Analysis of Astrocyte Territory Volume and Tiling in Thick Free-Floating Tissue Sections

All mice were used in accordance with the Institutional Animal Care and Use Committee (IACUC) at the University of North Carolina at Chapel Hill and the Division of Comparative Medicine (IACUC protocol number 21-116.0). Mice of both sexes at postnatal day 21 (P21) were used for these experiments. CD1 mice were obtained commercially (Table of Materials), and MADM9 WT:WT and MADM9 WT:KO mice were described previously⁹.

NOTE: This protocol requires brains with fluorescent protein expression in a sparse population of astrocytes. Fluorescent protein expression can be introduced genetically, virally, or by electroporation. Details of methods to sparsely label astrocytes are described in previously published studies and protocols^{1,7,8,9,10,11,12,13}.

1. Tissue collection and preparation

CAUTION: Paraformaldehyde (PFA) is a hazardous chemical. Perform all steps with PFA in a chemical fume hood.

1. Anesthetize mice with an injectable anesthetic (e.g., Avertin; 0.8 mg/kg) and ensure depth of anesthesia with a toe pinch. Use a peristaltic pump to perform intracardiac perfusion with ice-cold Tris-buffered saline (TBS) + Heparin at a rate of ~3 mL/min until the liver is clear (typically 3-5 min), followed by ice-cold 4% PFA in TBS at a rate of ~3 mL/min for 5 min.
   NOTE: Flow rate and times are optimized for postnatal day 21 (P21) mice. Adjustments to flow rate and perfusion time may be needed for mice of different ages.
2. Following perfusion, use a pair of operating scissors to detach the head and remove the skin to expose the skull. Next, use a pair of micro-dissecting scissors to remove the top of the skull to expose the brain. Use a pair of forceps or a small spatula to remove the brain and transfer it to a tube containing ice-cold 4% PFA and incubate overnight at 4 °C.
   NOTE: Be sure to use a tube with a flat bottom, such as a 7 mL scintillation vial, so that the brain does not become wedged in the bottom of the tube, which could compress the tissue and alter astrocyte volume.
3. The following day, pour out the PFA from the tube. Add 5 mL of 1x TBS to the tube to rinse the brain and remove residual PFA. Pour out the TBS and repeat this process two more times for a total of three rinses.
4. Add 4-5 mL of 30% sucrose in TBS and incubate the brain at 4 °C for 1-2 days, or until the brain sinks to the bottom of the tube.
   NOTE: Brains are ready for freezing once they have sunk but can be stored in sucrose solution at 4 °C for several weeks.
5. Prepare freezing medium in a 50 mL tube by mixing 15 mL of 100% optimal cutting temperature (OCT) compound with 30 mL of 30% sucrose in TBS. Mix on a nutator or an orbital shaker for at least 1 h to ensure the solution is fully mixed. Keep the tube upright for an additional hour to allow any bubbles to rise to the surface.
   NOTE: The freezing medium can be prepared in advance and stored at room temperature for several weeks.
6. Use a serological pipet to add freezing medium to a square embedding mold (Table of Materials). Take care to avoid introducing bubbles into the medium. For a P21 mouse brain, 3 mL is sufficient. Adjust the volume as needed for different ages so that the brain is completely submerged.
7. Add the brain to the medium and use a pair of #5 forceps to remove any brain stem that may be preventing the brain from sitting flat in the mold. Line up the front of the brain with one edge of the mold.
8. Transfer the mold to a flat surface cooled with dry ice. A metal lunch tin filled with dry ice works well for this purpose. Surround the mold with dry ice pellets to ensure slow and even freezing.
9. Once the freezing medium is completely white and solid, store the mold at -80 °C.
   ​NOTE: Brains can be stored at -80 °C for several years.

2. Cyrosectioning

NOTE: This sectioning method is intended to work with many different commercially available cryostats. With the cryostat used here (Table of Materials), the optimal specimen head cutting temperature is -23 °C, with an ambient chamber temperature between -23 °C and -25 °C.

CAUTION: The cryostat blade is extremely sharp. Use caution while manipulating the blade and operating the cryostat.

