This work was funded by NIH grants HD043680, MH106392, DA013137, and NS100624.

All animal protocols were reviewed and approved by the Animal Care and Use Committee at the University of South Carolina (federal assurance number: D16-00028).
1. Preparation of DiI/Tungsten bead tubing
<ol>
	<li>Dissolve 100 mg of polyvinylpyrrolidone (PVP) with 10 mL of ddH2O. Vortex the PVP solution lightly.</li>
	<li>Fill the tubing with the PVP solution (see Table of Materials) and leave it for 20 min. Then, expel the PVP solution through the other end of the tubing using a 10 mL syringe.</li>
	<li>Combine 170 mg of tungsten microcarrier beads with 250 &#181;L of methylene chloride. Vortex the tungsten bead suspension thoroughly.</li>
	<li>Combine 6 mg of lipophilic DilC18(3) dye with 300 &#181;L of methylene chloride. Vortex the DilC18(3) dye solution thoroughly. 
	NOTE: Perform steps 1.3&#8211;1.4 in a fume hood.</li>
	<li>Pipette 250 &#181;L of the tungsten bead suspension onto a glass slide. Wait for the suspension to air dry (~3 min).</li>
	<li>Add 300 &#181;L of DilC18(3) dye solution on the top of the tungsten bead suspension layer. Mix the DiIC18(3) dye solution and tungsten bead suspension thoroughly with a pipette tip and allow the mixture to air dry (~3 min).</li>
	<li>Once dried, use a razor blade to split the mixture into two 1.5 mL centrifuge tubes. Fill the tubes with water.</li>
	<li>Sonicate in a water bath (Amp I, 100%) until homogenous (~5 min). Ensure that the tip of the sonicator lies directly on the tubes with the bead suspension.</li>
	<li>Combine the two 1.5 mL of homogenized mixture into a 15 mL conical tube. Sonicate the mixture for another 3 min.</li>
	<li>Draw the tungsten beads-DilC18(3) dye mixture into the PVP-coated tubing using a 10 mL syringe. Feed the tubing into the preparation station (see Table of Materials).</li>
	<li>Rotate for 1 min on the preparation station. Carefully remove all the water from the tubing using a 10 mL syringe.</li>
	<li>Turn on the nitrogen gas and adjust the nitrogen flow to approximately 0.5 Liters per minute (LPM), rotate the tubing in the preparation station, and dry with nitrogen for 30 min.</li>
	<li>Remove the tubing from the preparation station and cut into 13 mm lengths (matching the loading size of the cartridge) using a tubing cutter. Keep the 13 mm lengths in scintillation vials in the dark.</li>
</ol>
2. Preparation of brain sections
NOTE: Adult male F344/N rats were pair housed in a controlled environment under a 12/12 light:dark cycle with ad libitum access to food and water. All animals were cared for using guidelines established by the National Institutes of Health in the Guide for the Care and Use of Laboratory Animals.
<ol>
	<li>Deeply anesthetize rats using 5% sevoflurane.</li>
	<li>Proceed to the following step when the rats are not responsive to noxious stimuli and reflexes are absent.</li>
	<li>Secure the rat in a supine position inside a chemical fume hood.</li>
	<li>Make an incision through the skin along the thoracic midline. Separate the diaphragm and open the chest with scissors. Insert a 20 G &#215; 25 mm needle into the left ventricle.</li>
	<li>Immediately cut the right atrium with scissors. Perfuse 50 mL of 100 mM PBS with 5 mL/min of flow rate. Perfuse 100 mL of 4% paraformaldehyde buffered in 100 mM PBS.</li>
	<li>Remove the entire rat brain right after perfusion.</li>
	<li>Postfix the entire brain for 10 min with 4% paraformaldehyde. 
	NOTE: Do not postfix in 4% paraformaldehyde for more than 30 min, because it will affect the labeling.</li>
	<li>Cut 500 &#181;m thick coronal sections using a rat brain matrix (see Table of Materials). Make the first cut and keep the blade in place. Make a second cut using a second blade and vertically remove the first blade, keeping the tissue on the blade surface.</li>
