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1.Euthanize animals according to IACUC protocol.
2.Harvest the mineralized tissues of interest, remove unwanted soft tissues, and trim samples into manageable units using a handheld or electric saw (Fig. 2).
3.Place samples in 10% neutral buffered formalin at 4°C until properly fixed. Fix samples for 3‒5 days.
4.Transfer specimens from formalin to sucrose/PVP solution for 7 days at 4°C.
5.Embed samples in Tissue-PlusTM O.C.T. Compound (cryo embedding medium), or other comparable polyvinyl alcohol/polyethylene glycol-based embedding mediums. Partially fill the cryomold with cryo embedding medium, and carefully place the intended cut plane parallel with bottom of mold.
6.Place the cryo mold on a base of dry ice to freeze the bottom of the sample, add more cryo embedding medium to fully cover the sample, and continue freezing on dry ice. Store samples at −20°C until use.
7.Remove the samples from the freezer, and place in the cryostat with a temperature between −20°C and −25°C.
8.Remove the sample from the cryomold, and adhere to the specimen disc with cryo embedding medium. Wait at least 5 minutes for the embedding medium to freeze, ‘gluing’ the sample to the disc.
9.Load the specimen disc into the specimen head of the cryostat (Fig. 3A). Insert a disposable ceramic-coated blade into the cryostat. Adjust the specimen head so that the blade evenly cuts the sample block.
10.Trim the block using the manual cutting wheel until you reach the region of interest. Brush off the initial sections to create a smooth, clean block face.
11.Set the section thickness to 18 µm.
12.Cut a piece of cryofilm slightly bigger than the sample size. Keep cryofilm chilled in the cryostat.
13.Using chilled forceps, grab the silver/gold tab on the cryofilm and peel off the non-adherent backing. Place the sticky side of the cryofilm over the sample (Fig. 3B). Use a roller or paintbrush to apply pressure to the block face, further adhering the tape to the sample.
14.Use forceps to grip the non-adherent gold tab of the cryofilm in one hand and use the manual cutting wheel to create an 18 µm section.
15.Place the collected section on a plastic slide with the gold tab facing up (tissue side up) (Fig. 3C).
16.Repeat to acquire as many sections as desired.
17.Store sections at 4°C, −20°C, or −80°C depending on length of storage required. At 4°C, store for no longer than a month.
18.Remove a section from a plastic slide with forceps, and using scissors, carefully trim off the excess tape around the tissue. At minimum, trim off the gold/silver tab.
19.Pipette a drop of chitosan adhesive for each section onto a glass slide. Drop size varies (50‒100 µL/section) based on section size.
20.Gradually lay the trimmed section TAPE SIDE DOWN onto the chitosan adhesive. The adhesive should cover the entire bottom of the sample to avoid bubbles between the slide and section.
21.Tilt the slide up on one of its long edges to allow excess chitosan adhesive to drain downward.
22.Place slides in this orientation on top of a paper towel in a slide box.
23.Place slide box with the lid propped open in the refrigerator for 2 days to allow the chitosan adhesive to fully dry.
24.Prepare glycerol-based mounting medium with TO-PROTM-3 Iodide nuclear counterstain (1:1000 dilution).
25.Remove sections from the refrigerator and lay the slides flat so that the tissues face upward.
26.Hydrate the sections with PBS for 10 minutes.
27.Remove PBS and cover the sections in the glycerol-based mounting medium (~500 µL/section).
28.Use the side of the pipette tip to evenly spread the mounting medium across the surface of the tissue.
29.Mount the coverslip. Tilt slides to remove excess glycerol and dry off the slides.
30.Load the slides into the microscope trays (Fig. 4A).
31.Insert the trays into the slide scanner (Fig. 4B), and close the outer door (Fig. 4C).
32.Select the appropriate scanning profile and exposure times for the fluorophores. You may also perform dark field imaging in this round to accentuate the mineralized tissue, and polarized light imaging to visualize aligned collagen.
33.Click preview scan to preview each slide.
34.Manually enter the slide names for each slide.
