Immunofluorescence Staining Using IBA1 and TMEM119 for Microglial Density, Morphology and Peripheral Myeloid Cell Infiltration Analysis in Mouse Brain

All experimental procedures were performed in agreement with the guidelines of the Institutional Animal Ethics committees, in conformity with the Canadian Council on Animal Care and the Animal Care Committee of Université Laval.

1. Immunostaining

1. Select three mouse brain sections containing the region of interest (ROI) (i.e., the hippocampus) with the help of a brain atlas. Place the sections in a plastic multi-well plate and cover them with 350 µL of phosphate-buffered saline (PBS) (Table 1).
   NOTE: For optimal results, the brains should be perfused with 4% paraformaldehyde and cut to a thickness of 50 µm with a vibratome. For a 24 multi-well plate, each well can hold up to six sections. The recommended volume of solution for each well is 350 µL (for up to three sections) and 500 µL for wells containing six sections. For a higher number of sections, it is recommended to use a 12 multi-well plate. Make sure that the selected volume of solution for each well completely covers the tissue and allows the sections to float. The recommended volumes apply for every solution used in the rest of the protocol.
2. Wash the samples by covering them with 350 µL of PBS and let them rest by placing the multi-well plate on top of a multipurpose shaker at room temperature (RT). Remove the PBS after 5 min and replace it 5x with fresh PBS.
   NOTE: To remove the solutions, a transfer pipette is recommended. When pouring in any solution, make sure to place the tip of pipette against the well wall to protect tissue integrity. Also make sure to use a new pipette for each new solution.
3. Remove PBS and add 350 µL of 10 mM sodium citrate buffer with pH = 6.0 (Table 1).
4. Seal the multi-well plate with paraffin film and let it float on a previously preheated water bath for 40 min at 70 °C.
5. Let the multi-well plate cool down for approximately 15 min.
6. Remove the sodium citrate buffer and wash the sections in PBS as done in step 1.2.
7. Remove PBS and add 350 µL of freshly made 0.1% NaBH₄ (Table 1) and let incubate for 30 min at RT.
8. Remove the solution of 0.1% NaBH₄ and wash the sections in PBS as done in step 1.2.
9. Remove PBS and add blocking buffer (Table 1) for 1 h at RT on top of a multipurpose shaker.
   NOTE: Make sure to prepare doubled volumes of blocking buffer, as the same solution will be used in the next step.
10. Remove the blocking buffer and replace by blocking buffer containing the mixture of primary antibodies (1:150 mouse IBA1 + 1:300 TMEM119). Seal the plate with paraffin film and let it incubate overnight at 4 °C.
11. The next day, warm samples at RT for approximately 15 min.
12. Wash the sections 5x for 5 min each in PBS with triton (PBST) (Table 1).
13. Remove PBST and add blocking buffer containing the mixture of secondary antibodies (1:300 donkey anti-mouse Alexa 488 for IBA1; 1:300 goat anti-rabbit Alexa 568 for TMEM119) for 1.5 h at RT. Starting from this point onward, protect the samples from light.
14. Remove blocking buffer and wash the sections 5x as done in step 1.2, except this time with PBST.
15. Remove the PBST and add 4′,6-diamidino-2-phenylindole (DAPI) [1:20000] for 5 min at RT.
16. Remove DAPI and wash the sections 3x for 5 min each in phosphate buffer (PB).
17. Mount the sections on a microscope slide. Let them dry while protected from light.
18. When dried, add some drops of mounting fluorescence medium and cover with a coverslip, avoiding bubble formation.
    NOTE: Store the slides while protected from light, inside a histological slide box, at 4 °C. The samples can be preserved for several months.

2. Imaging for density and distribution analysis

1. With the help of a widefield epifluorescence microscope, use a low magnification and the DAPI channel to locate the ROI (i.e., the CA1 region of the hippocampus).
2. Acquire images at 20x, using a numerical aperture (NA) of 0.5, with the DAPI, 488, and 568 channels and filters, at a resolution of 0.3 µm/pixel. Capture a mosaic picture covering the ROI. Alternatively, take individual pictures that will be stitched into a larger image.
   NOTE: A mosaic image is a super image constituted by smaller images. Mosaic images are usually used to overcome the limited area of the field-of-view of high magnifications. Some software includes a mosaic function; nevertheless, images can also be manually stitched together with other photo editing software by stitching the individual images into one. Remember to add the scale information to the file. For this type of analysis, it is recommended to have at least 300 microglial cells imaged per ROI/animal (corresponding to approximately 10−15 pictures for the hippocampus, for example), with a minimum of five animals per experimental condition. Figure 1A−C shows the images of colabelled microglia.
3. Save the image as a TIFF file.