	<li>Place the brain slices in a 24 well plate with 1 mL of 100 mM PBS in each well. Repeat this process until all slices have been cut.</li>
</ol>
3. Ballistic labeling and visualization of brain sections
<ol>
	<li>Remove the PBS from each targeted well.</li>
	<li>Load the cartridge with a piece of DiI/Tungsten bead tubing and place it into the applicator.</li>
	<li>Put a piece of filter paper between two mesh screens. Connect the applicator to the helium hose. Adjust the output pressure of helium to 90 pounds per square inch (psi).</li>
	<li>Place the applicator vertically by hand on the center of the targeted well at a distance of 1.5 cm between the sample and the mesh screen. Fire the DiI/Tungsten beads tubing. 
	NOTE: Be sure to remove all PBS from the targeted wells before shooting.</li>
	<li>Load the cartridge with the next DiI/Tungsten bead tubing. Continuously fire the DiI/Tungsten beads from the tubing on the remaining slices.</li>
	<li>Fill the 24 well plate with 100 mM of PBS. Wash with 500 &#181;L of fresh 100 mM of PBS 3x. Do not let the slices flip over while washing with PBS.</li>
	<li>Add 500 &#181;L of fresh 100 mM PBS and keep the slices at 4 &#176;C in the dark for 3 h.</li>
	<li>Transfer the brain slices onto glass slides using a fine brush. 
	NOTE: Three brain sections can be transferred onto each glass slide.</li>
	<li>Immediately add 1 mL of antifade mounting medium onto each section. Place a 22 mm x 50 mm coverslip over the brain sections. Dry the glass slides in the dark for 2 days.</li>
	<li>Turn on the confocal microscope system and switch to a 60&#215; objective.</li>
	<li>Set the confocal microscope system to have a magnification of 60&#215; (A/1.4, oil) and a Z-plane interval of 0.15 &#956;m (pinhole size 30 &#956;m, back-projected pinhole radius 167 nm). Use a 543 nm wavelength to acquire images of the neurons of interest.</li>
	<li>Obtain Z-stack images for the targeted neuron type based on brain region boundaries and morphological characteristics of neurons. 
	NOTE: Acquire at least three images fromm each animal.</li>
</ol>
4.&#160;Use of the methodology with cell culture
<ol>
	<li>Isolate primary cortical neurons from F344/N rats at postnatal day one using previously reported methodology14.</li>
	<li>Culture primary cortical neurons in a 35 mm glass bottom dish for one week. Change half of the culture medium with fresh neuron growth medium at third day after isolation. Wash the glass bottom dish 2x with 1 mL of 100 mM PBS.</li>
	<li>Fix with 4% paraformaldehyde for 15 min at room temperature. Repeat steps 3.2&#8211;3.6 to ballistically label the cells.</li>
	<li>Wash with 1 mL of 100 mM PBS 3x. Add 500 &#181;L of fresh 100 mM of PBS and keep it at 4 &#176;C in the dark for 3 h.</li>
	<li>Add 200 &#181;L of antifade mounting medium.</li>
	<li>Obtain Z-stack images for each targeted neuron using the parameters in step 3.10.</li>
</ol>
5.&#160;Neuronal analysis and dendritic spine quantification
<ol>
	<li>Blind neurons using code numbers to prevent experimenter bias.</li>
	<li>Establish selection criteria for neurons based on the brain region of interest. 
	NOTE: Selection criteria for neurons include continuous dendritic staining, low background, no dye clusters inside of the cells, minimal diffusion of the DiI dye into the extracellular space, correct morphology of pyramidal neurons (Figure 1).</li>
	<li>Open neuronal reconstruction software (See the attached Video 1).</li>
	<li>Load an image file by clicking on the &#8220;File Folder&#8221; image in the upper left-hand corner.</li>
	<li>Click &#8220;Soma&#8221; and mark the soma of the neurons on the image.</li>
