Mar 11, 2026
Version 2
Cryosectioning and Immunolabeling of Clinical Tissue Samples V.2
- Robert Dalton1,
- Ashleigh Abbott1,
- Janak Gaire1,
- Elisabeth Coller1,
- Jose Peaguda1,
- Matthew Scolieri1,
- Shane Priester1,
- Kyle D. Allen1
- 1University of Florida - Department of Biomedical Engineering
- Kyle D. Allen: Principal Investigator of the orthoBME Laboratory/corresponding author
Protocol Citation: Robert Dalton, Ashleigh Abbott, Janak Gaire, Elisabeth Coller, Jose Peaguda, Matthew Scolieri, Shane Priester, Kyle D. Allen 2026. Cryosectioning and Immunolabeling of Clinical Tissue Samples. protocols.io https://dx.doi.org/10.17504/protocols.io.n92lddq4nl5b/v2Version created by Robert Dalton
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 10, 2026
Last Modified: March 11, 2026
Protocol Integer ID: 312996
Keywords: Formaldehyde fixed, Cryosection, Clinical samples, Decalcification of clinical tissues, Osteoarthritis, Knee joint, Immunofluorescence, Microscopy, Confocal microscopy, immunolabeling of clinical tissue sample, immunofluorescence imaging, immune cell markers via immunofluorescence imaging, immunostaining clinical sample, immunofluorescence, clinical tissue sample, total knee arthroplasty, immune cell marker, tissue pathology, immunolabeling, fixation of the tissue, clinical tissue, clinical sample, cryosectioning, tissue, histopathological feature, meniscus, pathology
Funders Acknowledgements:
National Institute of Arthritis and Musculoskeletal and Skin Diseases
Grant ID: UC2AR082196
Abstract
This protocol describes the stepwise process for preparing, sectioning, and immunostaining clinical samples obtained from total knee arthroplasty (TKA) participants. Our protocol works for both hard (bone) and soft (synovium, infrapatellar fat pad, and meniscus) tissues. In our work, we utilize this procedure to visualize innervation, vasculature, and immune cell markers via immunofluorescence imaging. Following fixation of the tissue, these processes are used to characterize histopathological features and identify cellular and tissue pathology.
Materials
| Item | Vendor/Supplier | Catalog Number | CAS number(s) | |
| Decalcifying solution (Immunocal) | StatLab | SKU# 1214 (StatLab) | 64-18-6 | |
| Base molds | Epredia | 22-050-159 | ||
| Reagent reservoir | VistaLab Technologies | 3504-1000 | ||
| Optimum Cutting Temperature (OCT) medium | Fisher Scientific | 23-730-571 | Polyvinyl alcohol: 9002-89-5 Polyethylene glycol: 25322-68-3 Water: 7732-18-5 | |
| Dry ice | Fisher Scientific | NC0988943 | 124-38-9 | |
| Sucrose | Fisher Scientific | BP220-1 | 57-50-1 | |
| 2-methylbutane | Sigma-Aldrich | M32631-4L | 78-78-4 | |
| Superfrost plus slides | Fisher Scientific | 1255015 | ||
| Razor blade | Pincover Industrial | 62-0177 | ||
| Epredia MX35 Premier low-profile microtome blades | Fisher Scientific / Epredia | 3051835 | ||
| Epredia HP35 Coated high-profile microtome blades | Fisher Scientific / Epredia | 3150734 | ||
| Coplin glass staining jars | Alkali Scientific | JS005 | ||
| Slide-staining tray | Millipore Sigma | Z670146 | ||
| PAP pen | Abcam | ab2601 | ||
| Normal goat serum | Abcam | ab7481 | ||
| Normal donkey serum | Abcam | ab7475 | ||
| Bovine serum albumin | Millipore-Sigma | A5611 | ||
| Primary antibodies | ||||
| Secondary antibodies | ||||
| VECTASHIELD Vibrance Antifade Mounting Medium | Vector Laboratories | H-1700-10 | ||
| #1.5 24mm x 50mm coverslips | Globe Scientific | 1415-15 |
Troubleshooting
Fixation
Fix 0.5 cm x 0.5 cm x 0.5 cm (maximum size, ideally) tissue samples in 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) for 48 hours at 4 °C.
The volume of 4% PFA should be at least 15-20 times greater than the volume of the tissue.
See "Tissue Processing of Clinical Samples for Histology (paraffin, cryosectioning, and clearing)" protocol for 4% paraformaldehyde recipe and detailed fixation steps.
After 48 hours, decant the PFA into an appropriate hazardous waste container. Rinse the samples in 1X PBS to remove residual fixative. Decent into the hazardous waste container.
Incubate samples in 15-20x their volume of 1X PBS for 30 minutes at room temperature (RT; 20 – 22 °C).
Decant the 1X PBS solution into the hazardous waste container. Replace the 1X PBS with fresh 1X PBS. (Rinse 1)
Roughly 24 hours later, replace PBS (Rinse 2).