35.Use the tissue detection wizard to select a region of interest (ROI) on each slide, corresponding with the tissue. Save each ROI with the sample name. These ROI’s will be reused for subsequent imaging rounds.
36.Press the start scan button on the slide scanner to initiate scan.
37.Submerge slides in a coplin jar filled with 1x PBS until the coverslips fall off.
38.Apply the TRAP buffer to each of the slides.
39.Check the slides under a fluorescent microscope to determine with the sections are sufficiently decalcified (i.e., the mineral labels disappear). This usually takes 2 h 30 min at room temperature.
40.Prepare the TRAP staining solution by diluting the yellow fluorescent substrate in the TRAP buffer 1:50.
41.Apply the TRAP staining solution to the slides. Again, make sure to cover the entire tissue with the solution.
42.Place the slides under a UV light for 50 minutes.
43.Rinse slides with 1x PBS for 5 minutes 3 times.
44.Prepare glycerol-based mounting medium with TO-PROTM-3 Iodide nuclear counterstain (1:1000 dilution). Use this solution to coverslip the slides.
45.Repeat Steps 30 – 36 to image the slides in the slide scanner. Load the saved region of interest for each respective slide from the previous imaging round. Image slides with the ELF97 filter (TRAP stain) and the Cy5 (TO-PROTM-3 Iodide nuclear counterstain) channel.
46.Submerge slides in a coplin jar filled with 1x PBS until the coverslips fall off.
47.Prepare the AP buffer and the AP staining solution.
48.Incubate the slides for 20 min in the AP staining solution at room temperature.
49.Rinse the slides in 1x PBS for 5 min 3 times.
50.Prepare glycerol-based mounting medium with Hoechst 33342 nuclear counterstain (1:1000 dilution). Use this solution to coverslip the slides.
51.Repeat steps 30 – 36 to image the slides in the slide scanner. Load the saved region of interest for each respective slide from the previous imaging round. Image the DAPI (Hoechst stain) and Cy5 (AP stain) channels.
52.Submerge slides in a coplin jar filled with DI water until the coverslips fall off.
53.Rinse the slides DI water for 5 min 3 times to sufficiently remove the PBS.
54.Prepare the TB stain in DI water.
55.Incubate slides for 2 min in the TB solution.
56.Remove TB and rinse the slides DI water for 5 min 3 times.
57.Prepare fructose-based mounting medium. Use this solution to coverslip the slides.
58.Repeat steps 30 – 36 to image the slides in the slide scanner using a brightfield profile. Load the saved region of interest for each respective slide from the previous imaging round.
59.Open the .czi files for a given sample from each round of imaging in the Zen software.
60.For the ‘Round 1’ image, set a region of interest that you are interested in exporting. Save this ROI in the Zen software.
61.Load the saved ROI onto the images from Rounds 2 – 4.
62.Export the desired ROI’s as .jpg files. Create single channel files for each image (not composite images). Rename each image file with an appropriate description, i.e., Animal1_VertebralBody_DarkField.
63.Using Affinity Photo, create a ‘New Stack’, and open the single channel exported ROI .jpg files from all rounds of imaging.
64.Align images: Check ‘Automatically Align Images’ which will reposition the images to align them.
65.Create layers: In the right-hand menu under ‘Layers’ there will be a ‘Live Stack Group’. Right-click on the ‘Live Stack Group’ and click ‘Ungroup’ to split each image file into a layer.
66.Select all the layers and click on the blend mode dropdown to then click ‘Screen’. This feature will let you see through each layer.
67.If you would like to overlay the fluorescent layers onto the TB layer, change the opacity of the TB layer to 25% ‒ 40%.
68.Transform TB layer: In Affinity Pro, select the TB layer, and click the lock icon to lock the aspect ratio, and type *=percentage% in W: text box, and press ‘Enter’.
69.Manually adjust layers from the different round if they are not properly aligned.
70.Click ‘File’ > ‘Export’ to export the image as a Photoshop file (.psd), .tif, .jpg, or .png.