Scoring Central Nervous System Inflammation, Demyelination, and Axon Injury in Experimental Autoimmune Encephalomyelitis

All experiments conducted with mice were performed under animal use protocols approved by the Unity Health Toronto Animal Care Committee, following the guidelines set forth by the Canadian Council on Animal Care. Ensure to wear a lab coat, protective gloves, and eyewear throughout the laboratory procedures.

1. Harvesting and fixing the brain and spinal cord

1. Euthanize the mouse according to institutional policies. Lay the mouse prone on a dissecting table and decapitate using surgical scissors (cutting downwards).
2. Using Adson forceps (in non-dominant hand), grasp the skin on the top of the mouse’s head. Then make a 2.5 cm incision in the skin on the top of the head using surgical scissors.
3. Use fingers to push the skin on the head laterally to visualize the underlying skull.
4. Stabilize the mouse’s head by grasping eye sockets with standard Adson forceps.
5. Using fine scissors (dominant hand), make small snips in the skull along the midline from the cervical spine to the olfactory bulbs.
   NOTE: Embed only a few millimeters of scissor tips below the skull at a time to avoid damaging the underlying brain.
6. Use Adson forceps with teeth to reflect the skull laterally to reveal the underlying brain.
7. Hold the head using the non-dominant hand. Holding scissors closed (dominant hand), scoop the spinal cord out from the cervical spine and gently nudge the brain out from the skull, snipping the cranial nerves.
8. Place the brain in a conical tube containing 10 mL of 10% neutral buffered formalin that is labeled with the animal’s ID.
9. Incise the fur along the midline of the mouse’s torso from the neck to the tail.
10. Use fingers to push the skin laterally to visualize the spine.
11. Using surgical scissors, cut downwards through the spine at the level where the femurs attach to the hip.
12. Use surgical scissors to cut the body wall on each side of the spine from the hip to the neck. Trim away any attached organs.
13. Place the spine containing the spinal cord in the same tube of formalin that contains the brain. Allow the brain and spinal column to fix for 5–7 days.
    NOTE: The timing of fixation is important. Some antibodies will not work if tissue is over-fixed. If tissue is under-fixed it is difficult to extrude the spinal cord from the spinal column in step 2.

2. Grossing and processing of the spinal cord and brain

NOTE: The following steps take place in a fume hood. Before starting, prepare 2 x 10 cm clean Petri dishes, an Erlenmeyer flask fitted with a funnel lined with filter paper, two scalpels (one for cutting bone and one for cutting CNS tissues), lens paper, a pencil, embedding cassettes, and specimen jars pre-filled with 10% formalin.

1. Using scissors, cut a small piece of lens paper (the same width but twice the length of the cassette) and place it in one Petri dish.
2. Label a plastic cassette with the specimen ID using a pencil.
3. Pour the tube containing the fixed brain and spine into the funnel. Transfer the spine and brain to the empty Petri dish.
   NOTE: The used formalin will filter through into the Erlenmeyer flask and can be re-used at step 2.13.
4. Grossly divide the brain into six coronal pieces using a scalpel. Make one cut caudal to the cerebellum, one in the middle of the cerebellum, one just rostral to the cerebellum, and two cuts in the remaining rostral brain, creating 3 additional coronal slices of equal thickness.
5. Using forceps, transfer brain specimens onto one half of the lens paper in the Petri dish.
6. Cut the spinal cord into three pieces using the scalpel: the first cut is made at the bottom of the rib cage, and the second cut is made just below the curvature in the cervical spine.     
7. Using the same scalpel, trim the sacral spine piece at the caudal end until the spinal cord can be visualized.
8. Pick up the thoracic spine (non-dominant hand). Hold Adson forceps with teeth tightly closed (dominant hand) and gently push the end of the forceps into the smaller opening of the spinal column using a gentle twisting motion. The cord should emerge from the other end.
   NOTES: Any round-ended instrument that is the size of the spinal cord will work for this purpose. If the spinal cord does not pop out naturally, do not force it. Instead, use fine scissors to clip the bones along the side of the spine and reflect it open to reveal the spinal cord. Alternatively, fix the spinal cord and brain for a few additional days in formalin; however, the same fixation time should be applied to all specimens to avoid introducing variability in antibody staining.
9. Pick up the standard Adson Forceps (dominant hand). Still holding the spinal cord piece with the less-dominant hand, use forceps to gently pull the emerged cord out of the column. Place the spinal cord piece in the Petri dish containing the lens paper.
10. Repeat this process for the lumbar/sacral and cervical column pieces.     
11. Divide the three spinal cord pieces (cervical, thoracic, lumbar/sacral) into smaller cross-sectional pieces using a scalpel. Cut at least 15 segments, each of which should be less than 2 mm thick.
    NOTE: Ensure that the segments are shorter than they are wide, which will make the sections fall more easily in cross-section during the embedding process in step 3.
12. Arrange the spinal cord pieces on the same half of the lens paper containing brain pieces. Fold the lens paper over to sandwich the tissue pieces and place this in the labeled cassette.
    NOTE: The lens paper prevents small pieces of tissue from escaping the cassette during processing.
13. Transfer the cassette into a specimen jar containing recycled or fresh formalin.   
14. Repeat steps 2.1–2.13 for remaining specimens.
15. After 5-7 days of fixation, transfer cassettes from the specimen container into the first formalin bath in the automatic tissue processor (see Table of Materials). Run the tissue processor overnight according to the program described in Supplementary Table 1. Specimens are held in warm paraffin wax until embedding.