3. Imaging for morphology analysis

1. Using a confocal or structured illumination microscope, use the DAPI channel to locate the ROI at low magnification.
2. Using a 40x objective (i.e., NA 1.4 oil), locate an IBA1+/TMEM119+ cell inside the ROI. While live imaging, move in the Z-axis. As soon as the signal of the randomly selected microglia disappears, set this Z-level as the beginning of the Z-stack. Move along the Z-axis in the opposite direction until the signal of the microglia disappears and set that point as the end of the Z-stack.
   NOTE: Figure 2A−C shows images of IBA1+/TMEM119+ microglia.
3. Create a Z-stack in all three channels (DAPI, 488, 568) using a 0.33 µm Z-interval and pixel size of 0.15 µm/pixel. Add the scale information to the file.
   NOTE: The recommended Z-interval depends on the resolving power of the objective (e.g., for a 40x objective such as NA 1.4 oil, it is 0.33 µm). For morphology analysis, it is recommended to have at least 20 cells per animal with a minimum of five animals per experimental condition.
4. Save the file as a TIFF file.

4. Density and distribution analysis

1. Open FIJI/ImageJ with the nearest neighbor distance (NND) plugin installed. Open the 20x image.
   NOTE: Use a search engine with the keyword “Nearest Neighbor Distances Calculation with ImageJ” to find the installation instructions. The plugin Author is Yuxiong Mao.
2. To set the scale manually based on a scale imprinted on the image, select the straight line tool (Figure 3E), place the cursor on the edge of the scale, and, while pressing the shift key, draw a line as close as possible to the scale on the image (Figure 3I), select Analyze | Set scale, then enter the correct information (Figure 3J).
   NOTE: The scale can sometimes be contained in the metadata of the file and set automatically.
3. Select Image | Color | Make composite to create a composite image of all channels.
   NOTE: During image acquisition, FIJI/ImageJ will automatically create a composite in the RGB format.
4. On the menu bar, select Analyze | Set measurements. Check Area, Centroid, and Perimeter. On the tab Redirect to, click and select the opened file (Figure 3K).
5. Go to Image | Color | Channel tool to open the channel tool.
   NOTE: This menu will allow a specific color to be disabled. The DAPI channel can be useful to identify ROI and to confirm cells. It can be deactivated to make counting easier.
6. Draw a rough perimeter of the ROI with the freehand selection tool (Figure 3D).
7. Enable the selection brush tool by double-clicking the oval tool on the tool bar and make sure that the Enable selection brush box is checked (Figure 3G). This tool will be used to delineate the ROI more precisely. Select an appropriate brush size between 200−400.
8. Using the selection brush, adjust the perimeter to best fit the ROI. Press T on the keyboard to add to the ROI manager (Figure 3L).
9. Select Analyze | Measure or press the M key, and a results window will pop up. Copy and paste the results on a datasheet, then save the information regarding the area (i.e., the area of the ROI; Figure 3R).
10. After copying the area of the ROI, erase the information from the results window by clicking on it and pressing the Backspace key.
11. Go to the ROI manager window (Figure 3L), right-click the ROI trace, change the name to match the image’s name, then save.
12. Double-click the brush tool at the tool bar. Select the black color and a brush size of 10. Make sure that the option Paint of overlay is unchecked (Figure 3H).
13. In the TMEM119 channel, carefully place a black dot on the center the soma for each TMEM119+ microglia. Place a white dot on the center of the cells that are not positive for TMEM119 (to mark infiltrating macrophages). Repeat the same procedure for all cells contained in the ROI.
    NOTE: It is important that all dots (black and white) are located in the same channel. The identity of the channel can be verified (red, blue, or green) by looking at the color of the image window labels.
14. Select Image | Color | Split channel. A window for each channel will appear. Then, identify the channel that has the dot annotations and close the other two windows.
15. Redirect the new split channel image. Go to Analyze | Set measurements. On the tab Redirect to, click and select the split channel image (Figure 3K).
16. Select Image | Typ | 8-bit. Go to Image | Adjust and select Threshold (Figure 3O). To adjust the threshold, slide the button of the second bar, all the way to the left (threshold value = 0) in both bars.
    NOTE: This will leave only the black dots on the image, appearing white.
17. Select the ROI in the ROI manager window. Select Analyze | Analyze particle (Figure 3N). On Size (inchˆ2): write 1−20. Keep the pixel unit unchecked, check Display, summarize and add to manager, and press Ok. The summary window will pop up and give the number of points (Figure 3P). Copy and paste the information to the datasheet.
18. Select Plugins | NND. The NND window will pop up (Figure 3Q). Copy/paste all the information to the datasheet. Each number represents the distance each microglia has to the nearest neighboring microglia.
19. Go back to the threshold window and slide the first bar all the way to the right (threshold value = 255 in both bars), which will leave all the white dots visible, appearing white (Figure 3M).
20. Select Analyze | Analyze particle. The summary window that provides the number of points will pop up (Figure 3P). Copy and paste the information to the datasheet.
21. Go to the ROI manager select all the points, right-click, and save with the image’s name. This will allow saving of all the points in a zip file (Figure 3L). Select File | Save as, and save the file with a name that allows identification of the analyzed image.
22. Obtain the density of microglia (for each image) by dividing the number of IBA1+/TMEM119+ double-positive cells by the area of the ROI.
    NOTE: The values for each picture can be averaged for each animal. The data can then be presented as mean ± standard error of the mean (SEM) of all the animals.
23. Determine the NND by obtaining an average per picture of the NND values of all TMEM119+ cells.
    NOTE: The data can then be presented as mean ± SEM of all the animals.
24. Calculate the spacing index using the formula: NND² x density.
    NOTE: The data can then be presented as mean ± SEM of all the animals. The units for this measurement will be arbitrary units.
25. Quantify microglial clusters by identifying cells that have an NND under 12 µm.
    NOTE: Here, 12 µm is selected, as it is the approximate distance between two directly juxtaposing microglial cells touching each another with arborizations. If there are more than three microglia that meet this condition, return to the image and verify whether these cells are part of one or multiple clusters.
26. After confirming the number of clusters, write the number of clusters in the datasheet.
    NOTE: The number of clusters can be divided by the ROI area to obtain the density of cells/mm² for each animal. The data can then be presented as mean ± SEM of all the animals.
27. To determine the percentage of peripheral myeloid cell infiltration, calculate the % of IBA1+/TMEM119- cells over the total number of myeloid cells (TMEM119+/IBA1+ + TMEM119-/IBA1+) for each animal.
    NOTE: The data can then be presented as mean ± SEM of all the animals.

