This work was supported by a pilot grant under NIH P20 GM109005 (A.R.A.) and by the Center for Translational Neuroscience IDeA program award P30 GM110702.

Experiments were conducted in accordance with the ethical standards approved by the Institutional Animal Care and Use Committee at UAMS.
1. Animals and 5-Fu Injection Paradigm
<ol>
	<li>Purchase 6-month-old male C57Bl6/J wild-type mice and house them under a constant 12 h light/dark cycle until they reach 1 year of age.</li>
	<li>Dilute the 5-Fu in 0.9% sterile saline. Use 60 mg/kg as the required dose per mouse.</li>
	<li>Give the intraperitoneal injections of the 5-Fu (once per week for three weeks).
	<ol>
		<li>Give the intraperitoneal injections around the same time on each day. For example, between 0900-1200 h.</li>
	</ol>
	</li>
	<li>Euthanize the animals and extract the brains 30 days after the final injection.</li>
</ol>
2. Euthanasia Procedure and Brain Extraction
<ol>
	<li>Restrain the rodent and grab the base of the tail. With the other hand, place the thumb or first finger at the base of the rodent&#39;s neck. Quickly put pressure on the rodent&#39;s neck and push forward while the other hand holding the tail pulls backwards.</li>
	<li>Hold the rodent with one hand and the large surgical scissors with the other. Place the rodent&#39;s neck between the two blades and quickly decapitate the head.</li>
	<li>Take the severed mouse head, and use the fine scissors to trim the fur above the skull, up to the eyes. Next, place the tips of the scissors into each eye socket and close the scissors with slight force.</li>
	<li>Use spring scissors to cut the skull to the left and right of the cerebellum.</li>
	<li>Use forceps to lift the portion of the skull surrounding the cerebellum directly up and off.</li>
	<li>Use the spring scissors to cut along the midsagittal line of the skull.</li>
	<li>Use the forceps to lift the remaining portion of the skull off the brain.</li>
	<li>Use a spatula and a probe to scoop out the brain into the desired container. Using a stainless steel blade, cut each brain along the midsagittal plane. Perform the following Golgi-Cox method on the right hemispheres.</li>
</ol>
3. Golgi Staining and Tissue Preparation
<ol>
	<li>Immerse the freshly harvested half-brains into 5 mL mercuric chloride-based solution (solution A from the kit) in a 10 mL conical tube. The mercuric chloride solution is light-sensitive; cover the tube with foil. Store the sample in the dark at room temperature. 
	NOTE: Solution A is provided in the kit that is listed in the table of materials. Caution! Solution A (also called mercuric chloride solution) contains toxic reagents such as potassium dichromate, mercuric chloride, and potassium chromate. Wear protective gloves and work in a fume hood.</li>
	<li>After 1 day, slowly decant the solution into a temporary container (such as a large weigh boat), and dispose of it in a biohazard waste container. Immerse the brains (still in the original 10 mL conical tubes) with 5 mL of the mercuric chloride solution, and return the sample to the dark at room temperature for 13 more days.</li>
	<li>Add 30 g post-impregnation buffer to 90 mL distilled or deionized water (dH2O). Fill each well of a 6-well plate with 6.5 mL of the post-impregnation buffer, one well per brain 
	NOTE: Post-impregnation buffer is provided in the kit that is listed in the table of materials.
	<ol>
		<li>After 14 days (in total) in the mercuric chloride solution, rinse the tissue with dH2O, then transfer them into 6-well plates with post-impregnation buffer. Cover the plates with foil and store them at room temperature in the dark.</li>
	</ol>
	</li>
	<li>Pipette out the post-impregnation buffer after 1 day of immersion and renew with fresh post-impregnation buffer; store for 1 day in the dark at room temperature. If required, store this at 4 &#176;C for up to 1 month22.</li>
</ol>
4. Sectioning
<ol>
	<li>Label the 12-well plates on their lids with numbers in ascending order from left-to-right and up-to-down.</li>
	<li>Prepare two or three 12-well plates per brain if sectioning the entire brain.
	<ol>
		<li>Label the top of each 12-well plate lid with the identification number of the brain sectioned.</li>