Roughly 24 hours later, replace PBS with 0.05% NaN3 in 1X PBS for long-term storage (maximum of 6 months; replace 0.05% NaN3 in PBS solution after 6 months to prevent microbial contamination)
Decalcification (for hard tissues only; proceed to “Cryoprotection” if tissue is not naturally calcified/mineralized, e.g., bone)
Place cassettes containing sample in container with Immunocal solution (~40 mL per cassette).
Incubate samples on a shaker set at 100 rotations per minute at room temperature (20 – 22 °C).
Replace the Immunocal solution every 2-3 days. Collect Immunocal solution in a hazardous waste container and dispose of it accordingly.
Complete decalcification for bone samples ~2 cm x 2 cm x 2 cm is about 7-9 days at room temperature.
Ensure the sample is fully decalcified using X-ray or µCT or use forceps/tweezers to assess the spongy nature of the bone.
We recommend checking status of decalcification via µCT, as the “squeeze test” is not sensitive enough to determine whether the core of the sample is fully decalcified.
After decalcification, wash the specimen with distilled water (~40 mL per cassette) and place it in a shaker for 3-4 hours. Repeat this step two more times. Collect the waste in a hazardous waste container.
Cryoprotection
Place the sample in 15% sucrose solution (prepared in 1X PBS) overnight at 4 °C or until the sample sinks.
If the sample sinks immediately, keep the sample in 15% sucrose overnight at 4 °C.
Use ~ 10-15 mL of sucrose solution per sample.
If the sample does not sink after 24 hours, replace the 15% sucrose in 1X PBS with fresh 15% sucrose in 1X PBS.
If the sample still does not sink by the end of the second incubation period, continue to step 14.
Transfer the sample into 30% sucrose solution and incubate at 4 °C overnight or until the tissue sinks. If the tissue does not sink, replace the sucrose solution, and incubate for an additional day.
OCT Embedding
Gather all necessary materials. Place chux pad over a clean surface in the fume hood and retrieve sample(s) from the 4°C refrigerator.
Fill a Styrofoam container with at least 1-2 inches of dry ice.
Pour out sucrose solution into a Pyrex bottle, using cell strainer to catch the sample, if needed. Place the specimen on a Kimwipe and remove any excess liquid.
Line the bottom of the base mold with a layer of OCT and place specimen in the mold with the flattest side facing down (or oriented so that the region of interest is parallel with the surface or base of the mold). Fill the rest of the mold with OCT until sample is completely covered. Be careful not to introduce any air bubbles.
Place the reservoir on the dry ice and pour 20 mL of 2-methylbutane into the reservoir. Note that 2-methylbutane evaporates very quickly and should be kept at – 80°C until needed.
An alternative to using a reagent reservoir is to form a “pool” structure with aluminum foil by folding up the edges of a rectangular piece of foil. The 2-methylbutane can be poured into this “pool”/container.
Use tweezers or a hemostat to position the mold in the 2-methylbutane so that the bottom is completely submerged. Avoid any overflow onto the specimen and adjust orientation as needed to ensure sample freezes evenly. Keep mold in the 2-methylbutane until the sample has frozen/the OCT has turned completely white. If the sample is not completely covered by the OCT, add more OCT and continue until fully frozen.
Once frozen, samples can be immediately sectioned or stored.
For storage, wrap the mold containing the sample in aluminum foil and use tape to seal and label. Samples can be immediately sectioned or placed in zip-lock plastic bags and stored at – 80°C (long-term).
Once finished, pour 1 part of bleach per 9 parts of liquid waste into the Pyrex bottle. Let stand for a minimum of 30 minutes. After incubation, the waste-bleach solution can be safely disposed of down the drain with running water.
Cryosectioning
Bring the sample(s) to the cryostat. Check if the temperature of the cryostat is in the optimal range (between –15 °C to –22 °C. –20 °C is a good starting point).
Place paintbrushes, tweezers, labeled slides (labeled in pencil), and slide box in the cryostat chamber.
Slides should be labeled with the study name, sample ID, tissue type, date, your initials, slide number, and sectioning size.
For specimens embedded in OCT, use a razor blade to trim the block as needed, but be careful not cut too close to the specimen. Squeeze OCT medium onto the puck and place the tissue block on top of OCT. Let the specimen sit in the cryostat chamber until OCT solidifies and specimen is safely adhered to the puck. It takes ~2-3 minutes.
Slide the microtome blade into the blade position and lock the blade in place. Ensure the blade guard is covering the blade to prevent accidental injury.
Retract the specimen head fully (i.e., away from the user). Mount the puck and specimen on the specimen head.
Carefully bring the specimen head forward until the mounted specimen is within 5-10 mm of the blade.
Align the surface of the specimen with the angle of the blade.
Start trimming the surface OCT on trimming mode until the blade is close to the tissue, adjusting the cutting plane as needed. The blade should cut through the OCT block as evenly as possible.