3. Embedding and cutting brain and spinal cord sections

1. Transfer cassettes from the processor to the warm holding chamber of the paraffin embedding station (see Table of Materials).
2. Embed spinal cord and brain cross-sections from each mouse in a single paraffin block as follows (Figure 1):
   1. First, pour paraffin wax just to cover the bottom of the mould. Using fine forceps, place brain coronal and spinal cord cross-sectional pieces into the paraffin in the bottom of the mould.
   2. Transfer mould to cooling surface for several seconds to fix brain and spinal cord pieces in place. Move the mould back to the heated surface and fill it to the top with hot paraffin.
   3. Place cassette lid (that is labeled with specimen ID) on top of the mould. Pour more paraffin on top of the cassette lid. Transfer the mould to the cooling station to allow wax to set (cool for 30–60 min).
      NOTE: Embedding spinal cord sections requires practice. To improve success, use shorter lengths of spinal cord (<2 mm) as these are more likely to fall in cross-section. Surgical eye loupes can be worn to help distinguish if spinal cord pieces are in cross-section.
   4. Mount paraffin block on rotary microtome. Trim block until tissues of interest appear in paraffin section.
   5. Cut 5 µM ribbons of sections of each block and transfer them into a 42 °C  water bath.
   6. Collect sections on slides and place slides in a glass slide rack.
   7. Bake sections at 37 °C in a dry oven overnight. Cool slides before proceeding to staining.

4. De-paraffinization and rehydration of sections in preparation for staining

NOTE: Steps are performed in a fume hood. Before starting, prepare baths of solvents. Prepare 5 L of 1x PBS (1 L ddH₂O, 8 g NaCl, 0.2 g KCl, 1.44 g Na₂HPO₄, 0.24 g KH₂PO₄; pH = 7.4) with 0.05% Tween-20 (PBS-T) for all washing steps.

1. De-paraffinize by placing slides in two consecutive baths of xylene or a non-xylene based solvent on a shaker for 5 min each with gentle agitation.
2. Rehydrate tissues by transferring slides through successive baths of descending percentages of ethanol: 2 x 100% ethanol (5 min each), 2 x 95% ethanol (3 min each), and 1 x 70% ethanol (3 min). Hold in 95% ethanol for LFB staining, and rehydrate to 70% ethanol, and hold in PBS-T for immunohistochemistry (IHC).

