Note: The work presented was performed in compliance with recognized international standards, including the International Conference on Harmonization (ICH), the Council for International Organizations of Medical Sciences (CIOMS), and the principles of the Declaration of Helsinki. Use of human tissue was with the approval of University Health Network Research Ethics Board. The human brain samples were collected as a part of the Maritime Brain Tissue Bank. At the time of collection, informed consent was obtained from all patients.

1. Construction of photobleaching apparatus and solutions

1. Prepare stock solutions.
   1. Prepare 1 L of 1x stock Tris-buffered saline (1x TBS) solution (150 mM NaCl, 50 mM Tris-Cl, pH 7.4) by dissolving 8.77 g of NaCl and 6.06 g of Tris base in 800 mL of ddH₂O and adjust the pH to 7.4 using HCl. Bring up the volume to 1 L and autoclave.
   2. Prepare 10% (200x stock) sodium azide by dissolving 1 g of sodium azide in 10 mL of ddH₂O (10%, 200x stock).
2. For 1-3 standard size slides, use a single 100 mm x 100 mm transparent, square petri dish as a slide chamber. For a single slide chamber, add 0.25 mL of 200x sodium azide stock to 50 mL of 1x TBS to make a 0.05% azide-TBS solution. Stack 2-3 slide chambers vertically to process additional slides. Prepare an additional 50 mL of azide-TBS solution for each chamber.
3. Create a scaffold to elevate the slide chamber(s) such that a lamp head can fit underneath.
   1. For a 100 mm x 100 mm slide chamber, cut openings in the bottom and sides of a 100 mm x 100 mm x 30 mm plastic food container and invert the container. Ensure the side openings are large enough to fit an LED light source and ensure the bottom opening is large enough such that light from the light source reaches the sample chamber without impediment.
   2. Apply electrical tape to the scaffold to increase the grip between the scaffold and the sample chamber/benchtop. Use any alternative materials to construct the scaffold so long as it securely elevates the slide chamber without impeding the light from reaching the sample.
4. Remove any diffusers or opaque plastic from the desk lamp that may impede the LED light from directly reaching the sample (if possible) and orient the LED array upwards. Place the scaffold and slide chamber(s) above the LED array. Use a lamp with a flexible neck for easy manipulation.
5. Construct a reflective dome cover for the apparatus by lining the inside of a box large enough to cover the slide chamber and scaffold with aluminum foil. Use a 1 mL pipette tip box for a single chamber or a 150 mm x 150 mm x 150 mm cardboard box for multiple, vertically stacked chambers.

2. Photobleaching pre-treatment of tissue sections

NOTE: Tissue section preparation may vary depending on the source of tissue and fixation and embedding methods used. Here, brain tissue (orbitofrontal gyri) from a case of FTLD-T was fixed for ~2 days in formalin, run through a sucrose gradient, embedded in OCT, and cut to 10 µm thick sections using a cryostat.

1. In a 4 ºC cold room, cold cabinet, or refrigerator, orient the lamp under the scaffold and place the sample chamber on the scaffold. Pour 50 mL of azide-TBS solution into the sample chamber.
2. Submerge tissue sections mounted on standard glass microscope slides into the slide chamber containing azide-TBS using clean forceps. For multiple slides, ensure that the slides are placed in the chamber on a single layer.
3. Cover the apparatus with the reflective dome, turn on the LED lamp, and incubate for 48 h at 4 ºC.