1. Prepare sectioning medium by mixing 25 mL of 1x TBS and 25 mL of glycerol in a 50 mL tube. It is easiest to add the glycerol by pouring it directly into the tube and using the markers on the tube for measurement. Shake/vortex the tube to mix. This solution can be stored at room temperature for several weeks.
2. Prepare to collect tissue sections by adding 2 mL of sectioning medium per well to a 12-well plate. Label the top and bottom of the plate with sample information. Place the plate, along with other supplies (cryostat blade, razor blade, chucks, and paintbrushes), directly into the cryostat, and allow them to acclimate to temperature for ~5 min.
3. Move the brains into the cryostat chamber and allow them to acclimate to temperature along with the supplies.
4. Remove the frozen tissue block from the mold by cutting two corners of the mold with a razor blade and peeling the mold away from the tissue. Orient the block so that the front of the brain is facing upwards.
5. Add OCT to the chuck such that ~2/3 of the chuck is covered with OCT and immediately place the tissue block on the chuck, keeping the orientation described above. Press the block down into the OCT so that it is flat on the surface of the chuck.
6. Once the OCT is completely frozen and the tissue block is securely in place, clamp the chuck to the specimen head. Insert the cryostat blade and bring the blade close to the specimen.
7. Trim through the brain at 100 µm intervals until just before the brain region of interest is reached. To reduce the amount of OCT that dissolves in the sectioning media, use a razor blade to trim excess freezing medium from the sides of the tissue block.
8. Once the region of interest is reached, begin collecting 100 µm thick sections. Collect sections using either the anti-roll plate or by slowly advancing the cryostat with one hand while using a paintbrush in the other hand to guide the section off the block as it meets the blade. If the cryostat model has a pedal, then use the pedal to advance the cryostat so that both hands can be used to guide the section off the block.
9. Transfer the sections to the 12-well plate using a paintbrush or forceps. Each well can hold at least 10-12 sections. When adding the section to the medium, it is usually sufficient to touch the bottom edge of the section to the sectioning medium and allow the section to melt into the medium without getting the brush or forceps wet. If the brush touches the medium, be sure to dry it with a paper towel or tissue, as an overly wet brush will make it difficult to transfer tissue sections.
10. Secure the plate by wrapping it with foil or paraffin film. Make sure to clearly label it, then store it at -20 °C.
    ​NOTE: For most staining protocols, sections can be stored at -20 °C for at least 1-2 years without substantial loss of signal.

3. 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 3.1-3.9 can be skipped. If skipping immunostaining, perform three 10 min washes in TBS and proceed to step 3.10.

1. 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).
2. 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.
3. Prepare a glass pick by melting the end of a 5.75 in 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.
4. Wash the sections for 10 min each in Wash 1, 2, and 3 wells, followed by incubating the sections 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 one time.
5. 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.
6. 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.
7. 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.
8. 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 bleaching of the secondary antibodies.
9. 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.
10. 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:dH₂O 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.
11. Using a fine paintbrush, transfer the sections one at a time from the Wash 3 well to the Petri dish. This step removes triton and helps the sections to flatten out. Next transfer the sections from the Petri dish into the liquid on the slide.
12. 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.
13. 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.
14. 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 at least 2 h before imaging, or, image the following day.
    ​NOTE: Sections stained with antibodies against GFP and RFP can be stored for up to 2 weeks before imaging, provided they are completely sealed with nail polish.

4. Confocal imaging

NOTE: This protocol gives general image acquisition guidelines that are widely applicable to different confocal microscopes, rather than specific details for a particular confocal and software interface.

1. Use a 10x objective to locate individual cells for imaging, taking note of the specific brain region or sub-region (e.g., layer 5 of the visual cortex) as is relevant for the experiment. Use the focus knob to try and determine whether the entire astrocyte is contained within the section.
2. Switch to a higher magnification oil objective (e.g., 40x, 60x, or 63x) and bring the cell into focus. While looking through the eyepiece, use the focus knob to move from the top to the bottom of the cell.
   1. Visually inspect the cell to ensure that the entire cell is contained within the section. If the cell abruptly goes out of focus and is cut off at the edge of the section, exclude this cell and locate another one.
3. Using the confocal microscope's associated acquisition software, adjust the acquisition parameters to obtain an appropriate signal-to-noise ratio and level of detail (512 x 512 resolution will provide enough detail for territory analysis and will reduce the image acquisition time). Use 2x zoom if using a 40x objective, and no zoom if using a 60x or 63x objective.
4. Set the upper and lower boundaries of the z-stack. To ensure that the entire astrocyte is being captured, the first and last images of the z-stack should not contain any fluorescent labeling of the cell. For adequate sampling, use a step size of 0.5 µm.