	<li>Click &#8220;Tree&#8221; and select &#8220;User-Guided&#8221;.</li>
	<li>Trace all dendritic branches of interest. 
	NOTE: For pyramidal neurons, which are characterized by one apical dendrite and multiple basilar dendrites, only the apical dendrite is traced. Make sure all connecting branches are attached to one another.</li>
	<li>Click &#8220;Spine&#8221;.</li>
	<li>Define Detection Parameters and click &#8220;Detect All&#8221;. 
	NOTE: For brain slices, the consistent parameters utilized across brain regions are: 2.0 (Outer Range), 0.3 (Minimum Height), 100% (Detector Sensitivity), and 10 (Minimum Count). For cell culture, it is necessary to increase the Detector Sensitivity and decrease the Minimum Count.</li>
	<li>Classify spines by selecting &#8220;Classify All&#8221;. 
	NOTE: Dendritic spines are classified using an algorithm integral to the neuronal reconstruction software15.</li>
	<li>Save tracing by selecting the Disk Image in the upper left-hand corner.</li>
	<li>Perform neuronal and dendritic spine morphological analyses.
	<ol>
		<li>Open neuronal reconstruction quantitative analysis software (See attached Video 2).</li>
		<li>Load image by clicking &#8220;File&#8221; and &#8220;Open Data File&#8221;.</li>
		<li>Click &#8220;Analysis&#8221; and &#8220;Branched Structure Analysis&#8221; to analyze neuronal morphology and dendritic spine morphology.</li>
		<li>For neuronal morphology, click &#8220;Tree Totals&#8221; and select the box for &#8220;Dendrite Totals&#8221;.</li>
		<li>For dendritic spine morphology, click &#8220;Spines&#8221; and then select the box for &#8220;Spine Details&#8221;.</li>
		<li>Save output as a text (.txt) file by right-clicking the output table and selecting &#8220;Save to File&#8221;.</li>
		<li>Click &#8220;Analyze&#8221; and &#8220;Sholl Analysis&#8221;.</li>
		<li>Set the Starting Radius to 10 &#956;m and the Radius Increment to 10 &#956;m.</li>
		<li>Click the box &#8220;Dendrites&#8221; and click &#8220;Display&#8221;.</li>
		<li>Save output as a text (.txt) file by right-clicking the Output Table and selecting &#8220;Save to File&#8221;.</li>
	</ol>
	</li>
</ol>
6.&#160;Data analysis
<ol>
	<li>Analyze the neuronal morphology (i.e., dendritic branching complexity) data.
	<ol>
		<li>Add the number of dendrites at each branch order and divide by the total number of dendrites. Multiply by 100 to calculate the relative frequencies for the number of dendrites at each branch order.</li>
	</ol>
	</li>
	<li>Analyze the Sholl analysis data to examine neuronal arbor complexity and dendritic spine connectivity.
	<ol>
		<li>Calculate the mean and standard error of the mean for the number of intersections at each radius.</li>
		<li>Sum the number of spines dependent upon spine type (i.e., thin, stubby, mushroom) at each radius and divide by the total number of spines for that spine type. Multiply by 100 to calculate the relative frequencies for the number of spines at each radius.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DiOLISTIC+labeling+of+neurons+from+rodent+and+non-human+primate+brain+slices.">DiOLISTIC labeling of neurons from rodent and non-human primate brain slices.</a> Journal of Visualized Experiments. (41), (2010).</li><li>Spacek, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+the+Golgi+method:+a+time-lapse+study+of+the+early+stages+of+impregnation+in+single+sections.">Dynamics of the Golgi method: a time-lapse study of the early stages of impregnation in single sections.</a> Journal of Neurocytology. 18 (1), 27-38 (1989).</li><li>McLaurin, K. A., Li, H., Booze, R. M., Mactutus, C. 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None of the authors have conflicts of interest to declare.

