All procedures were approved by the University Health Network Animal Care Committee and performed in accordance with guidelines and regulations set by the Canadian Council on Animal Care.

1. Stereotactic injection

1. Pair-house adult female Sprague-Dawley rats (250-280 g) in cages with wood bedding and ad lib access to food and water. Maintain the animal colony in a regular 12 h light/dark cycle (lights on 06:30) with constant temperature and humidity.
2. Perform unilateral stereotactic injection of AAV directly to the SNpc on the right side of the brain (right or left side, according to the preferences of each lab) as previously described^8,10. Inject 2 μL of AAV1/2 at a final titer of 3.4 x 10¹² genomic particles/mL.

2. Brain sectioning and immunohistochemistry (IHC)

1. Anaesthetize the rat with 5% isoflurane by placing in an anaesthetizing chamber for 3 min. Other approved methods may be used for this step after appropriate institutional review.
2. Once the rat has reached a surgical plane of deep anesthesia, transfer it to a nose cone firmly affixed to a necropsy table. Secure the rat's fore-paws using tape and use toe pinch-response method to determine the depth of the anesthesia. The animal must be unresponsive before continuing.
3. Make a lateral incision below the sternum and cut through the diaphragm along the entire length of the rib cage to expose the pleural cavity. Lift and clamp the sternum with a hemostat and place above the head.
4. Clamp the heart using forceps and insert a butterfly needle connected to a perfusion pump into the posterior end of the left ventricle. Perfuse rat transcardially with 150 mL of heparinized saline, or until the eyes and skin are clear. Perfusion with 4% paraformaldehyde (PFA), instead of saline, may be preferred to facilitate immunostaining with certain antibodies or thinner brain sectioning.
5. Once perfusion is complete, decapitate with a guillotine and extract the brain to a brain matrix, ventral surface facing up.
6. Using a fresh razor blade, make a cut in the coronal plane 2 mm rostral to the optic chiasm. Slide the blade from side to side to avoid warping the brain while slicing.
7. Immerse the posterior portion of the brain in a pre-labeled vial containing approximately 20 mL of 4% PFA for 48 h of post-fixation at room temperature. The anterior portion of the brain may be flash frozen in 2-methylbutane chilled to -42 °C before storage at -80 °C.
8. After 48 h, transfer the fixed brains to a labeled vial containing 30% sucrose in phosphate buffered saline (PBS) and store at 4 °C until they sink (48-72 h).
9. Prepare a microtome by placing dry ice in the trough of the specimen stage, followed by 100% ethanol. Once the stage has cooled, squeeze optimal cutting temperature (OCT) compound onto the stage until it forms a circle 2 cm in diameter and 0.5 cm thick. Once it has partially frozen, carefully lower the brain onto the mound of OCT, ensuring the striatal cutting surface remains parallel with the stage.
10. Add more dry ice to the stage to help the brain to freeze. Once the brain has turned a cream color, clear the stage of dry ice.
11. Poke a hole into the right side of the brain with a 25G needle to distinguish the right and left hemispheres. Take care not to pass the needle through anatomical structures of interest.
12. Serially cut 40 μm sections in the coronal plane beginning at bregma -3.8 and ending at bregma -6.8.
13. Store six series in labeled tubes with anti-freeze solution (40% PBS, 30% 2-ethoxyethanol, 30% glycerol). Each series should contain 12 brain sections.
14. Select one set of sections for immunohistochemical staining, and wash off anti-freeze solution with 3 x 10 min washes in 0.2% PBS-T.
15. Block for 1 h at RT with gentle nutation in blocking solution (10% normal goat serum (NGS), 2% bovine serum albumin (BSA) in 0.2% PBS-T). Follow this with incubation with rabbit anti-tyrosine hydroxylase (TH) antibody (1:500) and mouse anti-α-syn antibody (1:500) in 2% NGS in 0.2% PBS-T overnight at room temperature.
16. Wash off primary antibody with 3 x 5 min washes in 0.2% PBS-T, followed by 1 h incubation with goat anti-rabbit Alexa Fluor 488 secondary antibody (1:500) and goat anti-mouse Alexa Fluor 555 secondary antibody in 2% NGS in 0.2% PBS-T. Ensure the sections are protected from light and nutating gently.
17. Wash off secondary antibody with 3 x 5 min washes in 0.2% PBS-T and mount the complete set of sections on slides protected from light and dust using a narrow paintbrush. Coverslip with fluorescence mounting medium and seal with clear nail varnish.

