In the field of life sciences and medical research, 3D cell culture technology is quietly transforming our understanding of the cellular world. Unlike conventional 2D monolayer culture, 3D cell culture, whether spheroids, organoids or other three-dimensional structures, can more faithfully mimic the microenvironment of in vivo cell growth and demonstrates great potential in precision medicine, drug development, regenerative medicine and other fields.
Science Sharing Logic
1 Why do 3D cells need to be made into wax blocks?
·3D cultures are usually tiny in size and fragile in structure. They tend to fragment after fixation and are highly susceptible to loss during washing. If whole-mount staining and observation are performed directly, the entire sample will be consumed in a single experiment, making multiple subsequent assays impossible.
·After 3D cells are prepared into paraffin blocks, dozens or even hundreds of sections can be cut from one block for various assays including HE staining, immunohistochemistry and immunofluorescence. This not only maximizes the utilization of precious samples, but also yields more stable and reproducible test results.
2 Full Workflow of Paraffin Block Preparation for 3D Cells
The core idea of preparing paraffin blocks for 3D cells is similar to that of conventional tissue blocks, covering key steps: fixation, dehydration, clearing, wax infiltration, embedding and sectioning. However, due to the unique properties of 3D cell samples, special attention should be paid to details in each step.
Step 1: Culture and Collection of 3D Cells
Taking the classic 3D spheroid culture as an example, researchers can culture cells in ultra-low attachment culture plates or transwell inserts to grow target cells into three-dimensional spheroids. Collection can be initiated once the spheroids reach an appropriate size. Gentle handling is required during collection to avoid damaging the three-dimensional structure of spheroids. Pre-staining of spheroids with eosin before collection can significantly improve their visibility in subsequent operations for easy positioning and manipulation.
Step 2: Fixation and Pre-treatment
·1. Carefully aspirate the culture medium and rinse gently with PBS.
·2. Fixation: Add 4% PFA and fix at room temperature for approximately 6 hours.
·3. Aspirate PFA and wash thoroughly with PBS.
·4. For 3D cells cultured in transwell inserts: Place the insert under a stereomicroscope. Use micro-scissors to carefully cut the circular PET membrane along the inner side of the plastic support ring of the insert. Wrap the membrane in moist weighing paper and place it into a tissue cassette for subsequent dehydration.
·5. For cells cultured in culture plates: Transfer the sample to a centrifuge tube and perform mild centrifugation to pellet the cells, then aspirate PBS. Prepare 2% low-melting-point agarose and dissolve it in a water bath. Add a small amount of agarose into the centrifuge tube and leave it on ice for 15–20 min until the agarose is fully solidified. Gently take out the agarose with adherent cell aggregates, wrap it in moist weighing paper, and place it into a tissue cassette for subsequent dehydration.
Step 3: Dehydration, Clearing and Wax Infiltration
·1. 70% ethanol, 30 min
·2. 80% ethanol, 30 min
·3. 95% ethanol, 30 min
·4. 100% ethanol I, II, III, 30 min each
·8. Xylene I, II, III, 30 min each
·9. Paraffin infiltration (60℃): Paraffin I, II, III, 60 min each.
Step 4: Embedding and Section Preparation
·Key point for embedding orientation:Embed the membrane slices and cell aggregates in an upright position as much as possible.
·Sectioning:Cut sections at a thickness of 3–5 μm using a paraffin microtome. The sections can be used for routine and special pathological staining including HE staining, IHC and mIF.
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