3. Immunofluorescence

1. To stain the tissue for phosphorylated tau using DAPI counterstain and Alexa 488- and Texas Red-conjugated secondary antibodies, first prepare solutions for antigen retrieval, permeabilization, blocking, and primary antibody binding.
   1. Prepare 500 mL of antigen retrieval buffer (10 mM citric acid, 2 mM ethylenediaminetetraacetic acid, 0.05% Tween 20; pH 6.2) by dissolving 0.92 g of citric acid and 0.37 g of ethylenediaminetetraacetic acid (EDTA) in 500 mL of ddH₂O. Adjust the pH to 6.2 with NaOH and add 0.25 mL of Tween 20.
   2. Prepare 500 mL of 0.025% Triton X-100 in TBS solution (TBS-Triton) by adding 0.125 mL of Triton X-100 to 500 mL 1x TBS.
   3. Prepare a 1% bovine serum albumin (BSA) solution in TBS (BSA-TBS buffer) by dissolving 0.1 g BSA in 10 mL TBS.
   4. Prepare a blocking solution by adding 0.2 mL of normal goat serum to 1.8 mL of 1% BSA/TBS.
   5. For each slide, prepare 150 µL of primary antibody solution (1:100 dilution) by pipetting 1.5 µL of anti-phospho-PHF-tau pSer202 + Thr205 (AT8) antibody into 148.5 µL of 1% BSA-TBS buffer and leave on ice.
2. To perform antigen retrieval, submerge the photobleached slides vertically in a slide collector containing 25 mL of antigen retrieval buffer. Secure the collector with tape and/or strings such that the collector does not fall into the water bath. Heat the collector in a water bath at 90 ºC for 30 min and allow the collector to cool to room temperature for 30 min before removing the slides. Do not remove the slides immediately as it will cause the sections to dry out.
3. Transfer the slides from the antigen retrieval collector into a staining jar filled with 30 mL of TBS-Triton and wash the sections for 5 min on an orbital shaker with gentle shaking. Repeat the wash once with fresh TBS-Triton. Wick away excess buffer with a lint-free tissue and outline the tissue with a hydrophobic pen. Take care not to let the slides dry out.
   1. For each slide, block the tissue by pipetting 200 µL of blocking solution onto the tissue and place the slide in a humidified chamber. Construct the chamber by placing a slide rack inside a pipette tip box containing a wet paper towel. Incubate at room temperature for 2 h on a level surface. Ensure that the blocking solution fully covers the tissue.
4. Remove the blocking solution by aspiration and pipette 100-150 µL of primary antibody solution onto the tissue. Ensure sufficient volume of antibody is present and that the section is on a level surface to avoid pooling of antibody solution to one side. Incubate at 4 ºC overnight in a humidified chamber.
5. Prepare the secondary antibody mixture and the DAPI nuclear counterstain.
   1. For each slide, prepare 150 µL of secondary antibody mixture (1:100 dilutions) by adding 1.5 µL of goat anti-mouse Alexa 488 and 1.5 µL of goat anti-mouse Texas Red to 147 µL of BSA-TBS and leave on ice.
   2. Prepare 0.1 µg/mL DAPI counterstain by serial dilution. Mix the stock solution thoroughly and dilute 1 µL of stock 5 mg/mL DAPI in 999 of TBS to make 1 mL of 5 µg/mL solution. For each slide, dilute 3 µL of 5 µg/mL solution with 147 µL of TBS to a final concentration of 0.1 µg/mL.
      CAUTION: DAPI is a known mutagen and should be handled with care.
6. Remove the primary antibody by aspiration. Submerge the slides in a glass staining jar containing 30 mL of TBS-Triton and wash for 5 min with gentle mixing on an orbital shaker. Repeat the wash step with fresh TBS-Triton. Wick away excess TBS-Triton and pipette 100 to 150 µL of secondary antibody mixture to each slide.
   1. Ensure the tissue is fully covered by the antibody mixture. Incubate for 2 h at room temperature in the humidified chamber in the dark.
7. Remove the secondary antibody mixture by aspiration and transfer the slide into a glass staining jar containing 30 mL of TBS (no Triton). Wash for 5 min with gentle mixing on an orbital shaker. Repeat the wash step with fresh TBS. Apply 100 to 150 µL of 0.1 µg/mL DAPI counterstain to each slide and incubate for 10 min at room temperature in the dark.
8. Transfer the slides to a glass staining jar containing TBS and wash 3 times with gentle mixing for 5 min each, using fresh TBS for each wash. Wick away excess buffer.
9. Apply 3 drops of aqueous mounting medium to the tissue. Using forceps, gently lower a clean glass coverslip onto the tissue, starting with one edge and slowly lowering the other edge to avoid trapping of air bubbles. Take care not to dislodge the coverslip if imaging immediately. Otherwise, allow the mounting medium to dry before storing at 4 ºC in the dark.

4. Fluorescence microscopy

1. Turn on the fluorescence lamp, the microscope and the computer and allow the lamp to warm up for 15 min. Place the stained tissue slides in the fluorescence microscope stage. Use the bright field image to locate the tissue at 10x magnification.
2. Apply a drop of ddH₂O to the coverslip surface and use a 20x water immersion objective lens (NA=1.0). Select the 4-line average plane scan setting. Set the pinhole size to 1 Airy unit that gives an optical slice of ~3 microns. Select the laser excitation and emission wavelengths for each fluorophore in separate tracks for best signal.
   NOTE: Alexa 488: λ_ex = 488 nm (argon laser) λ_em = 493-570 nm; Texas Red: λ_ex = 561 nm (DPSS 561 nm laser), λ_em = 601-635 nm; DAPI: λ_ex = 405 nm (Diode 405 laser), λ_em = 410-507 nm.
   1. Adjust the laser power and gain settings to optimize the signal intensity for each track. Collect the composite image and save. Use the same laser settings to compare fluorescence intensities in a different slide.
3. For visualization of fluorescence intensity in each channel, install the RGB profile tools macro for ImageJ¹⁰. Save the macro from the webpage as a text file (https://imagej.nih.gov/ij/macros/tools/RGBProfilesTool.txt). From the ImageJ menu, select Plugins -> Macros -> Install; select the text file to install the RGB profiles tool.
   1. Open the confocal image file in ImageJ and convert the composite images from the 3 stacks to RGB by performing the following: Image -> Color -> Channel Tool. Select "Composite" from the dropdown menu and check all three channels. Then, select Image -> Type -> RGB color.
   2.  Select the RGB profile tools icon and draw a line across the section in the image to be profiled. Save the intensity data as a spreadsheet for plotting.