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All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Immunostaining
NOTE: Perform all washes and incubations on an orbital platform shaker set to approximately 100 rpm. All steps are performed at room temperature, except the primary antibody incubation, which is performed at 4 °C. Prepare mounting media ahead of time by combining the ingredients in a 50 mL tube and mixing on a nutator overnight. Protect from light and store at 4 °C for up to 2 months. If the endogenous fluorescent signal is sufficient for imaging and analysis without the need for immunostaining, steps 1.1-1.9 can be skipped. If skipping immunostaining, perform three 10 min washes in TBS and proceed to step 1.10.
- Prepare a fresh solution of TBST (0.2% Triton in TBS) by adding 1 mL of 10% Triton-X to a 50 mL tube and filling the tube to 50 mL with 1x TBS. Prepare 2 mL of blocking and antibody solutions (10% goat serum in TBST) for each brain.
NOTE: For best results, make fresh TBST for each new staining experiment and use a high-quality source of 10% aqueous Triton-X (Table of Materials).
- Label a 24-well plate according to the schematic (Figure 1). Place different samples in different rows and different solutions in different columns. Add 1 mL of TBST to the first three columns ('Wash 1', 'Wash 2', and 'Wash 3'). Add 1 mL of blocking solution to the fourth column.
- Prepare a glass pick by melting the end of a 5.75 Pasteur pipet into a small hook using a Bunsen burner. Use this pick to transfer tissue sections from the 12-well plate into the Wash 1 column of the 24-well plate.
NOTE: For best results, screen the sections before beginning staining. To screen sections, transfer the sections one by one to a Petri dish filled with 1x TBS and examine the sections using a fluorescent microscope with a 5x or 10x objective to ensure an adequate number of fluorescently labeled cells are present. Staining four to six sections per well is recommended, though up to eight per well may be stained if necessary.
- Wash the sections for 10 min each in Wash 1, 2, and 3 wells, followed by incubating them for 1 h in the blocking solution. Use the glass pick to transfer the sections from one well to the next. The pick can be used to transfer multiple brain sections at once.
- Prepare 1 mL of primary antibody solution for each sample (e.g., Rabbit RFP 1:2000 or Chicken GFP 1:2000). Add the antibody to the antibody solution, vortex briefly, and then centrifuge for 5 min at ≥ 4,000 x g at 4 °C. Incubate the sections in primary antibody for two to three nights at 4 °C while shaking.
- After primary antibody incubation, aspirate the Day 1 TBST from the wash wells, add 1 mL of new TBST to each wash well, and move the sections into the first wash well. Wash the sections 3x for 10 min each.
NOTE: For aspiration, use a 9-in Pasteur pipet connected with tubing to a vacuum flask. House vacuum lines or an electric vacuum pump can be used as a vacuum source.
- While the sections are being washed, prepare secondary antibody solutions at a concentration of 1:200 by adding antibodies to the antibody solution (e.g., goat anti-rabbit 594 or goat anti-chicken 488). Vortex briefly, and then spin for 5 min at ≥ 4,000 x g at 4 °C.
- Incubate sections in secondary antibody for 3 h at room temperature. Protect the sections from light during this step and all the following steps to mitigate the bleaching of the secondary antibodies.
- After secondary antibody incubation, aspirate the Day 2 TBST from the wash wells, add 1 mL of new TBST to each wash well, and move the sections into the first wash well. Wash the sections 3x in TBST for 10 min each.
- During the final wash, take the mounting media out from 4 °C and allow it to warm to room temperature. Prepare a 2:1 mixture of 1x TBS:distilled water and add this to a Petri dish. Prepare a microscope slide by adding 800 µL of 2:1 TBS:dH2O to the surface.
NOTE: Using non-curing mounting media is strongly recommended. Hardening mounting media can alter the volume of the tissue which may confound the analysis of astrocyte territory volume. A recipe for a simple and inexpensive homemade mounting media is provided in the Table of Materials.
- Using a fine paintbrush, transfer the sections one at a time from the Wash 3 well to the Petri dish. This step removes the triton and helps the sections to flatten out. Next, transfer the sections from the Petri dish into the liquid on the slide.
- Use a paintbrush to help arrange the sections so that they are flat on the slide. Carefully remove excess liquid from the slide with a P1000 pipet, followed by vacuum aspiration.
NOTE: Add a P200 pipet tip to the end of the Pasteur pipet for finer control of vacuum aspiration.
- Once all excess liquid is removed from the slide, use a transfer pipet to immediately add one drop of mounting media to each section and gently lay a coverslip over the slide. Allow mounting media to spread for a few minutes, and then remove any excess mounting media that comes out from under the coverslip by vacuum aspiration.
NOTE: Do not allow the sections any time to dry before adding the mounting media. If the sections begin to dry before mounting media is added, this can impact the tissue volume and impede accurate data collection.
- Seal all four edges of the coverslip with clear nail polish. Let slides dry for 30 min at room temperature, and then store them flat at 4 °C. Wait for at least 2 h before imaging, or image the following day.
NOTE: Sections stained with antibodies against GFP (green fluorescent protein) and RFP (red fluorescent protein) can be stored for up to 2 weeks before imaging, provided they are completely sealed with nail polish.