		<li>Pipette 2 mL of 1x PBS into each well of each plate.</li>
	</ol>
	</li>
	<li>Set up the stage and turn on the vibratome; ensure that there is enough 1x PBS in the container to cover the tissue.
	<ol>
		<li>Cut two pieces of plastic paraffin film and fold them over twice. Place them over the holes for the specimen holder knobs to prevent 1x PBS from leaking onto the counter.</li>
		<li>Put tape down on the tissue block of the specimen holder and then put cyanoacrylate adhesive glue (see table of materials) on it.</li>
	</ol>
	</li>
	<li>Remove the brain from the 6-well plate and place it on a plastic or glass dish. Use the stainless steel double-edged blade to cut off about half of the cerebellum.
	<ol>
		<li>Cut the cerebellum caudally. Make sure that the portion of the cerebellum remaining is even.</li>
	</ol>
	</li>
	<li>Immediately place the flat surface of the remaining cerebellum on the glue. 
	NOTE: The dorsal part of the tissue (the cerebral cortex) should be facing left or right relative to the blade. The rostral end that previously contained the olfactory bulb attached should face upwards.
	<ol>
		<li>Wait at least a few minutes to make sure that the glue fully dries.</li>
	</ol>
	</li>
	<li>While the glue that is holding the brain in place is drying, pour 1x PBS into the specimen bath. After the glue is dry, place the specimen holder with the tissue into the specimen bath.
	<ol>
		<li>If the tissue sample is not fully covered, pour more 1x PBS into the specimen bath.</li>
	</ol>
	</li>
	<li>Attach the blade to the blade holder of the vibratome and slowly lower the blade into the specimen bath until it is fully submerged in 1x PBS. Continue lowering the blade until it is slightly below the top of the tissue.</li>
	<li>Set the vibratome speed to 7, the amplitude to 6, and the cutting angle to 12&#176; (see table of materials for vibratome model). Start the vibration and cut 200 &#181;m sections23.
	<ol>
		<li>Use a large paintbrush to move the tissue sections from the specimen bath into each designated well of the 12-well plates.</li>
	</ol>
	</li>
	<li>Continue to cut the tissue until the desired number of tissue sections is reached.</li>
</ol>
5. Post-staining
<ol>
	<li>For each of the following staining steps and rinses, use 2 mL of the indicated solution per well. Use a motorized pipet filler (see table of materials) to transfer the solutions into the wells. Use a transfer pipette (see table of materials) to transfer solutions out of the wells and into the proper waste containers.</li>
	<li>Rinse the sections in one 30 min wash of 0.01 M PBS-T (add 3.0 mL of detergent (see table of materials) to 1,000 mL 0.01 M PBS).</li>
	<li>Dilute ammonium hydroxide solution (Caution) 3:5 by volume with dH2O immediately before use. Stain the sections in the diluted ammonium hydroxide solution for 19-21 min.
	<ol>
		<li>Protect the light-sensitive ammonium hydroxide solution with foil. 
		NOTE: Ammonium hydroxide within solution has a concentration of ~10% and it is classified as &#34;hazardous.&#34; Ammonium hydroxide solution may produce burns if it contacts the skin. It may cause respiratory tract irritation if inhaled. Wear protective gloves and work in a fume hood.</li>
	</ol>
	</li>
	<li>Stain the sections in the post-staining buffer (add 90 g of reagent D in 500 mL dH2O) for 19-21 min. The post-staining buffer is light-sensitive; protect it with foil22.</li>
	<li>Rinse the sections in three, 5-10 min washes of 0.01 M PBS-T.</li>
</ol>
6. Mounting, Cleaning, and Covering
<ol>
	<li>Mount the sections on 1% gelatin-coated slides with the large paintbrush. Allow them to dry for 20-30 min at room temperature, and then place them in a Coplin jar overnight.</li>
	<li>Remove the slides with gloved hands from the Coplin jar and place them in a plastic rack (see table of materials). Place the plastic rack in the staining dish (see table of materials) containing 100% ethanol. Dehydrate them with three 5 min washes.</li>
	<li>Wash the slides twice for 5 min each in 99% xylene. Place the slides in a glass rack and lower the rack into a glass staining dish containing xylene with a metal handle (see table of materials). 
	NOTE: Caution, Xylene is harmful if inhaled and causes irritation if it touches the skin. It is highly flammable in liquid and vapor states. Wear protective gloves and work in a fume hood.</li>