5. LFB for myelin with H&E

1. Prepare a solution of 0.1% LFB (0.2 g LFB, see Table of Materials, 200 mL 95% ethanol, 0.5 mL glacial acetic acid). Mix and filter into an Erlenmeyer flask.  Store in a dark bottle until use.
2. Transfer sections from 95% ethanol into a glass slide rack that is placed in a staining dish containing LFB. Cover the dish and seal with with paraffin film to prevent evaporation.
3. Incubate sections at 56 °C oven overnight (maximum 16 h).
4. The following morning, transfer slides into a ddH₂O bath and hold.
5. Meanwhile, prepare (1) fresh 0.05% lithium carbonate solution (0.05 g lithium carbonate, 100 mL ddH₂O); (2) Eosin Y solution (add 2 g of eosin salt to 40 mL ddH₂0 and mix until dissolved and then mix with 160 mL of 95% ethanol).
   1. Prepare the following baths: 1 x lithium carbonate, 3 x 70% ethanol, 3 x 95% ethanol, 2 x 100% ethanol, 3 x ddH₂0. Place baths in order of use (see Supplementary Table 2).
6. Follow the steps as outlined in Supplementary Table 2.
7. After the slides are dry, visualize the demyelinating lesions under the microscope.

6. LFB for myelin without H&E

1. Perform this staining procedure for the analysis of myelin.
   NOTE: The procedure is identical to step 5 (see Supplemetary Table 2), but has an abbreviated work-flow. After step 4, continue procedure starting at step 10.

7. Antigen retrieval and peroxidase quenching for immunohistochemical (IHC) stains

NOTE: Before starting, prepare 100 mL of hydrogen peroxide in methanol (1 part 30% hydrogen peroxide solution in 9 parts 100% methanol, in a fume hood). Prepare 1 L of 10 mM citrate buffer with Tween-20 (2.94 g of trisodium citrate, dissolved in 1 L of ddH₂0 in a beaker on stir plate, bring pH to 6.0, and add 500 µl of Tween-20). Prepare PBS-T (see step 4). All washes are done in baths of PBS-T with gentle agitation (on a shaker) unless otherwise indicated.

1. Quench endogenous peroxidase by placing slides in 3% hydrogen peroxide in methanol for 15 min (in a fume hood). Wash slides twice in PBS-T (2 min each).
2. Transfer slides to a metal slide holder and place in 1 L of citrate buffer in a pressure cooker. Seal the lid and add a rubber stopper on the top of the steam escape vent.
3. Cook on high in the microwave until the yellow tab on the pressure cooker pops up, indicating that maximum pressure has been reached. Cook for an additional 5 min at maximum pressure, and then remove the pressure cooker from the microwave using protective gloves.
4. Depressurize by removing the stopper.  Remove the lid and let the slides cool in the citrate buffer for 20 min and then proceed to desired staining method.
   CAUTION: Stand back when releasing the steam as it can cause a scalding injury.

8. CD45 immunohistochemistry

NOTE: This IHC method is used to visualize infiltrating leukocytes. The avidin/biotin blocking steps are combined with the blocking and primary antibody incubation steps.

