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

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

1. Dissolve 100 mg of polyvinylpyrrolidone (PVP) with 10 mL of ddH₂O. Vortex the PVP solution lightly.
2. 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.
3. Combine 170 mg of tungsten microcarrier beads with 250 µL of methylene chloride. Vortex the tungsten bead suspension thoroughly.
4. Combine 6 mg of lipophilic DilC18(3) dye with 300 µL of methylene chloride. Vortex the DilC18(3) dye solution thoroughly.
   NOTE: Perform steps 1.3–1.4 in a fume hood.
5. Pipette 250 µL of the tungsten bead suspension onto a glass slide. Wait for the suspension to air dry (~3 min).
6. Add 300 µL 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).
7. Once dried, use a razor blade to split the mixture into two 1.5 mL centrifuge tubes. Fill the tubes with water.
8. 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.
9. Combine the two 1.5 mL of homogenized mixture into a 15 mL conical tube. Sonicate the mixture for another 3 min.
10. 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).
11. Rotate for 1 min on the preparation station. Carefully remove all the water from the tubing using a 10 mL syringe.
12. 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.
13. 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.

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.

1. Deeply anesthetize rats using 5% sevoflurane.
2. Proceed to the following step when the rats are not responsive to noxious stimuli and reflexes are absent.
3. Secure the rat in a supine position inside a chemical fume hood.
4. Make an incision through the skin along the thoracic midline. Separate the diaphragm and open the chest with scissors. Insert a 20 G × 25 mm needle into the left ventricle.
5. 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.
6. Remove the entire rat brain right after perfusion.
7. 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.
8. Cut 500 µm 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.
9. 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.

3. Ballistic labeling and visualization of brain sections

1. Remove the PBS from each targeted well.
2. Load the cartridge with a piece of DiI/Tungsten bead tubing and place it into the applicator.
3. 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).
4. 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.
5. Load the cartridge with the next DiI/Tungsten bead tubing. Continuously fire the DiI/Tungsten beads from the tubing on the remaining slices.
6. Fill the 24 well plate with 100 mM of PBS. Wash with 500 µL of fresh 100 mM of PBS 3x. Do not let the slices flip over while washing with PBS.
7. Add 500 µL of fresh 100 mM PBS and keep the slices at 4 °C in the dark for 3 h.
8. Transfer the brain slices onto glass slides using a fine brush.
   NOTE: Three brain sections can be transferred onto each glass slide.
9. 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.
10. Turn on the confocal microscope system and switch to a 60× objective.
11. Set the confocal microscope system to have a magnification of 60× (A/1.4, oil) and a Z-plane interval of 0.15 μm (pinhole size 30 μm, back-projected pinhole radius 167 nm). Use a 543 nm wavelength to acquire images of the neurons of interest.
12. 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.

4. Use of the methodology with cell culture

1. Isolate primary cortical neurons from F344/N rats at postnatal day one using previously reported methodology¹⁴.
2. 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.
3. Fix with 4% paraformaldehyde for 15 min at room temperature. Repeat steps 3.2–3.6 to ballistically label the cells.
4. Wash with 1 mL of 100 mM PBS 3x. Add 500 µL of fresh 100 mM of PBS and keep it at 4 °C in the dark for 3 h.
5. Add 200 µL of antifade mounting medium.
6. Obtain Z-stack images for each targeted neuron using the parameters in step 3.10.

5. Neuronal analysis and dendritic spine quantification

1. Blind neurons using code numbers to prevent experimenter bias.
2. 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).
3. Open neuronal reconstruction software (See the attached Video 1).
4. Load an image file by clicking on the “File Folder” image in the upper left-hand corner.
5. Click “Soma” and mark the soma of the neurons on the image.
6. Click “Tree” and select “User-Guided”.
7. 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.
8. Click “Spine”.
9. Define Detection Parameters and click “Detect All”.
   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.
10. Classify spines by selecting “Classify All”.
    NOTE: Dendritic spines are classified using an algorithm integral to the neuronal reconstruction software¹⁵.
11. Save tracing by selecting the Disk Image in the upper left-hand corner.
12. Perform neuronal and dendritic spine morphological analyses.
    1. Open neuronal reconstruction quantitative analysis software (See attached Video 2).
    2. Load image by clicking “File” and “Open Data File”.
    3. Click “Analysis” and “Branched Structure Analysis” to analyze neuronal morphology and dendritic spine morphology.
    4. For neuronal morphology, click “Tree Totals” and select the box for “Dendrite Totals”.
    5. For dendritic spine morphology, click “Spines” and then select the box for “Spine Details”.
    6. Save output as a text (.txt) file by right-clicking the output table and selecting “Save to File”.
    7. Click “Analyze” and “Sholl Analysis”.
    8. Set the Starting Radius to 10 μm and the Radius Increment to 10 μm.
    9. Click the box “Dendrites” and click “Display”.
    10. Save output as a text (.txt) file by right-clicking the Output Table and selecting “Save to File”.

6. Data analysis

1. Analyze the neuronal morphology (i.e., dendritic branching complexity) data.
   1. 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.
2. Analyze the Sholl analysis data to examine neuronal arbor complexity and dendritic spine connectivity.
   1. Calculate the mean and standard error of the mean for the number of intersections at each radius.
   2. 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.