	<li>Remove the slides from the xylene one at a time and quickly cover the tissue with ~0.25 mL of mounting media (see Material Table). Next, take the slide covers and lay them over the media.
	<ol>
		<li>Push any trapped air bubbles underneath the slide covers to the edge with a blunt object, such as the end of a paintbrush.</li>
	</ol>
	</li>
	<li>Place the slides in an area away from sunlight and allow them to dry for 2-3 days.</li>
	<li>Proceed to acquiring images using a microscope and analyzing with the appropriate software.</li>
</ol>
7. Acquire Image Stack
<ol>
	<li>Open the imaging program (see Materials Table). Place the slide onto the stage of the microscope. On the menu at the top of the program, click &#34;Acquisition&#34; and then click &#34;Live Image.&#34;</li>
	<li>Set the microscope to 10X. Identify the neuron of preference and then bring the cursor, which appears like a red X, onto the center of the soma. Left click to set the reference point.</li>
	<li>Set the microscope to 40X. Click &#34;Move&#34; from the menu at the top of the program and then click &#34;To Reference Point.&#34;</li>
	<li>Begin creating the image stack by manually scrolling with the fine objective above the tissue until it is slightly out of focus.
	<ol>
		<li>Click &#34;Set Top&#34; in the bottom right hand corner of the program under the heading &#34;Image Acquisition.&#34;</li>
		<li>Scroll slightly under the tissue until it is slightly out of focus and then click &#34;Set Bottom&#34; under &#34;Image Acquisition.&#34; Next, click &#34;Acquire Image Stack.&#34;</li>
	</ol>
	</li>
	<li>After the image stack is acquired, type the desired name for image file in the window that appears. Save as &#34;MBF Ascii file.&#34;
	<ol>
		<li>When prompted, select the file type as &#34;MBF JPEG2000 stack file&#34; and then click &#34;Save.&#34;</li>
	</ol>
	</li>
</ol>
8. Neuronal Tracing
<ol>
	<li>In the toolbar underneath the menu, click the box titled &#34;Contour Name.&#34; Select &#34;Soma&#34; from the drop down menu.</li>
	<li>Manually trace the soma. Then right click and select &#34;Finish Cell Body&#34; when tracing is complete.</li>
	<li>Select &#34;AutoNeuron&#34; to bring up another tab titled &#34;AutoNeuron Workflow.&#34;</li>
	<li>Under &#34;Configuration Type,&#34; select &#34;New.&#34; Set the parameters and then click &#34;Next Step&#34; to bring up &#34;Soma Detection&#34; subheading.
	<ol>
		<li>To set the parameters, select &#34;Brightfield&#34;. Next, under &#34;Max Process Diameter&#34;, click &#34;Start Measuring&#34;. Using the cursor, go to the base of the dendrite and manually set the diameter.</li>
	</ol>
	</li>
	<li>Click &#34;Yes&#34; in the window that requests the user to trace the entire image. Manually trace the soma. Unclick and then click &#34;Next step&#34; to bring up &#34;Seed Placement&#34; subheading.</li>
	<li>Click &#34;Validate Seeds&#34; and then click &#34;Next Step&#34; to bring up &#34;Neuron Reconstruction&#34; subheading.</li>
	<li>Under &#34;Neuron&#34;, click &#34;Interactive.&#34; Perform interactive tracing by following the direction from the program of the dendrites and right-clicking to complete the highlighted area traced by the program. After tracing all the dendrites, click &#34;Next Step.&#34;</li>
	<li>After a new window appears, save the new configuration with a desired title. Click &#34;Save.&#34;</li>
</ol>
9. Analysis
<ol>
	<li>Open the analysis program (see the table of materials).</li>
	<li>Click &#34;File&#34; from the top menu and then click &#34;Open Data File&#34;. Select the file of interest.</li>
	<li>Under the analysis subheading, select &#34;Sholl Analysis.&#34;</li>
	<li>In the &#34;Sholl Analysis&#34; window, set the starting radius to 10 &#181;m. Under &#34;Analysis,&#34; click the boxes &#34;Dendrites&#34; and &#34;Branch Orders.&#34;</li>
	<li>Right click the newly opened windows and click &#34;Excel Export.&#34; Click &#34;Save.&#34;</li>
	<li>Under the analysis, click &#34;Branched Structure Analyses.&#34; Select &#34;All Possible Analyses.&#34;</li>
	<li>Right click the newly opened window and click &#34;Excel Export&#34;24.</li>
</ol>

