Mar 09, 2026
Tissue Processing for Clinical Samples for Histology (paraffin, cryosectioning, and clearing)
- Ashleigh Abbott1,
- Robert Dalton1,
- Lauryn Lafayette1,
- Shane Priester1,
- Janak Gaire1,
- Folly Patterson1,
- Jose Peaguda1,
- Kyle D. Allen1
- 1University of Florida J. Crayton Pruitt Family Department of Biomedical Engineering
Protocol Citation: Ashleigh Abbott, Robert Dalton, Lauryn Lafayette, Shane Priester, Janak Gaire, Folly Patterson, Jose Peaguda, Kyle D. Allen 2026. Tissue Processing for Clinical Samples for Histology (paraffin, cryosectioning, and clearing). protocols.io https://dx.doi.org/10.17504/protocols.io.j8nlkq6zwl5r/v1
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 07, 2026
Last Modified: March 09, 2026
Protocol Integer ID: 284501
Keywords: Tissue staining, Knee Joint, Formalin fixed, Paraffin embedded, Safranin O, Fast Green, Cartilage, Osteoarthritis, Formalin Fixed Paraffin Embedded, FFPE, Microscopy, knee replacement, imaging cartilage, cartilage degradation, clinical samples, fixed clinical tissues, Clearing, Tissue clearing, Clinical tissues, Cryosectioning, tissue processing for clinical sample, pathology relevant to osteoarthritis, osteoarthritis, clinical tissue, tissue processing, clinical sample, allocating clinical sample, precious clinical tissue, light sheet microscopy, total knee arthroplasty, histology, fixation of the tissue, immunofluorescence, samples from patient, tissue, several imaging modality, removed tissue, imaging modality, meniscus, bone, infrapatellar fat pad, histopathology
Funders Acknowledgements:
National Institute of Arthritis and Musculoskeletal and Skin Diseases
Grant ID: UC2AR082196
Abstract
The following protocol outlines our approach to allocating clinical samples for three imaging modalities: histopathology, immunofluorescence, and light sheet microscopy. We have used this protocol to process clinical fixed samples from patients with total knee arthroplasty. Once the surgeon has excised and removed tissues to place the implant, these tissues are transferred in saline to be processed for histology and biochemical assays. Following fixation of the tissues, these processes are used to characterize histological features and identify pathology relevant to osteoarthritis. Our protocol works for both hard (bone) and soft (synovium, infrapatellar fat pad, and meniscus) tissues. Our approach enables researchers to pursue several imaging modalities using the same source sample, which is imperative when working with small or precious clinical tissue.
Materials
1. Thermometer
2. pH meter or pH strips
3. Ice bucket and ice
4. Temperature controlled shaker
5. Tissue collection containers and cassettes (labeled)
6. Forceps, tweezers
7. Dry ice or metal bead bath beads stored in -80 °C
8. Kim wipes
9. #10 scalpel blade and handle
10. Razorblade
11. Forceps
12. Camera and camera log
13: Serological pipettes and pipette-aid
14. Glass vials
15. 15 mL tubes
16. Notecards
17. Personal protective equipment (Gloves, lab-coat, masks, fume hood, downdraft table, safety goggles, etc.)
Optional (but recommended): X-ray system (e.g., IVIS CT machine)
Reagents:
| Name | Vendor | CAS number(s) | Catalog Number or Product Code | RRID | |
| Phosphate buffered saline (PBS 10X) | Fisher Bioreagents | Water: 7732-18-5 Sodium chloride: 7647-14-5 Potassium chloride: 7447-40-7 Sodium phosphate dibasic: 7558-79-4 Potassium phosphate monobasic: 7778-77-0 | BP399-20 | AB_2861614 | |
| Paraformaldehyde (PFA) powder | Sigma-Aldrich | 30525-89-4 | P6148-1KG | N/A | |
| Sodium hydroxide (NaOH) | Sigma-Aldrich | 1310-73-2 | 06203-1KG | N/A | |
| Hydrochloric acid (HCl) | Fisher Scientific | 7647-01-0 | A144S-500 | N/A | |
| Decalcifying solution (ImmunocalTM ) | StatLab | 64-18-6 | SKU# 1214, StatLab) | N/A | |
| Sucrose | Fisher Bioreagents | 57-50-1 | BP220-1 | N/A | |
| 2-Methylbutane | Sigma-Aldrich | 78-78-4 | M32631-4L | N/A |
Troubleshooting
Before start
This protocol will review the process from sample creation and storage in PBS to the processing and allocation of samples for the 3 pathways; structural characterization (paraffin) , 2D immune/innervation characterization (cryosectioning), and innervation characterization 3D (PEGASOS clearing).
