Mar 09, 2026
Processing of Clinical Tissue for Formalin Fixed Paraffin Embedded (FFPE) Samples
- Ashleigh Abbot1,
- Robert Dalton1,
- Lauryn Lafayette1,
- Shane Priester1,
- cruzcarlos 2,
- Kyle Allen1,2
- 1University of Florida J. Crayton Pruitt Family Department of Biomedical Engineering;
- 2University of Florida Pain Research and Intervention Center of Excellence
Protocol Citation: Ashleigh Abbot, Robert Dalton, Lauryn Lafayette, Shane Priester, cruzcarlos , Kyle Allen 2026. Processing of Clinical Tissue for Formalin Fixed Paraffin Embedded (FFPE) Samples. protocols.io https://dx.doi.org/10.17504/protocols.io.q26g7edj1lwz/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 08, 2026
Last Modified: March 09, 2026
Protocol Integer ID: 283831
Keywords: Knee joint, FFPE, Clinical tissues, Histology, Paraffin histology, Decalcification of bone, Formalin fixed, Paraffin, Wax infiltration, Embedding, Decalcification, Wax embed, Formalin fixed paraffin embedded, processing clinical tissue, clinical tissues for formalin, pathology relevant to osteoarthritis, paraffin histology, cartilage degeneration in osteoarthritis, clinical tissue, processing of tissue, tissue type, cartilage degeneration, storage of the tissue, osteoarthritis, formalin fixed paraffin embedded, total knee arthroplasty, tissue, histology, infrapatellar fat pad, removed tissue, samples from patient, fixed paraffin
Funders Acknowledgements:
National Institute of Arthritis and Musculoskeletal and Skin Diseases
Grant ID: UC2AR082196.
Abstract
This protocol outlines a step-by-step workflow for processing clinical tissues for formalin fixed paraffin embedded (FFPE) samples. Following fixation and storage of the tissues, these processes are used to characterize histological features and identify pathology relevant to osteoarthritis. 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 other processes. Our protocol works for both hard (bone) and soft (synovium, infrapatellar fat pad, and meniscus) tissues. Paraffin histology is commonly performed to identify structural features and pathology of disease, particularly cartilage degeneration in osteoarthritis. Processing of tissues for paraffin histology is an important and impactful step that will influence the results of the sections and images downstream. Therefore, our protocol has been optimized to best preserve all tissue types and structure for histological characterization.
Materials
Ice bucket and ice
Microtome blade
Razor blade
Microscope slides
Microtome
Slide warmer
Microscope
Kim Wipes
Tweezers
Paint brushes
Pencil
Water bath
Slide boxes
Personal protective equipment (Gloves, lab-coat, masks, fume hood, downdraft table, safety goggles, etc.)
Reagents:
| Name | Vendor | CAS number | 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 | |
| Ethanol | Decon Labs | 64-17-5 | 2701 | N/A | |
| Paraffin Wax | Fisherbrand | Paraffin waxes and hydrocarbon waxes: 8002-74-2 Polyisobutylene: 9003-27-4 2,6-Di-tert-butyl-p-cresol: 128-37-0 | 22900700 | N/A |
Troubleshooting
Overview of Processing Clinical Tissues for Histology Analysis
Note: this protocol focuses on the structural characterization pathway shown in the flow chart below.
Flow chart of processing used in our study. This protocol focuses on structural characterization (paraffin) only.
Sample Dehydration and Wax Infiltration
After allocating samples to paraffin, the sample will be placed in a cassette labeled with pencil and notecards placed inside the cassette.
Note: pencil can mostly withstand ethanol and other reagents used in this process, but pen and sharpie will be washed off.
During processing, all paraffin-designated samples will be placed in a large container of 1X PBS (depending on number of samples).
The samples will then be dehydrated using an ethanol ladder of 30%, 50% and 70%.
Using ~10-20X volume of ethanol to sample, the samples will be placed in 30% ethanol for 20 minutes, followed by 20 minutes in 50% and lastly placed in 70% ethanol.
Once dehydrated, the samples will be stored briefly in 70% ethanol until they are infiltrated at the molecular pathology core.
Paraffin wax infiltration is done in an infiltration machine using high pressure to infiltrate each sample with wax.
Note: depending on tissue type, the length of infiltration can be adjusted to the settings available on the machine.
Sample Embedding
Once infiltrated, the samples are then embedded in paraffin wax using a Leica EG1150 embedding machine.
Note: we use a temperature range of 60-65°C above the paraffin melting temperature.
Fill metal embedding tray with wax and place sample in the tray.
Keep the sample submerged in the wax at all times. Removal from the wax may introduce air bubbles into the sample, which would require re-infiltration.