1. Prepare blocking buffer (2% BSA, 2% rabbit serum in 1x PBS).
2. Transfer slides to glass slide tray and wash twice with PBS-T (2 min each). Prepare avidin/blocking solution (4 drops/mL of avidin in 2% BSA/2% rabbit serum in 1 x PBS, see Table of Materials).
3. Dry off excess PBS-T around tissue using a laboratory tissue paper. Using a hydrophobic pen, draw a circle around the tissue and place slide in the humid chamber.
4. Apply avidin/blocking solution onto each section (400 µL/slide).
5. Cover humid chamber and incubate for 30 min at room temperature.  During this step, prepare anti-CD45 antibody (Supplementary Table 3) in blocking buffer containing biotin (4 drops/mL biotin, 2% BSA/2% rabbit serum in 1x PBS).
6. Tap off blocking solution from slide onto a lint-free laboratory wipe. Dab around tissue using a laboratory wipe to remove excess liquid.
7. Place the slide back into a humid chamber. Add 400 μL of CD45 antibody solution (see Table of Materials) onto the section. Incubate overnight at 4 °C in a covered humid chamber.
8. The next day, drain off primary antibody and wash slides 3 x in PBS-T (5 min each).
9. Dry area around section using a lint-free laboratory and then add 400 μL of secondary antibody (1:200 dilution in blocking buffer) onto each section. Incubate at room temperature for 1 h.
10. Meanwhile, prepare ABC reagent (see Table of Materials) by adding 2 drops of reagent A to 5 mL of 1x PBS and mix. Add 2 drops of reagent B to the same solution and mix (prepare ~30 min before use).
11. Wash the slides in 3 changes of 1x PBS-T (5 min each) and place the slides in a humid chamber.
12. Add 400 μL ABC reagent to sections. Cover humid chamber and incubate for 30 min at room temperature.
13. Wash the slides in 3 changes of 1x PBS-T (5 min each). Meanwhile, prepare an appropriate amount of DAB solution in a foil-covered 15 mL centrifuge tube according to the manufacturer’s instructions (see Table of Materials).
14. Take one slide and focus on a spinal cord section under a microscope. Add 400 μL DAB to the slide and start lab timer.
15. Visualize the section while it is developing and stop timer when leukocytes are brown. Transfer the slide into a ddH₂0 bath to stop the reaction. Hold in water for 5 min. The same development time is used for the remaining slides. 
    CAUTION:  DAB is a carcinogen. Dispose of DAB waste and post-DAB ddH₂O as hazardous waste.
16. Counterstain slides with Mayer’s Hematoxylin for ~4-10 min (see Supplementary Table 2). Rinse slides under running tap water for 10 min.
17. Dehydrate in 95% ethanol (1 x 3 min), followed by absolute 100% ethanol (2 x 3 min each).
18. Moving into a fume hood, transfer the slides into xylene or a xylene-substitute solvent for 5 min. Coverslip with mounting medium, and allow the slides to dry for 1-2 days in the fume hood.
    CAUTION:  If using xylene, use double gloves and forceps to handle slides when cover-slipping, since it is toxic and can dissolve gloves.
19. Clean the slides using xylene and scan using a slide scanner at 20 x magnification.

9. SMI-32 IHC for axonal damage

NOTE: This protocol uses a mouse SMI-32 antibody, which reacts against non-phosphorylated neurofilament heavy, which can accumulate in injured axons²⁵. Since this antibody has been raised in mouse and detects a mouse antigen, it is recommended that a Mouse on Mouse (MOM) kit be utilized. In this procedure, the avidin/biotin blocking step is done as a separate step from the primary antibody incubation. Before starting this protocol, de-paraffinize, rehydrate, quench endogenous peroxidase activity, and conduct antigen retrieval as described in step 4 and step 7.

1. Wash sections twice with 2 x PBS-T (2 min each). Remove extra liquid around sections using a laboratory tissue and draw a circle around tissues using a hydrophobic pen.
2. Add 400 μL blocking buffer (2% (w/v) goat serum in 1x PBS-T) with avidin (4 drops/mL) to the sections. Incubate for 15 min at room temperature.
3. Dip the slides twice in 1x PBS-T. Add 400 μL of blocking buffer with biotin (4 drops/mL) to the slide and incubate for 15 min at room temperature.
4. Wash the slides in a 1 x PBS-T bath for 2 min. Meanwhile, prepare MOM blocking reagent by adding 2 drops of stock solution (see Table of Materials) to 2.5 mL of 1x PBS.
5. Add 400 μL of MOM reagent onto the section. Incubate for 1 h at room temperature.
6. Wash the slides twice in a 1x PBS-T bath for 2 min. Meanwhile, prepare MOM diluent by adding 300 μL of protein concentrate stock solution to 3.75 mL of 1x PBS.
7. Add 400 μL of MOM diluent and incubate for 5 min at room temperature. Meanwhile dilute the SMI-32 antibody (see Table of Materials) in the MOM diluent.
8. Tap off the diluent from slides and add 400 μL of SMI-32 antibody solution onto section. Cover the humid chamber and incubate for 30 min at room temperature.
9. Wash slides twice in a 1x PBS-T bath for 2 min, each with gentle agitation. Meanwhile, dilute MOM anti-mouse IgG working reagent in the diluent  (10 μL of stock in 2.5 mL of diluent).
10. Add 400 μL of anti-mouse IgG working reagent per slide. Incubate for 10 min.
11. Wash slides twice in a 1x PBS-T bath for 2 min each. Continue staining following the steps outlined in CD45 staining protocol (step 8.11 – 8.19). 