Preparation of Paraformaldehyde (PFA) and Storage Solvents
Prepare the following solutions prior to surgery day dissection:
1X PBS: Dilute 10X PBS by mixing 1 part of 10X PBS and 9 parts of distilled water (i.e. 10 mL of 10X PBS for 90 mL DI water)
Keep 1-2L refrigerated at 4°C and 1-2L room temperature for use in 4% PFA
4% PFA solution (Note: Prepare PFA in the chemical fume hood and use appropriate personal protective equipment)
Note: recommended to make 2-3L of 4% PFA for each dissection. The below protocol is written for 1L of 4% PFA.
On a stirring hotplate with a magnetic stir bar, heat a volume of 500mL 1X PBS to 55 °C.
Tip: Can preheat 1X PBS in water bath set to 55°C prior start
Ensure PBS does not boil or go above 60°C
Weigh out a quantity of PFA powder equal to 4% volume of the final solution (For 1L final volume, weigh 40 g of paraformaldehyde).
Add PFA powder to the heated 1X PBS and stir, maintaining the temperature at 55°C (keep the temperature below 60°C to prevent breakdown of PFA) until the PFA powder no longer forms large clumps.
Add 2 pellets of NaOH or 1-2 drops of 2M NaOH solution to the PFA solution. The cloudy solution should become clear.
Note: this can take around 20 minutes
Tip: do not add more than 4 pellets per 1L of PFA to prevent issues with pH balancing in step h. PFA can break down at extremely high (basic) pH
Remove from heat and add PBS to reach a volume of ~950mL
Note: can use cold PBS to speed up the cooling process in part f.
Allow PFA to cool to room temperature before adjusting pH (in the hood).
When PFA is cooled, adjust pH to 7.4 as needed using HCl or NaOH.
(Optional: Filter out undissolved particles using a fine filter paper).
Note: ensure that pH meter is calibrated prior to use._
Bring final volume to 1L with 1X PBS.
**Store at 4 °C for immediate use (within one week). For long term storage, store at – 20 °C.**
Prep containers for human sample cassettes at least one day prior.
Make at least 2 1L containers for each dissection.
Label the container with donor ID, date, study, “4% paraformaldehyde in 1X phosphate-buffered saline”, and the date and pH of the 4% PFA. Additional labels depend on the organization of your project/tissue bank; examples include the dissection number and tissue type (e.g., 49A, 49B) A=hard tissues, B=soft tissues, C=extra containers, as needed.
Pour the PFA (stored at 4°C) into the respective containers (one for hard tissue and one for soft tissue).
Keep PFA containers on ice in a cooler during the full duration of the dissection.
Post-dissection Fixation
Place the cassette in a container with 4% PFA. Fix the tissue for 48 hours at 4 °C.
Samples should be, ideally, a maximum of 0.5 cm x 0.5 cm x 0.5 cm. The volume of 4% PFA should be at least 15-20 times greater than the volume of the tissue.
Decant the 4% PFA into a proper hazardous waste receptacle. Rinse cassettes with 1X PBS to remove excess fixative. Incubate in PBS for 30 minutes.
Replace the PBS immediately after the first incubation to rid the sample of most of the residual PFA to prevent further fixation (Rinse 1).
Roughly 24 hours later, replace PBS (Rinse 2).
Roughly 24 hours later, replace PBS with 0.05% NaN₃ in 1X PBS.
