Name: ballistic labeling of pyramidal neurons in brain slices and in primary cell culture
Uploaded: 2020-04-02T14:00:00.000+00:00
Duration: 9 min 40 s
Description: Watch this Scientific Journal Video about Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture at JoVE.com

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

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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 ...

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Neuroscience

6.6K Views.  University of South Carolina. Dit protocol maakt het mogelijk om mogelijke morfologische veranderingen in neuronen en dendritische stekels te evalueren die neurochemische en gedragsafwijkingen kunnen onderstrepen. Het is uniek en nuttig voor het visualiseren van neuronen in verschillende hersengebieden bij ratten, die in combinatie met geavanceerde reconstructie software onderzoekers in staat stelt om de mogelijke mechanismen die ten grondslag liggen aan neurocognitieve disfunctie te verduidelijken. Begin met het voorbere...

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

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; ...

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

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

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

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

Rapid Genotyping of Animals Followed by Establishing Primary Cultures of Brain Neurons

High-resolution analysis of the morphology and function of mammalian neurons often requires the genotyping of individual animals followed by the analysis of primary cultures of neurons. We describe a set of procedures for: labeling newborn mice to be genotyped, rapid genotyping, and establishing low-density cultures of brain neurons from these mice. Individual mice are labeled by tattooing, which allows for long-term identification lasting into adulthood. Genotyping by the described protocol is fast and efficient, and allows for automated extraction of nucleic acid with good reliability. This is useful under circumstances where sufficient time for conventional genotyping is not available, e.g., in mice that suffer from neonatal lethality. Primary neuronal cultures are generated at low density, which enables imaging experiments at high spatial resolution. This culture method requires the preparation of glial feeder layers prior to neuronal plating. The protocol is applied in its entirety to a mouse model of the movement disorder DYT1 dystonia (&#916;E-torsinA knock-in mice), and neuronal cultures are prepared from the hippocampus, cerebral cortex and striatum of these mice. This protocol can be applied to mice with other genetic mutations, as well as to animals of other species. Furthermore, individual components of the protocol can be used for isolated sub-projects. Thus this protocol will have wide applications, not only in neuroscience but also in other fields of biological and medical sciences.

We describe procedures for labeling and genotyping newborn mice and generating primary neuronal cultures from them. The genotyping is rapid, efficient and reliable, and allows for automated nucleic-acid extraction. This is especially useful for neonatally lethal mice and their cultures that require prior completion of genotyping.

High-resolution analysis of the morphology and function of mammalian neurons often requires the genotyping of individual animals followed by the analysis ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

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

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

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

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

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

Modified Roller Tube Method for Precisely Localized and Repetitive Intermittent Imaging During Long-term Culture of Brain Slices in an Enclosed System

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of their normal in vivo interactions. Slices obtained from a variety of transgenic mouse lines or use of viral vectors for expression of fluorescently tagged proteins or reporters in wild type brain slices allow for high-resolution imaging by fluorescence microscopy. Although several methods have been developed for imaging brain slices, combining slice culture with the ability to perform repetitive high-resolution imaging of specific cells in live slices over long time periods has posed problems. This is especially true when viral vectors are used for expression of exogenous proteins since this is best done in a closed system to protect users and prevent cross contamination. Simple modifications made to the roller tube brain slice culture method that allow for repetitive high-resolution imaging of slices over many weeks in an enclosed system are reported. Culturing slices on photoetched coverslips permits the use of fiducial marks to rapidly and precisely reposition the stage to image the identical field over time before and after different treatments. Examples are shown for the use of this method combined with specific neuronal staining and expression to observe changes in hippocampal slice architecture, viral-mediated neuronal expression of fluorescent proteins, and the development of cofilin pathology, which was previously observed in the hippocampus of Alzheimer's disease (AD) in response to slice treatment with oligomers of amyloid-&#946; (A&#946;) peptide.

Presented here is a modified roller tube method for culturing and intermittent high-resolution imaging of rodent brain slices over many weeks with precise repositioning on photoetched coverslips. Neuronal viability and slice morphology are well maintained. Applications of this fully enclosed system using viruses for cell-type specific expression are provided.

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of ...

Single Cell Multiplex Reverse Transcription Polymerase Chain Reaction After Patch-clamp

The cerebral cortex is composed of numerous cell types exhibiting various morphological, physiological, and molecular features. This diversity hampers easy identification and characterization of these cell types, prerequisites to study their specific functions. This article describes the multiplex single cell reverse transcription polymerase chain reaction (RT-PCR) protocol, which allows, after patch-clamp recording in slices, to detect simultaneously the expression of tens of genes in a single cell. This simple method can be implemented with morphological characterization and is widely applicable to determine the phenotypic traits of various cell types and their particular cellular environment, such as in the vicinity of blood vessels. The principle of this protocol is to record a cell with the patch-clamp technique, to harvest and reverse transcribe its cytoplasmic content, and to detect qualitatively the expression of a predefined set of genes by multiplex PCR. It requires a careful design of PCR primers and intracellular patch-clamp solution compatible with RT-PCR. To ensure a selective and reliable transcript detection, this technique also requires appropriate controls from cytoplasm harvesting to amplification steps. Although precautions discussed here must be strictly followed, virtually any electrophysiological laboratory can use the multiplex single cell RT-PCR technique.

This protocol describes the critical steps and precautions required to perform single cell multiplex reverse transcription polymerase chain reaction after patch-clamp. This technique is a simple and effective method to analyze the expression profile of a predetermined set of genes from a single cell characterized by patch-clamp recordings.

The cerebral cortex is composed of numerous cell types exhibiting various morphological, physiological, and molecular features. This diversity hampers ...

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 ...

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

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

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

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

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

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

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