10. LFB and H&E scoring for the presence of demyelinating lesions

NOTE: The following is an analysis approach that can be applied to gain rapid insights into the severity of inflammatory demyelination. This analysis is conducted in sections of the cord sampled at different levels (cervical, thoracic, and lumbar, at least 3 sections per/level). Refer to the Allen Brain Atlas for Mouse spinal cord²⁶ to help identify the anatomical level of the spinal cord. This analysis requires TIFF files. If scanned images are in .czi format, follow instructions in Supplementary Table 4 to convert czi files to TIFF files.

1. Open the TIFF image of the section to be analyzed by dragging and dropping the file into the ImageJ. Observe the spinal cord section in 4 quadrants: dorsal, left lateral, right lateral, and anterior (Figure 2A,B).
2. Score for the presence of demyelinating lesions in each quadrant as depicted in Figure 2.
   NOTE: The presence of a lesion results in a score of 1 in that quadrant, for a total of 4 possible points per section. A score of 1 is assigned even if there are multiple lesions within a quadrant.
3. Add up all points for each mouse and divide by the total number of quadrants sampled to obtain the fraction of quadrants with sub-meningeal lesions.

11. Calculating the area fraction of LFB-staining in spinal cord white matter

NOTE: This analysis measures the percent area fraction of spinal cord white matter that is stained with LFB.

1. Open an LFB-stained section saved as a TIFF file by dragging/dropping the file into ImageJ.
2. Click on Image > Type > 8-bit to produce a grey scale image. Click on Image > Adjust > Threshold. Adjust the bottom slider in order that the red overlay captures all of the dark regions (myelin) seen by eye (Figure 2C–E). Click on Apply.
3. Click on Analyze > Tools > ROI Manager to open the ROI Manager.
4. Select the polygon drawing tool on the ImageJ tool bar. This drawing tool allows one to outline a region of spinal cord with the computer mouse.
5. Outline the dorsal region of spinal cord and add the polygon to the ROI Manager by clicking t on the keyboard.
6. Outline the anterior-lateral region of the spinal cord white matter and add the polygon to the ROI manager.
   NOTE: When tracing, exclude large blood vessels, tears in the tissue, artifacts, and areas adjacent to the dorsal horn (Figure 2A,B arrow), which are normally less myelinated. Be as accurate as possible when tracing since including too much white background will skew the results.
7. Click on Analyze > Set Measurements and select Area Fraction.
8. Click on Measure in the ROI Manager. This will give the fraction of the outlined area that is stained with LFB.
9. From the newly produced results box, copy the values into Excel and close the image window. Calculate the average % fraction area stained for the dorsal and ventro-lateral regions. Average these to get a value for that section.
10. Repeat steps 11.1–11.9 for cervical, thoracic, and lumbar sections (N = 3/level/mouse).    
11. Calculate the average percent myelin fraction for all sections for each mouse. Percent demyelination is estimated by subtracting the % area staining from 100.

12. Analysis of the number of CD45⁺ cells and SMI-32⁺ axon ovoids

1. Drag and drop a CD45 or SMI-32 stained TIFF image into ImageJ. Click on Analyze > Set Scale > Global > Ok to set the scale bar across all images. Note that this is done only for the first image.
2. Select the polygon drawing tool and trace around the grey matter using the computer mouse. Click on Analyze > Measure and record the results in Excel as “Grey Matter Area”.
3. With the drawn polygon still on the image, remove the grey matter from the image by clicking on the Delete key (keyboard). This action leaves only the white matter to be analyzed.
4. Outline the entire spinal cord section using the polygon drawing tool. Exclude any regions of tissue that are missing or damaged.
5. Click on Analyze > Measure and record the results as “Total Tissue Area” in the Excel file.
6. Click on Image > Colour > Colour Deconvolution. From the vectors drop-down window select H DAB. Three new windows will appear – keep the brown hued window (DAB channel) and delete other image windows.
7. Click on Image > Adjust > Threshold. Adjust the bottom slider to threshold the image so that the red overlay captures the same amount of brown staining detected by eye. Click on Apply.
8. Click on Process > Binary > Watershed. At this step compare the original image and the binary image to assure that they are in agreement.
9. Click on Analyze > Analyze Particles. Select Show Overlay and modify the settings: Size = 5 – 150 μm², Circularity = 0.4 – 1. Click on OK.
   NOTE: These settings allow for individual cells and small groups of cells that were not split with the Watershed function to be included as opposed to being excluded in the analysis.
10. In the results window, record the last number in the left-most column in Excel, which represents the total particle count. Then, calculate the white matter area (total tissue area – grey matter area in mm²). Express total particle count per white matter area (count/mm²).
11. Repeat steps for each saved RGB image. Once all spinal cord sections have been analyzed for one mouse, average the particle count per mm² tissue for that mouse.      
    NOTE: The workflow used to analyze spinal cord leukocytes can also be applied to brain regions.