3. Confocal microscopy and image acquisition

1. Capture IHC images using software coupled to a confocal microscope at 10x magnification. Open the pinhole to 1.5 AU to capture a wide plane totaling ~1.5 μm and set the focus on the injected side of the brain.
2. On the Acquisition tab, check the Tile Scan imaging option and set the dimensions to 10 x 4.
3. Under the Acquisition Mode panel, set the Zoom to 1.1. This helps to avoid any obvious stitching marks between tile scan images.
4. Set the Frame Size to 1024 x 1024 pixels and the Averaging to 2 to ensure high quality image acquisition.
5. In the Channels panel, set track 1 to Alexa488 and track 2 to Alexa555.
6. Load the slide onto the stage and choose a section with strong TH staining. Click on Live on the Acquisition panel.
7. In the Channels panel, set the Laser Strength and Gain to levels that maximize signal and limit noise from the background. Use the range indicator to ensure that the signals are not overexposed (as indicated by a dark red overlay).
8. Repeat the above step with multiple slides to ensure staining is consistent between slides as the laser strength/gain cannot be adjusted between slides.
9. On the Acquisition tab, check the Positions box.
10. At this point, you are ready to begin imaging. Using the eyepiece, choose the first section showing positive TH staining, set the focus at the point of interest (i.e., SNpc) and then move the stage to the midline of the section. This saves the position in the x, y, and z axes and will image a tile scan capturing the whole section.
11. Repeat the above step for all sections throughout the SNpc giving a complete set of images of the SNpc. If detailed analysis of the uninjected side is required, steps 3.10 to 3.11 should be repeated by setting the focus on the uninjected side.

4. Image analysis and quantitation

1. Separate image files using appropriate software and import image files to automated image analysis software.
2. Define a region of interest by selecting the Pen annotation tool to draw an annotation around the SNpc.
   NOTE: In sections which have a large amount of dopaminergic neuron loss, temporarily increasing the emittance/absorption can help to clearly define the SNpc (Figure 2).
3. Move to the Analysis tab and from the drop-down Analyze menu, select Real-Time Tuning. This opens a separate window on the section image allowing for real-time modification of analysis parameters (Figure 3).
4. Under the Analysis Magnification section, select the appropriate image zoom.
5. Under the Cell Detection section, select nuclear dye as the dye used for TH staining (Alexa Fluor 488).
6. Adjust the Nuclear Contrast Threshold, Minimum Nuclear Intensity, Nuclear Segmentation Aggressiveness, and Nuclear Size settings while carefully watching the Real-Time Tuning window.
   NOTE: Accurate representation of each individual cell as a single cell in the Real-Time Tuning window is vital for accuracy. These settings are on an arbitrary scale depending on the software used, but correct adjustment is needed to allow the software to accurately differentiate between individual cells, and between cells and the background (Figure 3).
7. Repeat this process with a minimum of 10 separate samples to ensure a uniform agreement of what constitutes a cell across different sections.
   NOTE: Additional cell markers (such as α-syn or NeuN) can be identified within the same analysis platform using the Marker 1 or Marker 2 sections on the analysis tab.
8. Once an appropriate number of images have been sampled and Real-Time Tuning has been adjusted accordingly, save the analysis settings in the Settings Actions drop-down menu.
9. Select all images to be analyzed and click on Analyze.
10. Choose the analysis setting you have just saved and in the Region of Analysis window, check the Annotation Layer(s) box. Then, check Layer 1 and click on Analyze.
    NOTE: For a single brain, the analysis typically takes about 5 mins. The completed result will clearly show each item that has been counted as a cell (Figure 4).
11. Once complete, export the summary analysis data for all sections. There is an option to export Object Analysis Data, which will give detailed data, including cell size of each individual cell detected. This dataset could be used to examine changes in cell size in response to a toxin/therapeutic.
12. Add the Total Cells from each section analyzed per animal and the Total Analyzed Area (mm²). Divide the total number of cells by the total area analyzed to calculate the number of cells/mm² in the SNpc for each rat