Rinses can be separated at minimum by 3-4 hours and maximum a few days.
Note: after the first rinse, the timing of rinse 2 and 3 can be more flexible as long as they are not left in plain PBS (no bacteriostatic preservative) for 3-2 weeks, which increases the risk of microbial contamination or continued fixation.
The final rinse and storage of samples in 0.05% NaN₃ in 1X PBS is to prevent microbial contamination.
Replace the 1X PBS and 0.05% NaN₃ with fresh 1X PBS and 0.05% NaN₃ every six months, at a minimum.
Processing Prep
Note: this depends on the experimental design; in our case, we were preparing samples for microtome sectioning, cryosectioning, and optical clearing (for light sheet microscopy) after sample procurement.
Clearing
- Label glass scintillation vials (1 vial/sample)
- Label should include sample ID, date, initials, storage condition, “for clearing”
- Note: use glass vials to prevent issues with clearing reagents and plastic.
- 0.05% NaN₃ in 1X PBS (~10-15 mL/sample)
Cryosectioning
- Label 15 mL tubes (1 tube/sample)
- Label should include sample ID, date, initials, storage condition, “for cryosectioning”
- 15% sucrose in 1X PBS (~10 mL/sample)
Paraffin
- Notecards labeled with pencil to be placed inside cassettes
- Label should include sample ID
- 1L 1X PBS and container for sample storage
- Samples stored in PBS until they are sent through paraffin processing pathway (dehydration with ethanol ladder)
Setup:
- 0.05% NaN₃ in 1X PBS (~1L) or about 10 mL per sample
- 15% sucrose in 1X PBS (~1L) or about 10 mL per sample
- Serological pipettes and pipette-aid
- Bio-board (disposable dissection board)
- #10 scalpel blade and handle
- Razorblade
- Forceps
- Camera and camera log
Decalcification
NOTE: (skip this step for soft tissues i.e., synovium, fat pad, or meniscus)
Place cassettes containing sample in container with Immunocal solution (~40 mL per cassette)
Incubate samples in 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 CT or use forceps/tweezers to assess the spongy nature of the bone.
We recommend checking status of decalcification via CT, 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 1X PBS (~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.
Once the sample is decalcified, it can be further processed for our various imaging pipelines.
Sample Allocation and Recordkeeping
Retrieve necessary sample cassettes from storage.
Take picture of cassette and tissue prior to processing
This is to create a visual record of where each subsequent sample was sourced.
Using the scalpel and/or razorblade, make two cuts horizontally across the tissue. Tissue slices/segments should be at least 2 mm in width.
The cuts should be made so that each segment is of roughly the same composition and is as representative of the natural physiology as possible. For example, the meniscus should be cut perpendicular to the apex and the periphery (i.e., if cutting the body of the meniscus, the cut should be made in the coronal plane).
Capture a picture of sample + cassette after the sample is cut into segments
Record which segment will be assigned to each pathway
Note: if the sample is cut in unevenly sized segments, larger segments are designated in the following hierarchy: cryosection 3e paraffin 3e clearing. Cryosectioning generally requires larger sections than paraffin-embedded sectioning, and clearing is most optimal with smaller than in larger sample sizes.
If the sample cannot be split in three segments of appropriate size, another sample (ideally, one that is spatially close to the original, in the physiological sense) is also processed.
Each segment is placed in the appropriate container
Cryo-designated segments are placed in 15 mL tubes, which are filled with 15% sucrose ~10 mL/sample, and incubated at 4 °C until the samples sink (see Cryosectioning protocol for further steps)
Clearing-designated samples are placed in a glass scintillation vial with 0.05% NaN₃ in 1X PBS
Paraffin-designated samples are placed back into their original cassettes and submerged 1X PBS. Either the same day or once you have all the samples, then start the ethanol ladder beginning with 30% ethanol. Further steps are outlined in the paraffin processing protocol.
Note: do not begin dehydrating until you have all the cassettes, as you do not want earlier samples to be dehydrated longer.
Acknowledgements
This work was funded by the National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant ID: UC2AR082196.