Be sure to keep track of sample ID/order in all subsequent steps.
Optional: if available, move metal trays with the samples to the vacuum oven. Turn vacuum to ~15 kPa to pull air bubbles from the samples and wait 10-15 minutes.
Goal: to have melted wax under negative pressure that is NOT boiling and just barely melted. If you are too hot, you will cook your sample. Wax will melt differently at lower pressures.
Use this step if you have poor infiltration.
If vacuum oven is unavailable use the following step:
Place metal embedding tray with sample in the warming compartment of the embedding station for a total of 1 hour, using tweezers to agitate the sample in the tray roughly every 15 minutes. This is to remove any residual air bubbles.
Again, make sure not to remove the sample from the wax to prevent reversing infiltration.
Move samples back to the embedding station. Use warm tweezers to move the sample and make sure all major air bubbles are removed. If necessary, repeat step 10/11.
Do not remove the sample from the wax as this will reverse the infiltration step.
Position sample appropriately and hold with tweezers. Carefully move metal tray to the cold plate on the embedding machine and let the wax on the bottom begin to harden and set the sample. After ~30 seconds press the cassette onto the sample tray and add more wax to fill the entire cassette. Let the sample set at room temperature for 24 hours.
Note on orientation for embedding:
The sectioning plane of interest should be parallel to the bottom of the embedding tray for sectioning. I.e. you want the surface of interest on the bottom of the cassette (parallel plane)
Note this depends on sample type and plane of interest. Consider how you will mount the sample when embedding, as well. Also, place the flattest side down when possible.
Optional: Can leave sample on cooling plate for a few hours to set sample, if desired.
After the sample solidifies overnight, you can remove the sample from the metal tray by placing on the cold plate for 15+ minutes to loosen and remove wax from the sample holder.
Do not try to pry the sample out of the cassette at room temperature. This is likely to crack the wax block, which will then need to be re-embedded.
Preparation for Sectioning
Turn on water bath and set temperature to 42°C (takes approximately 30 minutes).
Note: turn water bath down to 38°C if sectioning fatty tissues
Turn on slide warmer and set temperature to 40-50°C.
Begin preparing sample by trimming the edges of the paraffin block carefully with a razor blade.
Make sure there is wax at all four sides (do not cut too closely to the sample). It is better to leave more wax than to not have enough.
Edges should be straight and parallel, otherwise the sections will not form a ribbon while sectioning.
Do not cut all the way down to the cassette as this will loosen the sample and potentially cause it to detach from the cassette.
Set up microtome
Use the control panel to back up the sample holder on the microtome. This is located on the right-hand side of the base. Avoid moving the sample holder all the way forward or back, as the microtome does not take consistent section thicknesses at the beginning or end of its travel.
Load the sample by pulling the lever and placing the sample at the proper orientation.
Note: it is best to mount the sample such that the largest portion of tissue is closest to the blade when sectioning.
Note: for bone cut the cartilage first and the bone after (orientation).
Sectioning
First, use the trim function (~20 um) to take larger slices and remove the layer of wax covering the tissue. Unlock the microtome arm and begin sectioning until reaching the tissue.
Once the tissue is reached, switch to the section function (to be optimized for 10-20 µm) and set to the desired sectioning thickness. Begin sectioning and use the paintbrushes or forceps to pick up and uncurl the paraffin wax ribbons that will form. After taking 10 slices, gently place the ribbon of slices in the water bath by anchoring one end to the edge of the water bath.
Section mostly between 10-12 µm but may need to increase thickness for difficult tissues.
Using a slide, after the sample is wrinkle free, pick up the samples ~3-4/slide.
NOTE: If having issues with wrinkling even with the water bath, you can use a dilute ethanol bath to float the samples on prior to placing the sectioned ribbons in the water bath.
This is helpful for when the sections come out shriveled or shrunk, this will help the sample ease the tension and release the wrinkles.
The ethanol (15%) helps act as a surfactant in the water allowing the wax to relax.
To transfer the slides from the ethanol bath to the water bath, use a cooled slide (the back non-charged side) to pick up and transfer the ribbon.
Use a KimWipe to wipe off any excess water and then look at the slide under the microscope.
Collect 10-15 slides per sample.
Exceptions: You see something interesting in a particular area and want to collect more slides. You are having trouble getting good sections after extensive troubleshooting. You need extra slides for more stains.
Save the sectioned blocks in case you need to take more sections from a sample.
The slides are then dried for 12+ hours on a slide warmer (~40-50°C) before being stored.
Slides are stored in slide boxes until they are selected for staining. Please see the staining protocols.
Acknowledgements
This work was done in a grant funded by the National Institute of Arthritis and Musculoskeletal and Skin Diseases
Grant ID: UC2AR082196.