5. Image analysis

NOTE: This protocol describes the steps for performing image analysis using commercially available image analysis software (i.e., Imaris; see Table of Materials). Other versions of this software may be used with minor modifications to the workflow. This protocol also requires MATLAB to run the Convex Hull XTension file (Supplemental File).

1. Before beginning analysis, install the Convex Hull XTension file XTSpotsConvexHull.m by pasting the file in the rtmatlab subfolder of the XT folder.
2. Use the Imaris File Converter to convert images to '.ims' format. Open an image file in the image analysis software and view the image in the 3D viewing mode by selecting the 3D View button.
3. Before analysis, inspect the cell to ensure that it meets the inclusion criteria.
   1. Ensure that the entire cell is present within the image (i.e., no part of the cell should be cut out of the image). Rotate the image to inspect the front and back edges.
   2. Ensure that the cell has only one cell body. Click on the Slice button, drag the cursor on the left of the screen to move through the z-stack, and verify that the labeled cell contains only one cell body.
   3. Ensure there is an individual astrocyte that can be distinguished from any neighboring astrocytes that are labeled with the same color.
4. Create a surface
   1. With the 3D View button selected again, create a new surface by clicking on the blue Add new Surfaces icon.
      1. In the Create tab that appears in the bottom left of the software window, make sure that only the Segment only a Region of Interest and Object-Object Statistics boxes are checked for Algorithm Settings, and the Start creation with Slicer view box for Creation Preferences is unchecked. Click on the blue arrow to proceed.
         NOTE: A region of interest may not be needed if there is no additional fluorescent signal in the image outside of the cell of interest. In this case, leave Segment only a Region of Interest unchecked and proceed to step 5.4.3.
   2. Create a region of interest around the cell by entering dimensions into the boxes under Size of the Surface Creation tool, or dragging the edges of the yellow box (Figure 2A). Click on the blue arrow to proceed.
   3. Select the correct Source Channel from the drop-down list for analysis (e.g., Channel 1, or the channel that contains the fluorescent cell fill). The value in the Surfaces Detail box is determined by the software and based on image acquisition parameters. Ensure that this value remains constant for all the samples in the experiment. Click on the blue arrow to proceed.
   4. Adjust the Threshold (Absolute Intensity) by dragging the yellow bar or by inputting values in the box so that the gray surface fills as much of the cell's signal as possible without going beyond the boundary of the cell. A small amount of fluorescent signal will be visible at the edges of the surface (Figure 2B). Click on the blue arrow to proceed.
   5. The last panel of the Surface Creation tool sets the minimum quality value, which is the software's default Lower Threshold value. In the drop-down list, ensure that Number of Voxels Img=1 is selected. Check that only the left power button for Lower Threshold is green (active); the right power button should be red (inactive). The software defaults the Lower Threshold value to 10. Leave this unchanged and click on the green arrow to finish generating the surface.
   6. Carefully inspect the newly generated surface. Delete any surface pieces that are not part of the cell by selecting them, clicking on the pencil Edit tool, and clicking on the Delete option.
   7. Unify the remaining surface pieces by clicking on the funnel Filter icon to select all, clicking on the pencil Edit icon, and clicking on the Unify option (Figure 2C).
      NOTE: With a large image file, the software may occasionally crash during the subsequent steps of this protocol. It is a good idea to save the file at this point.
5. Generate spots close to the surface
   1. Click on the orange Add new Spots icon. With the new Spots (e.g., "Spots 1") selected, in the Create tab, ensure only the Object-Object Statistics box is checked for Algorithm Settings, and the Start creation with Slicer view box for Creation Preferences is unchecked. Click on the blue arrow to proceed.
   2. Select the correct Source Channel from the drop-down list (e.g., Channel 1), and then input an Estimated XY Diameter value of 0.400 µm in the box. Ensure only the Background Subtraction box is selected. Click on the blue arrow to proceed.
      NOTE: 0.400 µm is appropriate for 1024 x 1024 or 512 x 512 images using 63x, 60x, or 40x objective. For other imaging conditions, the diameter must be determined based on imaging conditions and remain constant throughout all analysis. The appropriate value will create enough spots to cover all positive signal but will not add spots on the background signal.