13. Measurement of sNF-L using a SIMOA assay

1. Collect 100–200 μL blood from live mice by saphenous bleed using capillary microtainer tubes or via cardiac puncture using a syringe and 25 G needle (terminal procedure). For the latter, transfer blood into a 1.5 mL microcentrifuge tube.
2. Allow the blood to clot at room temperature for 30–60 min.
3. Centrifuge samples at 2660 x g for 5 min at 4 °C and pipette the upper fraction (serum) into a new microcentrifuge tube. Store the serum at -80 °C until ready to use.
4. Preparation: Allow calibrators and controls (NF-light assay kit, see Table of Materials) to warm to room temperature for 1 h prior to use.  Take the enzyme substrate (RGP) out of the refrigerator and place in a 30 °C waterbath for at least 30 min, vortexing every 10 min.
5. Thaw the serum samples on ice. Once thawed, gently vortex samples and centrifuge at 10000 x g for 5 min to pellet any debris.
6. Load the plate: Vortex the calibrators and the controls. Load the calibrators in duplicate, controls in duplicate, and serum samples in duplicate onto the 96-well plate provided with the kit. Serum samples are diluted in a 1:3 ratio with diluent provided in the kit.  Seal the plate.
7. Turn on the SIMOA machine (see Table of Materials) in the order of computer, program, and then machine. Allow the machine to initialize and run Start of Day maintenance.
8. Vortex the magnetic beads for 30 s.
9. Load Reagents tab on the screen, double clicking the rack position where the reagent bottle is to be placed, and then scanning the barcode of the bottle.
10. Place magnetic beads in the shaking position on the rack (positions 1–3).
11. Continue, loading the detector, the SBG reagent, and RGP reagent into the machine.
12. Click setting on the software. Ensure sample mode is on plate. In the setup run tab, name the experiment.
13. Assign wells on the plate for the calibrators as follows: Click the well, then select the assay. Select calibrate A then click on ascending. Highlight wells A1-8 then click replicates (2) to assign the coordinates for the calibrators to be run in duplicate. Run using the neat protocol.
    NOTE: Refer to the certificate of analysis (manufacturer website) to get the concentrations of each calibrator for each particular lot number. Once wells are assigned, it is not possible to navigate away from this screen without losing the work, until the plate is locked in place.
14. Assign the samples following the steps below:
    1. From the bottom right of the screen, click button to assign the samples. Highlight wells where the samples are to be located.
    2. Check off the assay to be run from the list, chose the number of replicates, and specify the onboard dilution of 4x.
    3. With the wells still highlighted, enter a prefix for the sample ID and the starting number and then click on generate to produce sequential IDs for the samples. Repeat steps for control samples.
15. Load plate into the plate holder (A1 to A12).
16. On the screen interface, click on done programming samples and proceed to system resources tab. Check that all reagent containers are full and the waste containers are empty.
17. Click on Run.
18. When the run is complete, check the calibration curve under “Data Reduction” tab by selecting the assay and plate name.
19. Go to “Run History” and use the filters to find the most recent run. Select run and then all results. Click on export, and save the .csv file. Click on  report and then select batch calibration report and select the recent run. Preview the report and export it.
20. Run the end of day maintenance recommended by the manufacturer. Turn off the program, the machine, and then the computer.