed by the restaining of sections to determine the neurochemical identity. We report that sections that were filled with biocytin, fixed with paraformaldehyde (PFA), stained, and coverslipped can be removed and restained with a second primary antibody days later. This restaining involves the removal of the coverslip, the washing of sections in a buffer solution, and the incubation of primary and secondary antibodies to reveal the neurochemical identity. The method is advantageous for eliminating data loss due to an inability to recover morphology and for narrowing down the neurochemical markers to be tested based on morphology.

This protocol presents a method for the morphological recovery of neurons patched during electrophysiological recordings using biocytin filling and subsequent immunohistochemical postprocessing. We show that thick biocytin-filled sections that were stained and coverslipped can be restained with a second primary antibody days&#160;or months later.

Electrophysiological recordings of cells using the patch clamp technique have allowed for the identification of different neuronal types based on firing ...

Large-scale Reconstructions and Independent, Unbiased Clustering Based on Morphological Metrics to Classify Neurons in Selective Populations

This protocol outlines large-scale reconstructions of neurons combined with the use of independent and unbiased clustering analyses to create a comprehensive survey of the morphological characteristics observed among a selective neuronal population. Combination of these techniques constitutes a novel approach for the collection and analysis of neuroanatomical data. Together, these techniques enable large-scale, and therefore more comprehensive, sampling of selective neuronal populations and establish unbiased quantitative methods for describing morphologically unique neuronal classes within a population.
The protocol outlines the use of modified rabies virus to selectively label neurons. G-deleted rabies virus acts like a retrograde tracer following stereotaxic injection into a target brain structure of interest and serves as a vehicle for the delivery and expression of EGFP in neurons. Large numbers of neurons are infected using this technique and express GFP throughout their dendrites, producing &#34;Golgi-like&#34; complete fills of individual neurons. Accordingly, the virus-mediated retrograde tracing method improves upon traditional dye-based retrograde tracing techniques by producing complete intracellular fills.
Individual well-isolated neurons spanning all regions of the brain area under study are selected for reconstruction in order to obtain a representative sample of neurons. The protocol outlines procedures to reconstruct cell bodies and complete dendritic arborization patterns of labeled neurons spanning multiple tissue sections. Morphological data, including positions of each neuron within the brain structure, are extracted for further analysis. Standard programming functions were utilized to perform independent cluster analyses and cluster evaluations based on morphological metrics. To verify the utility of these analyses, statistical evaluation of a cluster analysis performed on 160 neurons reconstructed in the thalamic reticular nucleus of the thalamus (TRN) of the macaque monkey was made. Both the original cluster analysis and the statistical evaluations performed here indicate that TRN neurons are separated into three subpopulations, each with unique morphological characteristics.

This protocol describes large-scale reconstructions of selective neuronal populations, labeled following retrograde infection with a modified rabies virus expressing fluorescent markers, and independent, unbiased cluster analyses that enable comprehensive characterization of morphological metrics among distinct neuronal subclasses.

This protocol outlines large-scale reconstructions of neurons combined with the use of independent and unbiased clustering analyses to create a ...

Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation. There are several approaches to Golgi staining, all of which have been useful for determining the morphological characteristics of dendritic arbors and produce a clear background. The present Golgi-Cox method, (a slight variation of the protocol that is provided with a commercial Golgi staining kit), was designed to assess how a relatively low dose of the chemotherapeutic drug 5-flurouracil (5-Fu) would affect dendritic morphology, the number of spines, and the complexity of arborization within the hippocampus. The 5-Fu significantly modulated the dendritic complexity and decreased the spine density throughout the hippocampus in a region-specific manner. The data presented show that the Golgi staining method effectively stained the mature neurons in the CA1, the CA3, and the dentate gyrus (DG) of the hippocampus. This protocol reports the details for each step so that other researchers can reliably stain tissue throughout the brain with high quality results and minimal troubleshooting.