   3. The last panel of the Spots Creation tool sets the minimum quality value, which is the software's default Lower Threshold value; ensure that this value remains unchanged unless there's a noticeable difference in the placement/distribution of the spots (e.g., due to a poorer quality staining). Click on the green arrow to finish creating the spots.
   4. To view the spots better, change the transparency of the surface by selecting the Surface and clicking the multicolor Color tool. Under Material, select the material that makes the surface transparent enough to view spots throughout the cell (the fourth to last material in the list) (Figure 2D,E).
   5. Re-select the Spots, click the Filter icon and select Shortest Distance to Surfaces Surfaces=Surfaces X, where 'Surfaces X' is the surface being analyzed (e.g., Surfaces 1), from the drop-down list. Ensure that both Lower Threshold and Upper Threshold power buttons are green (active).
      1. Input a minimum distance of -1.0 µm (distance of the spots from the inside of the surface) in the Lower Threshold box, and a maximum distance of 0.1 µm (distance from the outside) in the Upper Threshold box. Click on the Duplicate Selection to new Spots button to finish generating spots close to the surface.
         NOTE: The minimum and maximum distance values may vary with different image acquisition parameters, such as objective, zoom, and resolution. These numbers must be determined empirically. The transparent surface helps judge whether the values capture all spots that accurately represent the shape of the surface.
6. Generate convex hull and collect territory measurement
   1. Ensure that the newly created Spots is selected in the upper left panel of the software window (e.g., "Spots 1 Selection ['Shortest Distance…]"). Click on the gear Tools icon, and then the Convex Hull plugin. Click on the plugin only once and wait for the MATLAB window to appear and then disappear (this may take several seconds). A solid hull will appear in the 3D view.
   2. In the upper left panel, select Convex Hull of Spots X Selection ['Shortest Distance…], where 'Spots X Selection' is the newly created spots (e.g., Spots 1 Selection), and then click on the hull to select it (Figure 2F).
   3. Click on the graph Statistica icon, choose the Selection tab, ensure Specific Values is selected from the top drop-down list, and then choose Volume from the bottom drop-down list. Record the volume of the hull. Save the file and proceed to the next image.
7. Measuring territory overlap volume for neighboring cells with different fluorescent labels
   NOTE: This section of the protocol may be used if the samples contain neighboring astrocytes that express distinct fluorescent labels (e.g., Astrocyte 1 expresses GFP only and Astrocyte 2 expresses RFP only). This labeling was achieved using viral or genetic strategies as previously described^9,10 (Figure 3A).
   1. Using the steps detailed in steps 5.4-5.6, create the surface, the spots close to the surface, and the convex hull for each astrocyte (Figure 3B).
   2. With Convex Hull of Spots 1 Selection… selected, click on the pencil Edit icon and click on Mask Selection. In the Mask Channel window, ensure that Channel 1 (or the channel representing astrocyte 1) is selected and make sure that the box next to the duplicate channel before applying mask option is checked. Click on OK to create the channel Masked CH1.
   3. With Convex Hull of Spots 2 Selection… selected, click on the pencil Edit icon and click on Mask Selection. In the Mask Channel window, select Channel 2 (or the channel representing astrocyte 2) from the drop-down menu and make sure that the box next to the duplicate channel before applying mask option is checked. Click on OK to create the channel Masked CH2 (Figure 3C).
   4. Click on the Coloc button at the top of the screen and use the Coloc tool to create a co-localization channel of the two masked channels. Set Channel A a Channel 3 – Masked CH1 and Channel B a Channel 4 – Masked CH2 (Figure 3D).
   5. Drag the yellow histogram bar to the left for both channels. Use the slice view below to observe the co-localization signal, which will be visible in gray. Note that since the fluorescent signals of the two cells are not actually co-localized, the threshold will need to be moved to the left so that false co-localizations begin to appear at points of cell-cell contact.
   6. Click on Build Coloc Channel on the right-hand side to complete the process. Click on the 3D view to return to the standard 3D view. A Colocalization Result channel will now be visible in gray and appears in the display adjustment box.
   7. Create the surface, the spots close to the surface, and the convex hull of the Colocalization Result channel using the steps described in 5.4-5.7 (Figure 3E,F).
   8. Record the volume of the convex hull. This is the territory overlap volume. Divide this number by the territory volume of one of the astrocytes to calculate the percentage of territory overlap. Choose the control or wild-type astrocyte if one of the astrocytes is genetically manipulated with respect to the other.