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The authors declare that they have no competing financial interests.

Compared to more modern techniques, the Golgi-Cox method has several advantages that make it the preferred method for examining spine morphology: 1) The staining can be used for essentially any tissue, 2) A basic light microscope setup is all that is needed to acquire Golgi-based images, 3) The Golgi-Cox imaging is faster than confocal imaging, and 4) Golgi stained sections are viable for several months to years longer than samples that are fluorescently-labeled. Even with these advantages, the Golgi-Cox method still has certain limitations. First, the entire process is very time consuming, requiring several weeks of staining followed by analyzing the images. As seen with this protocol, it takes 14 days in solution A and 1 day in solution B before the sectioning can begin. Additionally, performing sectioning, post-staining, and mounting will take 2-3 hours per brain. Each slide must dry for 1 day before cleaning and covering them. After which, the amount of time it will take to image and analyze the sections will depend on the individual. Second, there may be issues of consistency between the data from analysts due to differences in categorizing spines30. For this reason, it is recommended that one person performs the imaging and analysis portion of each experiment, which further extends the amount of time for completion of the project.
There are several critical steps of the Golgi-Cox method that require exact timing and attention. When staining the sections with the ammonium hydroxide solution, it is imperative that the sections remain in the solution for at least 19 min, but not over 21 min. If the sections are not in the ammonium hydroxide solution for enough time, they will not be properly stained, meaning that the neurons will look less defined when imaging, thus making it harder to analyze them. If the sections are in the ammonium hydroxide solution for over 21 min, they may become over-stained, thus making it difficult to identify the individual neurons. Therefore, when switching out the solutions, the researcher must move quickly. Another important step is the transfer of the sections onto slides. Before moving the sections with the large paintbrush, it is best to first add a small amount of the PBS-T onto the slide. This allows the sections to easily be placed onto the slide and maneuvered without potentially breaking apart. One more critical step is the drying step after the sections are placed on a slide. If the slides do not dry for at least 20 min at room temperature, then certain sections will most likely fall off during the dehydration steps. If the slides dry for longer than 30 min, they will most likely become brittle and break apart during the dehydration steps.
When performing this procedure, there are certain steps that may be troublesome. For example, if the entire cerebellum is cut off before placing the brain on the specimen holder and sectioning, the brain may crack apart before the DG is fully sectioned, rendering any further sections unusable. Therefore, it is important to only cut caudally through about half of the cerebellum. Another step that may negatively affect the sections is the vibratome speed and the amplitude. While the recommended settings are for 7 and 6 respectively, upon cutting, certain sections may disintegrate in the PBS or appear uneven. To correct this, finely adjust either the vibratome speed or the amplitude, one at a time until the sections are in-tact and even, between settings of 6-8.
Even though commercial kits for Golgi staining are available, they often lack exact details for each individual step. With this protocol, we report in detail all the steps that were used to allow other labs to reliably stain tissue at a high quality with minimal troubleshooting. The materials used are available to most neuroscience labs. Aberrations in dendrite morphology and alterations in the number of dendritic spines are related to a multitude of neurological disorders31. Furthermore, these perturbations of the dendritic arborization might represent a significant morphological hallmark of the chemotherapy induced injury in the brain32. In the future, we will examine how the structural alterations induced by 5-Fu may relate to the behavioral, cellular, and molecular changes.

Dendrites are the largest portion of neurons that receive and process presynaptic input1. Their dendritic processes have a complex geometry, where the proximal branches have a larger diameter than the distal branches. As dendrites develop, they form several connections with other neurons in a process referred to as dendritic arborization. The extent and pattern of this branching determines the amount of synaptic inputs that a dendrite can adequately process2.
Dendritic arborization is a necessary process for activity-dependent plasticity and proper development of neuronal circuits. Extension, retraction, branching, and synaptogenesis are intricate processes that include intrinsic genetic programs and influences from extrinsic factors. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation3,4. The alterations in dendritic complexity are associated with pathophysiological and behavioral changes5. Abnormalities are related to several disease states, including Fragile X Syndrome and Down Syndrome6.
Dendritic spines are the specialized subcellular compartments of the dendritic arbors that receive excitatory input within the central nervous system. There are three morphological classes of dendritic spines, with the name of each class based on their size and shape: 1) mushroom spines, which have complex postsynaptic densities with more glutamate receptors than other spines7; 2) stubby spines, which lack a stem; and 3) thin spines, which consist of a protracted, narrow stem and a globular head8. The dendritic spine volume is used in part to define them, with thin spines generally smaller (0.01 &#181;m3) compared to mushroom spines (0.8 &#181;m3)9,10. The spines stabilize with maturation. For example, the thin spines either retract after a few days or develop into mushroom spines. Alternatively, the mushroom spines are relatively stable and can survive for an extended period.The strength of the neuronal connections is thought to be based on the number of spines and/or their volume11,12,13.
The classical Golgi staining method and its more modern variations have all been useful for examining the dendritic spine morphology and density. One unique aspect of the Golgi staining is that it randomly stains about 5% of the total neurons, which allows for the tracing of individual neurons14,15. Although the exact mechanism in which the Golgi method stains individual neurons is still unknown, the principle of the method is based on the crystallization of silver chromate (Ag2CrO4)16,17. There are three main types of the Golgi method: the rapid Golgi, the Golgi-Cox, and the Golgi-Kopsch18,19. All three methods start with an initial incubation phase in chromium salts for several days to months, but there are certain key differences between them. The rapid Golgi uses osmium tetroxide in the first step, whereas the Golgi-Kopsch includes paraformaldehyde. The staining in both the rapid-Golgi and the Golgi-Kopsch is followed by an incubation in a 1-2% silver nitrate solution for about 7 days. The Golgi-Cox method uses mercuric chloride and potassium dichromate instead of silver nitrate and has an impregnation time of 2-4 weeks. The tissues are then sectioned and quickly placed in a diluted ammonia solution, followed by a photographic fixer to remove salts. Of the three types, the Golgi-Cox method is thought to be the best at staining the dendritic arbors without much background interference, in part, because the crystal artifacts do not occur on the surface of the tissue (unlike in the rapid Golgi method)17,20,21.
The present method is a slight variation of the protocol provided with a commercial Golgi staining kit, and was designed to assess how a relatively low dose of the 5-Fu would affect the dendritic morphological characteristics and the spine density. Any data acquired could provide further insight into how chemotherapeutic treatment affects the neuronal circuitry.