Here we present a Golgi-Cox protocol in extensive detail. This reliable tissue stain method allows for a high-quality assessment of the cytoarchitecture in the hippocampus, and throughout the entire brain, with minimal troubleshooting.

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching ...

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical ...

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

During neuronal development, axons navigate the cortical environment to reach their final destinations and establish synaptic connections. Growth cones -the sensory structures located at the distal tips of developing axons- execute this process. Studying the structure and dynamics of the growth cone is crucial to understanding axonal development and the interactions with the surrounding central nervous system (CNS) that enable it to form neural circuits. This is essential when devising methods to reintegrate axons into neural circuits following injury in fundamental research and pre-clinical contexts. Thus far, the general understanding of growth cone dynamics is primarily founded on studies of neurons cultured in two dimensions (2D). Although undoubtedly fundamental to the current knowledge of growth cone structural dynamics and response to stimuli, 2D studies misrepresent the physiological three-dimensional (3D) environment encountered by neuronal growth cones in intact CNS tissue. More recently, collagen gels were employed to overcome some of these limitations, enabling the investigation of neuronal development in 3D. However, both synthetic 2D and 3D environments lack signaling cues within CNS tissue, which direct the extension and pathfinding of developing axons. This protocol provides a method for studying axons and growth cones using organotypic brain slices, where developing axons encounter physiologically relevant physical and chemical cues. By combining fine-tuned in utero and ex utero electroporation to sparsely deliver fluorescent reporters along with super-resolution microscopy, this protocol presents a methodological pipeline for the visualization of axon and growth cone dynamics in situ. Furthermore, a detailed toolkit description of the analysis of long-term and live-cell imaging data is included.

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

This protocol demonstrates a straightforward and robust method to study in situ axon growth and growth cone dynamics. It describes how to prepare ex vivo physiologically relevant acute brain slices and provides a user-friendly analysis pipeline.

During neuronal development, axons navigate the cortical environment to reach their final destinations and establish synaptic connections. Growth cones ...

Rapid Golgi Stain for Dendritic Spine Visualization in Hippocampus and Prefrontal Cortex

Golgi impregnation, using the Golgi staining kit with minor adaptations, is used to impregnate dendritic spines in the rat hippocampus and medial prefrontal cortex. This technique is a marked improvement over previous methods of Golgi impregnation because the premixed chemicals are safer to use, neurons are consistently well impregnated, there is far less background debris, and for a given region, there are extremely small deviations in spine density between experiments. Moreover, brains can be accumulated after a certain point and kept frozen until further processing. Using this method any brain region of interest can be studied. Once stained and cover slipped, dendritic spine density is determined by counting the number of spines for a length of dendrite and expressed as spine density per 10 &#181;m dendrite.

The protocol describes a modification of the rapid Golgi method, which can be adapted to any part of the nervous system, for staining neurons in the hippocampus and medial prefrontal cortex of the rat.

Golgi impregnation, using the Golgi staining kit with minor adaptations, is used to impregnate dendritic spines in the rat hippocampus and medial ...

Microscopic Imaging of Neurons in a Mouse Brain Section Using Golgi-Cox Metallic Staining

<a href='https://app.jove.com/v/55358/imaging-neurons-within-thick-brain-sections-using-the-golgi-cox-method'>Source: Louth, E. L.,&#160; et al. Imaging neurons within thick brain sections using the Golgi-Cox method. J. Vis. Exp. (2017)</a>This video demonstrates imaging neurons in a Golgi-Cox stained thick brain section by setting upper and lower focus boundaries and capturing overlapping images. Multiple locations are imaged to ensure detailed neuronal projections, which are combined to create a 3D representation.

Source: Louth, E. L.,&#160; et al. Imaging neurons within thick brain sections using the Golgi-Cox method. J. Vis. Exp. (2017)This video demonstrates ...

Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method

Samenvatting

Meer video's verkennen