In this protocol, we describe a versatile labeling technique for neurons from both rat brain and those grown in vitro. Furthermore, we report the methodology for utilizing neuronal reconstruction software and neuronal reconstruction quantitative analysis software to assess neuronal morphology and dendritic spines. The assessment of neuronal morphology and dendritic spines provides an opportunity to determine alterations in dendritic branching complexity, neuronal arbor complexity, dendritic spine morphology, and synaptic...

In 2000, Gan et al. described a rapid labeling technique for individual neurons and glia in the nervous system that combined various lipophilic dyes, allowing for the simultaneous labeling of many brain cells with different colors1,2. More recently, a ballistic labeling technique was described by Seabold et al.3 that introduced fluorescent dyes (Dil) into the neurons of brain slices. A versatile staining technique, ballistic labeling is appreciated for its ability to be utilized in multiple animal species and across a wide range of ages. Furthermore, it can be combined with immunostaining to identify subpopulations of brain cells3. Compared with traditional techniques (e.g., Golgi-Cox silver impregnation, microinjection)4, ballistic labeling affords an opportunity to more clearly distinguish morphological characteristics, including dendritic spines, a feature that is critical for drawing inferences about neuronal complexity and synaptic connectivity5.
Excitatory pyramidal neurons are characterized by a single, large apical dendrite, multiple shorter basal dendrites, and thousands of dendritic spines6. Pyramidal neurons are found in multiple brain regions related to higher order cognitive processing, including the prefrontal cortex (PFC) and hippocampus. In the PFC, pyramidal neurons are observed in layers II/III and layer V, with each exhibiting unique morphology. Specifically, pyramidal neurons in layer II/III of the PFC have a shorter apical dendrite and less branching than pyramidal neurons in layer V6. Within the hippocampus, pyramidal neurons are located in both the CA1 and CA3 regions, with each displaying distinct morphologies. Specifically, pyramidal neurons in the CA1 region exhibit a more distinctive apical dendrite, with branching occurring farther from the soma, relative to the CA3 region6.
Dendritic spines on pyramidal neurons in both the PFC and hippocampus are the primary site of excitatory synapses7. Morphological characteristics of dendritic spines, which are classically characterized into three primary categories (i.e., thin, stubby, or mushroom8), have been related to the size of the excitatory synapse9. Thin spines, characterized by a long, thin neck, small bulbous head, and smaller postsynaptic densities, are more unstable and develop weaker connections. However, mushroom spines, which have a larger dendritic spine head, are recognized for forming stronger synaptic connections, an effect resulting from their larger size. In sharp contrast, stubby spines are devoid of a spine neck, exhibiting an approximately equal head and neck volume ratio8. Within the hippocampus, branched spines may also be observed, whereby the spine has multiple heads that emerge from the same dendritic spine neck10. Therefore, the morphological changes of dendritic spines could reflect functionality and structural capacity. Furthermore, studies have demonstrated that the size and shape of dendritic spines relates to their structural plasticity, leading to the idea that small spines are involved in learning and attention, whereas larger, more stable spines, are involved in long-term processes, including memory11. Additionally, the distribution of dendritic spines along the dendrite may be associated with synaptic connectivity5,12.
Thus, the present methodological paper has three goals: 1) Present our protocol for ballistic labeling, which has been utilized with a success rate (i.e., neurons meeting selection criteria and appropriate for analysis) of 83.3%5,12,13 and across multiple brain regions (i.e., PFC, nucleus accumbens, hippocampus); 2) Demonstrate the generalizability of the technique and its application to neurons grown in vitro; 3) Detail the methodology utilized in neuronal reconstruction software and the inferences that can be drawn from such data.