5. Morphology analysis

1. Open FIJI/ImageJ.
2. Open the 40x image using Image J or FIJI. Select A popup window will appear asking if the images should be opened in a stack. Click OK. Next, select Image | Stacks | Z project to open the Z-Projection window. Include all slices, from the first through last slice. Ensure that the Max Intensity is selected under Projection Type, and click OK
3. Click on the new window with the Z project. Select Image | Colors | Split channels. Conduct the traces on the images of the IBA1 channel.
   NOTE: The other channels (TMEM119 and DAPI) can be kept open and consulted as needed during the microglial morphology analysis.
4. On the menu bar, select Analyze | Set measurements. Check the Area, Centroid, and Perimeter. On the tab Redirect to, select the opened file (Figure 3K).
5. Set the scale as described in steps 4.2.
6. To measure the soma size in the IBA1 channel, draw a rough perimeter of the soma with the freehand selection tool (Figure 3D).
7. Enable the selection brush tool by double-clicking the oval tool on the tool bar, followed by checking Enable the selection brush box (Figure 3G). Select a selection brush size between 10−20 (Figure 3B).
8. Using the selection brush, adjust the trace to best fit the soma. Zooming in will enable precision during this step (Figure 2I).
9. Press the T key to add the soma trace to the ROI manager (Figure 3L).
10. Select Analyze | Measure or press the M key. A results window will pop up. Copy and paste the results on a datasheet (Figure 3R).
11. To save the information regarding the soma area, go to the ROI manager window, right-click on the ROI, change the name to match the image’s name, specify that the trace is for soma, then save the file.
12. To measure arborization area in the IBA1 channel, click on a microglial process extremity with the polygon selection tool, which will start the polygon shape (Figure 3C).
13. Following the tips of the microglial processes, go around the microglia by clicking at the tips of each process extremity to form a polygon that best represents the area covered by the microglial arborizations (Figure 2D−H).
    NOTE: Make sure that the polygon connects all the microglial process extremities. The lines forming the polygon should never intersect. When clicking around a microglial process tip, be careful to avoid cutting off any part of the process. It is sometimes useful to add extra points to go around a process. The number of points forming the polygon is not directly linked to the number of distal processes and thus is not relevant for the study.
14. To close the polygon, click on the starting point of the polygon.
15. Press the T key to add the trace to the ROI manager (Figure 3L). Select Analyze | Measure or press the M key. A results window will pop up. Copy and paste the results on a datasheet (Figure 3R).
16. To save the information regarding the arborization area, go to the ROI manager window, right-click the ROI, change the name to match the image’s name, specify for arborization, then save the file.
17. Determine the soma area by averaging all soma areas for each animal.
    NOTE: The data can be presented as mean ± SEM of all the animals.
18. Determine arborization area by averaging all the arborization areas for each animal.
    NOTE: The data can be presented as mean ± SEM of all the animals.
19. Calculate the morphology index by using the formula soma area/arborization area for each microglial cell and average per animal.
    NOTE: The data can be presented as mean ± SEM of all the animals.