<table><tbody><tr><td>superGolgi Kit&amp;nbsp;</td><td>Bioenno Lifesciences</td><td>30100</td><td>&amp;nbsp;Contains hazardous materials.&amp;nbsp;</td></tr><tr><td>PBS 10x powder concentrate</td><td>Fisher&amp;nbsp;</td><td>BP665-1</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>9002-93-1</td><td></td></tr><tr><td>Permount&amp;nbsp;</td><td>Fisher&amp;nbsp;</td><td>SP 15-100</td><td></td></tr><tr><td>Slide cover&amp;nbsp;</td><td>Fisher&amp;nbsp;</td><td>12-546-14</td><td></td></tr><tr><td>7 mL Transfer pipette&amp;nbsp;</td><td>Globe Scientific&amp;nbsp;</td><td>135030</td><td></td></tr><tr><td>10 mL Falcon tubes&amp;nbsp;</td><td>BD Biosciences&amp;nbsp;</td><td>352099</td><td></td></tr><tr><td>Foil&amp;nbsp;</td><td>Fisher&amp;nbsp;</td><td>01-213-105</td><td></td></tr><tr><td>12-well plate&amp;nbsp;</td><td>BD Biosciences&amp;nbsp;</td><td>353043</td><td></td></tr><tr><td>200 proof Ethanol&amp;nbsp;</td><td>Pharmco-AAPER</td><td>111000200</td><td></td></tr><tr><td>Xylene&amp;nbsp;</td><td>Acros Organics&amp;nbsp;</td><td>1330-20-7</td><td>Hazardous.&amp;nbsp;</td></tr><tr><td>Permabond 200</td><td>Permabond LLC</td><td>GF2492</td><td></td></tr><tr><td>25 mL serological pipette</td><td>Sigma</td><td>SIAL1489</td><td></td></tr><tr><td>Parafilm</td><td>Midsci</td><td>HS234526C&amp;nbsp;</td><td></td></tr><tr><td>Vibratome&amp;nbsp;</td><td>World Precision Instruments&amp;nbsp;</td><td>NVSLM1</td><td></td></tr><tr><td>C57Bl/6 Male Mice&amp;nbsp;</td><td>The Jackson Laboratory&amp;nbsp;</td><td>000664</td><td></td></tr><tr><td>Axio Imager 2</td><td>ZEISS</td><td>Multiple components, see website for details.&amp;nbsp;</td><td></td></tr><tr><td>AxioCam MRc Camera</td><td>ZEISS</td><td>426508-9902-000</td><td></td></tr><tr><td>Staining Dish , Green</td><td>Tissue-Tek</td><td>62541-12</td><td></td></tr><tr><td>Staining Dish Set&amp;nbsp;</td><td>Electron Microscopy Sciences&amp;nbsp;</td><td>70312-20</td><td></td></tr><tr><td>Motorized Pipet Filler&amp;nbsp;</td><td>Fisher&amp;nbsp;</td><td>03-692-168</td><td></td></tr><tr><td>Neurolucida&amp;nbsp;</td><td>mbf Bioscience&amp;nbsp;</td><td></td><td></td></tr><tr><td>Neurolucida Explorer&amp;nbsp;</td><td>mbf Bioscience&amp;nbsp;</td><td></td><td></td></tr><tr><td>Prism&amp;nbsp;</td><td>GraphPad</td><td></td><td></td></tr></tbody></table>

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.