Once the blade begins sectioning the tissue itself, switch to sectioning mode at 15-30 microns thick. Use cold forceps or fine paint brushes to hold the section down as it is being sectioned.
Some troubleshooting of cutting temperature may be needed. If the section rolls up excessively, the specimen may be too cold. If the section sticks to the blade or easily tears, the specimen may be too warm.
Once a section is fully cut, place the section onto a cold slide. Use cold forceps and paint bushes to unroll and orient the section, taking care to not puncture or tear the section. Make sure the sample is as flat as possible before placing a finger on the underside of the slide to warm up the section. The section will adhere to the slide as the OCT melts. Collect multiple sections per slide (depending on the size of the specimen), leaving room to create a hydrophobic barrier around and between the sections (at least ~3-5 mm between sections). Do not place sections towards the edges of the slide, as this will create difficulty with staining in later steps.
Continue sectioning and collecting the sections on glass slides. Record any observations (e.g., wrinkling of a certain section, missed sections, etc.), as well as a basic drawing of the sample, while you are sectioning for later reference.
Slides can be immediately processed for immunostaining or stored in -20 °C (for few weeks) or – 80°C (for long-term storage). Wrap the slide box with aluminum foil to prevent frost buildup on the box. Label the box and aluminum foil with study name, date, initials, sample ID, tissue type(s), and box number on the top and side of both the slide box and aluminum foil.
Immunostaining
Prepare the following solutions:
1X PBS: Dilute 10X PBS by taking 1 part of 10X PBS to 9 parts of distilled water
1X PBST: Add 0.05% to 0.1% Triton-X or Tween-20 to 1L PBS. Tween-20 is a gentler detergent than Triton X and recommended for surface staining. Triton X can be used for permeabilization of cell membranes in order to stain intracellular structures/proteins.
Blocking solution: Prepare blocking solution consisting of 1% bovine serum albumin, 4% normal goat serum or normal donkey serum (depending on the secondaries used), 0.05% Triton-X or Tween-20 in PBS
Antibody mixture: Prior to incubation in primary and secondary (if using unconjugated primary antibodies) solution, dilute antibodies in blocking solution. Dilution should be predetermined.
Bring the slides/tissue sections (freshly sectioned or stored at -20 °C or -80 °C) to room temperature (RT).
Rehydrate tissue sections in 1X PBST for 10 mins in Coplin jars.
Aspirate excess PBST and apply PAP pen to create a hydrophobic barrier around tissue sections. Take care to avoid PAP pen liquid coming in contact with the tissue sections.
Place slides in slide-staining tray containing a shallow layer of deionized water. Block sections in blocking solution for 1 hour at RT.
Aspirate the blocking solution and incubate tissue sections in primary antibody or antibodies diluted (For multi-labeling experiments, use antibodies raised in different hosts) in blocking buffer for overnight at 4 °C.
Aspirate primary antibody solution and wash the slides in 1X PBST (3 times, 5 min interval between washes).
Incubate tissue sections in secondary antibodies raised in host species of primary antibodies (For multi-labeling experiments, use spectrally well-separated secondaries and ensure the instrument used for imaging is equipped for the selected fluorophores) prepared in blocking solution for 1-1.5 hour at RT in dark. Here onwards, protect slides from exposure to light.
Aspirate the secondary antibody solution and wash sections with 1X PBST (3 times, 5 min interval between washes). Optional: Add DAPI (1:1000) to counterstain nuclei in the second wash.
Aspirate PBST. Move quickly so that tissue sections do not dry out; place a drop of mounting medium (VECTASHIELD Vibrance hardset) on each tissue section, plus any remainder on the slide area between the sections (~66uL in total for a 24mm x 50mm slide cover).
Coverslip the slides by carefully lowering the glass coverslip onto mounting medium droplets. Allow gravity and capillary action to draw the mounting medium across the slide. There should be a minimal amount of bubbles. Bubbles on tissue sections will impact imaging.
If there are bubbles on top of any tissue sections, DO NOT try to press the bubbles off of the sections - this can disrupt the tissue morphology and cause high background staining if cells are crushed.
Allow slides to cure for at least two hours at RT before imaging. If not immediately imaging, cure slides overnight at room temperature (protected from light).
If your tissue sections are thicker than 10 µm, you will need to seal the slides with plastic sealant or nail polish. After the slides have cured, apply a thin layer of sealant/nail polish around the edges of the coverslip, taking care not to brush the sealant over the tissue sections. Allow sealant/nail polish to set overnight, protected from light.
Store slides at 4 °C. Signal intensity should be stable for several months.
Protocol references
Fra-Bido, Sigrid, et al. “Optimized immunofluorescence staining protocol for imaging germinal centers in secondary lymphoid tissues of vaccinated mice.” STAR Protocols, vol. 2, no. 3, Sept. 2021, p. 100499, https://doi.org/10.1016/j.xpro.2021.100499.