<table border="0" cellpadding="0" cellspacing="0" style="width:868px;" width="869">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">20Gx25mm PrecisionGlide needle</td>
			<td style="width:263px;">BD</td>
			<td style="width:143px;">305175</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">24-well cell culture plate</td>
			<td>Costar</td>
			<td>3562</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">35 mm Glass Bottom Dishes</td>
			<td>MatTek Corporation</td>
			<td>P35G-1.5-20-C</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Antibiotic-Antimycotic solution</td>
			<td>Cellgro</td>
			<td>30004CI</td>
			<td>100X</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">B-27 supplement</td>
			<td>Life Technologies</td>
			<td>17504-044</td>
			<td>50X</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Barrel liner</td>
			<td>BIO-RAD</td>
			<td>165-2417</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Borax</td>
			<td>Sigma</td>
			<td>B9876</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Boric acid</td>
			<td>Sigma</td>
			<td>B0252</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cartridge holder</td>
			<td>BIO-RAD</td>
			<td>165-2426</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Confocal imaging software</td>
			<td style="width:263px;">Nikon</td>
			<td style="width:143px;">EZ-C1</td>
			<td>version 3.81b</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Confocal microscope</td>
			<td style="width:263px;">Nikon</td>
			<td style="width:143px;">TE-2000E</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cover glass</td>
			<td>VWR</td>
			<td>637-137</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DilC18(3)</td>
			<td>Fisher Scientific</td>
			<td>D282</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMEM/F12 medium</td>
			<td>Life Technologies</td>
			<td>10565-018</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dumont #5 Forceps</td>
			<td>World Precision Instruments</td>
			<td>14095</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dumont #7 Forceps</td>
			<td>World Precision Instruments</td>
			<td>14097</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">F344 rat</td>
			<td style="width:263px;">(Harlan Laboratories, Indianapolis, IN)</td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Glucose</td>
			<td>VWR</td>
			<td>101174Y</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">GlutaMax</td>
			<td>Life Technologies</td>
			<td>35050-061</td>
			<td>100X</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">HBSS</td>
			<td>Sigma</td>
			<td>H4641</td>
			<td>10X</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Helios diffusion screens</td>
			<td>BIO-RAD</td>
			<td>165-2475</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Helios gene gun kit</td>
			<td>BIO-RAD</td>
			<td>165-2411</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Helios gene gun system</td>
			<td>BIO-RAD</td>
			<td>165-2431</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Helium hose assembly</td>
			<td>BIO-RAD</td>
			<td>165-2412</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Iris Forceps</td>
			<td>World Precision Instruments</td>
			<td>15914</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Iris Scissors</td>
			<td>World Precision Instruments</td>
			<td>500216</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Methylene chloride</td>
			<td>Fisher Scientific</td>
			<td>D150-1</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Neurobasal medium</td>
			<td>Life Technologies</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Neurolucida 360 software</td>
			<td style="width:263px;">mbf bioscience</td>
			<td style="width:143px;"></td>
			<td>dendritic spine analysis</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>158127-500G</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Paraformaldehyde</td>
			<td>Sigma</td>
			<td>P6148</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Poly-L-Lysine</td>
			<td>Sigma</td>
			<td>P9155</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Polyvinylpyrrolidone</td>
			<td>Fisher Scientific</td>
			<td>5295</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ProLong Gold antifade reagent</td>
			<td>Fisher Scientific</td>
			<td>P36930</td>
			<td>mounting medium</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rat brain matrix, 300 - 600g, Coronal, 0.5mm</td>
			<td>Ted Pella</td>
			<td>15047</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sevoflurane</td>
			<td>Merritt Veterinary Supply</td>
			<td>347075</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium Bicarbonate</td>
			<td>Life Technologies</td>
			<td>25080</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">SuperFrost Plus Slides</td>
			<td>Fisher Scientific</td>
			<td>12-550-154%</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Syringe kit</td>
			<td>BIO-RAD</td>
			<td>165-2421</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tefzel tubing</td>
			<td>BIO-RAD</td>
			<td>165-2441</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Trypsin-EDTA</td>
			<td>Life Technologies</td>
			<td>15400-054</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tubing cutter</td>
			<td>BIO-RAD</td>
			<td>165-2422</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tubing Prep station</td>
			<td>BIO-RAD</td>
			<td>165-2418</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tungsten M-25 Microcarrier 1.7 &#181;m</td>
			<td>BIO-RAD</td>
			<td>165-2269</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Vannas Scissors</td>
			<td>World Precision Instruments</td>
			<td>500086</td>
			<td></td>
		</tr>
	</tbody>
</table>

ballistic labeling of pyramidal neurons in brain slices and in primary cell culture