The effects of the 5-Fu treatment on the dendritic arborization and complexity in the hippocampus of Golgi-stained brain sections were quantified and traced using a commercially available imaging software. After tracing, the dendritic arborization, the spine density, and the spine morphology were analyzed using Sholl analysis and the dendritic complexity index (DCI). Sholl analysis is a quantitative analytical method that can be used to determine the dendritic arbor morphology25. Starting from the soma, circles 10 &#181;m apart from each other overlay the dendritic tracings. The length of the dendrites is determined by the number of circles that the dendrites cross over. The branch point analysis, a method to ascertain the complexity of a dendritic arbor, is based on the total number and order of branch points.
A branch point is defined as when a branch splits into two sub-branches. The branch order is a measure of the complexity based on how many times the branches divide. For example, a first-order branch point would be the original dendrite extending from the soma, while a second-order branch point would be the point at which the branch divides into two sub-branches. By examining both the branch points and the branch order, the complexity of the dendritic arbor (based on the number of branch points) and at what points the branching occurs (a smaller branch order is closer to the soma, while a larger branch order is further from the soma) can be determined. These parameters were applied to the DCI. The CA1 and CA3 were divided into apical and basal portions and analyzed independently26,27.
The dendrites of the CA1 and CA3 pyramidal neurons and granule neurons in the DG were analyzed. All neurons selected for each experimental group fulfilled the following criteria: 1) the dendrites had dark and consistent Golgi staining across their entire length, 2) the dendrites were visibly in-tact, and 3) each neuron had enough space between them to prevent interference during the analysis28. Figure 1 demonstrates a neuron that fulfills the above criteria. Data were expressed as mean &#177; SEM. To determine the statistical differences between the sham and the 5-Fu groups, a paired two-tailed t-test was used. All the statistical analyses were conducted using analytical software (see table of materials), and p &#60;0.5 was considered significant. A mixed-factors analysis of variance (ANOVA) was used to evaluate the effects of the 5-Fu treatment and the Sholl radius. Fisher LSD post-hoc tests were used following ANOVA when appropriate29.
In both CA1 (t = 7.68, p &#60;0.01; Figure 2A) and CA3 (t = 7.54, p &#60;0.01; Figure 3A) apical pyramidal dendrites, there was a significant overall reduction in spines following the 5-Fu treatment. While the 5-Fu treatment did not significantly affect the basal spine density of CA1 (t = 1.79, p = 0.15; Figure 2B), it did significantly decrease CA3 basal pyramidal dendritic spines (t = 5.57, p &#60; 0.05; Figure 3B). Furthermore, the Fisher LSD post-hoc tests revealed a decrease in the dendritic arborization of the CA1 apical pyramidal dendrites at 40-100 &#181;m (Fisher&#39;s LSD, p &#60; 0.001; Figure 2C) and 130-160 &#181;m (Fisher&#39;s LSD, p &#60;0.05; Figure 2C) away from the soma compared to the saline treated controls. A similar phenomenon was seen in the CA1 basal apical dendrites at 30-60 &#181;m (Fisher&#39;s LSD, p &#60;0.001; Figure 2D) and 70-110 &#181;m (Fisher&#39;s LSD, p &#60;0.05; Figure 2D) away from the soma29.
Regarding the CA3 apical pyramidal dendrites, there was not a significant difference in the segmental dendritic length following the 5-Fu treatment compared to the saline treated controls (F (28,87) = 0.91; p = 0.59; Figure 3C). However, for the CA3 basal pyramidal dendrites, there was a significant difference between the 5-Fu treatment and the saline treated controls in the segmental dendritic length (F (25, 78) = 1.85; p &#60;0.05; Figure 3D). The Fisher&#39;s LSD revealed that the 5-Fu treatment decreased the dendritic arborization at 90-100 &#181;m (p &#60;0.01; Figure 3D), 70-80 &#181;m and 110-120 &#181;m (Fisher&#39;s LSD, p &#60;0.05; Figure 3D) away from the soma29.
To determine the dendritic complexity, a branch order analysis was used to compare the 5-Fu treated and the saline treated control groups. Within the DG, the branch order analysis showed a significant difference between each treatment and branch order (F (8, 36) = 25.61, p &#60; 0.001; Figure 4). This was further analyzed using the Fisher&#39;s LSD, which indicated that there was decreased length at the 4th and 6th order following the 5-Fu treatment (p &#60; 0.001; Figure 4)29.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55696/55696fig1.jpg" /> 
Figure 1: Golgi stained neuron that represents the criteria for imaging and further analysis. All three spine types can easily be seen following the Golgi staining. Staining is consistent across the entire length of the dendrite and the dendrite is isolated from other neurons. Scale bar, 5 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55696/55696fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55696/55696fig2.jpg" /> 
Figure 2: Golgi stained neurons demonstrated significant differences in spine density between the 5-Fu treated and the saline treated groups in the apical CA1, but not the basal CA1. There was a significant difference in the spine length between 5-Fu treated and saline treated groups in both apical and basal CA1. (a) In the CA1 apical pyramidal dendrites, 5-Fu significantly decreased spine density. (b) In the basal dendrites, there were no significant changes in the overall density of spines. (c) Sholl analysis revealed that the 5-Fu treatment significantly decreased dendritic arborization at 40-100 &#181;m and 130-160 &#181;m away from the soma in the CA1 apical pyramidal dendrites. (d) In the basal dendrites, the 5-Fu treatment decreased the dendritic arborization at 30-60 &#181;m and 70-110 &#181;m away from the soma. Average &#177; SEM (n = 6). *p &#60;0.05; **p &#60;0.01, ANOVA29. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55696/55696fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/55696/55696fig3.jpg" /> 
Figure 3: Golgi stained neurons demonstrated significant differences in spine density between 5-Fu treated and saline treated groups in both apical and basal CA3. There was a significant difference in spine length between 5-Fu treated and saline treated groups in apical CA3, but not basal CA3. (a) In the CA3 apical pyramidal dendrites, 5-Fu significantly decreased spine density. (b) In the basal dendrites, 5-Fu significantly decreased the overall density of spines. (c) There was no significant effect of 5-Fu treatment in the CA 3 apical area. (d) In the basal dendrites, 5-Fu treatment decreased dendritic arborization 70-120 &#181;m away from the soma. Average &#177; SEM (n = 6). *, p &#60;0.05; **, p &#60;0.01, ANOVA29. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55696/55696fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/55696/55696fig4.jpg" /> 
Figure 4: Golgi stained neurons within the DG showed decreased length at the 4th and 6th order following 5-Fu treatment compared to the saline treated group. Length per branch order in pyramidal neurons of the CA1 hippocampal region. Average &#177; SEM (n = 6). *p &#60;0.05; **p &#60;0.01, ANOVA29. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55696/55696fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method at JoVE.com

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

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 ...

assessment-hippocampal-dendritic-complexity-aged-mice-using-golgi-cox

Nanyang Technological University

Research

JoVE Journal

Neuroscience

15.0K Views.  University of Arkansas for Medical Sciences. 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.

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

Preparation of Acute Hippocampal Slices from Rats and Transgenic Mice for the Study of Synaptic Alterations during Aging and Amyloid Pathology

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The hippocampus can be extracted quickly and easily from rats and mice and slices remain viable for hours in oxygenated artificial cerebrospinal fluid. Moreover, basic electrophysisologic techniques are easily applied to the investigation of synaptic function in hippocampal slices and have provided some of the best biomarkers for cognitive impairments. The hippocampal slice is especially popular for the study of synaptic plasticity mechanisms involved in learning and memory. Changes in the induction of long-term potentiation and depression (LTP and LTD) of synaptic efficacy in hippocampal slices (or lack thereof) are frequently used to describe the neurologic phenotype of cognitively-impaired animals and/or to evaluate the mechanism of action of nootropic compounds. This article outlines the procedures we use for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and Alzheimer's disease (AD)1-3. Use of aged rats and AD model mice can present a unique set of challenges to researchers accustomed to using younger rats and/or mice in their research. Aged rats have thicker skulls and tougher connective tissue than younger rats and mice, which can delay brain extraction and/or dissection and consequently negate or exaggerate real age-differences in synaptic function and plasticity. Aging and amyloid pathology may also exacerbate hippocampal damage sustained during the dissection procedure, again complicating any inferences drawn from physiologic assessment. Here, we discuss the steps taken during the dissection procedure to minimize these problems. Examples of synaptic responses acquired in "healthy" and "unhealthy" slices from rats and mice are provided, as well as representative synaptic plasticity experiments. The possible impact of other methodological factors on synaptic function in these animal models (e.g. recording solution components, stimulation parameters) are also discussed. While the focus of this article is on the use of aged rats and transgenic mice, novices to slice physiology should find enough detail here to get started on their own studies, using a variety of rodent models.

This article outlines procedures for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and age-related neurodegenerative diseases, such as Alzheimer&#x2019;s disease.

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The ...

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 ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

Biology

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 ...

Immunohistochemical Visualization of Hippocampal Neuron Activity After Spatial Learning in a Mouse Model of Neurodevelopmental Disorders

Induction of phosphorylated extracellular-regulated kinase (pERK) is a reliable molecular readout of learning-dependent neuronal activation. Here, we describe a pERK immunohistochemistry protocol to study the profile of hippocampal neuron activation following exposure to a spatial learning task in a mouse model characterized by cognitive deficits of neurodevelopmental origin. Specifically, we used pERK immunostaining to study neuronal activation following Morris water maze (MWM, a classical hippocampal-dependent learning task) in Engrailed-2 knockout (En2-/-) mice, a model of autism spectrum disorders (ASD). As compared to wild-type (WT) controls, En2-/- mice showed significant spatial learning deficits in the MWM. After MWM, significant differences in the number of pERK-positive neurons were detected in specific hippocampal subfields of En2-/- mice, as compared to WT animals. Thus, our protocol can robustly detect differences in pERK-positive neurons associated to hippocampal-dependent learning impairment in a mouse model of ASD. More generally, our protocol can be applied to investigate the profile of hippocampal neuron activation in both genetic or pharmacological mouse models characterized by cognitive deficits.

We describe an immunohistochemistry protocol to study the profile of hippocampal neuron activation following exposure to a spatial learning task in a mouse model characterized by cognitive deficits of neurodevelopmental origin. This protocol can be applied to both genetic or pharmacological mouse models characterized by cognitive deficits.

Induction of phosphorylated extracellular-regulated kinase (pERK) is a reliable molecular readout of learning-dependent neuronal activation. Here, we ...

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.

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 ...