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of pyramidal neurons and dendritic spines, a ballistic labeling technique can be utilized. In the present protocol, pyramidal neurons are labeled with DilC18(3) dye and analyzed using neuronal reconstruction software to assess neuronal morphology and dendritic spines. To investigate neuronal structure, dendritic branching analysis and Sholl analysis are performed, allowing researchers to draw inferences about dendritic branching complexity and neuronal arbor complexity, respectively. The evaluation of dendritic spines is conducted using an automatic assisted classification algorithm integral to the reconstruction software, which classifies spines into four categories (i.e., thin, mushroom, stubby, filopodia). Furthermore, an additional three parameters (i.e., length, head diameter, and volume) are also chosen to assess alterations in dendritic spine morphology. To validate the potential of wide application of the ballistic labeling technique, pyramidal neurons from in vitro cell culture were successfully labeled. Overall, the ballistic labeling method is unique and useful for visualizing neurons in different brain regions in rats, which in combination with sophisticated reconstruction software, allows researchers to elucidate the possible mechanisms underlying neurocognitive dysfunction.

In Figure 2A, the typical pyramidal neurons in the hippocampal region in the rat brain sections were identified by ballistic labeling technology, characterized by one large apical dendrite and several smaller basal dendrites around the soma. Figure 2B shows the neuron in the neuronal reconstruction quantitative analysis software after the soma was detected, dendritic branches were traced, and spines were detected. Subsequently, the data were analyzed using neuro...

Watch this Scientific Journal Video about Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture at JoVE.com

Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of ...

ballistic-labeling-pyramidal-neurons-brain-slices-primary-cell

Research

JoVE Journal

Neuroscience

6.5K Views.  University of South Carolina. We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

Video: Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

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

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

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

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

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Primary Culture of Mouse Dopaminergic Neurons

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions such as motor control, motivation, and working memory. Nigrostriatal dopaminergic neurons selectively degenerate in Parkinson&#39;s disease (PD). This progressive neuronal loss is unequivocally associated with the motors symptoms of the pathology (bradykinesia, resting tremor, and muscular rigidity). The main agent responsible of dopaminergic neuron degeneration is still unknown. However, these neurons appear to be extremely vulnerable in diverse conditions. Primary cultures constitute one of the most relevant models to investigate properties and characteristics of dopaminergic neurons. These cultures can be submitted to various stress agents that mimic PD pathology and to neuroprotective compounds in order to stop or slow down neuronal degeneration. The numerous transgenic mouse models of PD that have been generated during the last decade further increased the interest of researchers for dopaminergic neuron cultures. Here, the video protocol focuses on the delicate dissection of embryonic mouse brains. Precise excision of ventral mesencephalon is crucial to obtain neuronal cultures sufficiently rich in dopaminergic cells to allow subsequent studies. This protocol can be realized with embryonic transgenic mice and is suitable for immunofluorescence staining, quantitative PCR, second messenger quantification, or neuronal death/survival assessment.

Dopaminergic neurons play a vital regulatory role in the brain. Their loss is associated with Parkinson's disease. In this video, we show how to generate primary cultures of central dopaminergic neurons from embryonic mouse mesencephalon. Such cultures are useful to study the extreme vulnerability of these neurons to various stresses.

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Isolation, Culture and Long-Term Maintenance of Primary Mesencephalic Dopaminergic Neurons From Embryonic Rodent Brains

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes involved in physiological functions and vulnerability and death of these neurons is imparative to understanding the underlying causes and unraveling the cure for this common neurodegenerative disorder. Primary cultures of mesDA neurons provide a tool for investigation of the molecular, biochemical and electrophysiological properties, in order to understand the development, long-term survival and degeneration of these neurons during the course of disease. Here we present a detailed method for the isolation, culturing and maintenance of midbrain dopaminergic neurons from E12.5 mouse (or E14.5 rat) embryos. Optimized cell culture conditions in this protocol result in presence of axonal and dendritic projections, synaptic connections and other neuronal morphological properties, which make the cultures suitable for study of the physiological, cell biological and molecular characteristics of this neuronal population.

The causes of degeneration of midbrain dopaminergic neurons during Parkinson&#8217;s disease are not fully understood. Cellular culture systems provide an essential tool for study of the neurophysiological properties of these neurons. Here we present an optimized protocol, which can be utilized for in vitro modeling of neurodegeneration.

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...