Assaying Circuit Specific Regulation of Adult Hippocampal Neural Precursor Cells

Adult neurogenesis is a dynamic process by which newly activated neural stem cells (NSCs) in the subgranular zone (SGZ) of the dentate gyrus (DG) generate new neurons, which integrate into an existing neural circuit and contribute to specific hippocampal functions. Importantly, adult neurogenesis is highly susceptible to environmental stimuli, which allows for activity-dependent regulation of various cognitive functions. A vast range of neural circuits from various brain regions orchestrates these complex cognitive functions. It is therefore important to understand how specific neural circuits regulate adult neurogenesis. Here, we describe a protocol to manipulate neural circuit activity using designer receptor exclusively activated by designer drugs (DREADDs) technology that regulates NSCs and newborn progeny in rodents. This comprehensive protocol includes stereotaxic injection of viral particles, chemogenetic stimulation of specific neural circuits, thymidine analog administration, tissue processing, immunofluorescence labeling, confocal imaging, and imaging analysis of various stages of neural precursor cells. This protocol provides detailed instructions on antigen retrieval techniques used to visualize NSCs and their progeny and describes a simple, yet effective way to modulate brain circuits using clozapine N-oxide (CNO) or CNO-containing drinking water and DREADDs-expressing viruses. The strength of this protocol lies in its adaptability to study a diverse range of neural circuits that influence adult neurogenesis derived from NSCs.

The goal of this protocol is to describe an approach for analyzing behavior of adult neural stem/progenitor cells in response to chemogenetic manipulation of a specific local neural circuit.

Adult neurogenesis is a dynamic process by which newly activated neural stem cells (NSCs) in the subgranular zone (SGZ) of the dentate gyrus (DG) generate ...

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high fidelity. Exploration of LTP and LTD mechanisms, single neuron dendritic computation, and experience-dependent changes in circuitry, would not have been possible without this classical preparation. However, with a few exceptions, most basic research using acute hippocampal slices has been performed using slices from rodents of relatively young ages, ~P20-P40, even though synaptic and intrinsic excitability mechanisms have a long developmental tail that reaches past P60. The main appeal of using young hippocampal slices is preservation of neuronal health aided by higher tolerance to hypoxic damage. However, there is a need to understand neuronal function at more mature stages of development, further accentuated by the development of various animal models of neurodegenerative diseases that require an aging brain preparation. Here we describe a modification to an acute hippocampal slice preparation that reliably delivers healthy slices from adult and aging mouse hippocampi. The protocol&#8217;s critical steps are transcardial perfusion and cutting with ice-cold sodium-free NMDG-aSCF. Together, these steps attenuate the hypoxia-induced drop in ATP upon decapitation, as well as cytotoxic edema caused by passive sodium fluxes. We demonstrate how to cut transversal slices of hippocampus plus cortex using a vibrating microtome. Acute hippocampal slices obtained in this way are reliably healthy over many hours of recording, and are appropriate for both field-recordings and targeted patch-clamp recordings, including targeting of fluorescently labeled neurons.

This is a protocol for acute slice preparation from adult and aging mouse hippocampi that takes advantage of transcardial perfusion and slice cutting with ice-cold NMDG-aCSF to reduce hypoxic damage to the tissue. The resulting slices stay healthy over many hours, and are suitable for long-term patch-clamp and field-recordings.

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high ...

Preparation of Acute Slices from Dorsal Hippocampus for Whole-Cell Recording and Neuronal Reconstruction in the Dentate Gyrus of Adult Mice

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in molecular, anatomical and functional criteria suggesting a division into different sub-circuits along its rostro-caudal extent. Owing to differential connectivity and function the most fundamental distinction is made between the dorsal and the ventral hippocampus, which are preferentially involved in spatial and emotional processing, respectively. Accordingly, in vivo work regarding spatial memory formation has focused on the dorsal hippocampus.
In contrast, electro-physiological in vitro recordings have been preferentially performed on intermediate-ventral hippocampus, largely motivated by factors like slice viability and circuit integrity. To allow for direct correlation of in vivo data on spatial processing with in vitro data we have adapted previous sectioning methods to obtain highly viable transverse brain slices from the dorsal-intermediate hippocampus for long-term recordings of principal cells and interneurons in the dentate gyrus. As spatial behavior is routinely analyzed in adult mice, we have combined this transversal slicing procedure with the use of protective solutions to enhance viability of brain tissue from mature animals. We use this approach for mice of about 3 months of age. The method offers a good alternative to the coronal preparation which is frequently used for in vitro studies on dorsal hippocampus. We compare these two preparations in terms of quality of recordings and preservation of morphological features of recorded neurons.

We present a protocol to prepare acute-slices from the dorsal-intermediate hippocampus of mice. We compare this transversal preparation with coronal slicing in terms of quality of recordings and preservation of morphological features of recorded neurons.

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in ...

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.