Strategy for Biobanking of Ovarian Cancer Organoids: Addressing the Interpatient Heterogeneity across Histological Subtypes and Disease Stages

The best clinical management in cutting-edge translational research for patients with advanced ovarian cancer has highest priority at our institution. With the biobanking of patient-derived organoids, we have achieved major progress towards individually tailored management. This enables us to evaluate patient-specific characteristics and to identify predictive biomarkers.

We have made significant advances in recent years in improving radical multivisceral surgery, chemotherapy, anti-angiogenic therapy, and also PARP inhibitor therapy in patients with advanced ovarian cancer and improved survival rates even with that. Now the next big step would be to better predict which patient would benefit from which specific treatment. Currently, clinical treatment decisions are based on general histopathological features and on information on homologous recommendation deficiency or tested in paraffin-embedded tumor tissue.

Biologic models that evaluate specific responses to chemotherapy and to targeted treatments have not yet been implemented in clinical routine. We have identified key characteristics of ovarian cancer tissue, specifically the culture requirements necessary to sustain its renewal potential in vitro, and use this knowledge to achieve robust long-term growth of organoid lines and establish biobank from samples at various stages of the disease. Our day-to-day experience in biobanking of ovarian cancer organoids effectively shows a remarkable heterogeneity in the biology of this malignancy, which could pave the way to new classification or provide the basis for new therapeutic solutions targeting stemness potential.

To begin, take the freshly isolated ovarian cancer tissue. Wash the fresh tissue in a Petri dish thoroughly using PBS without calcium and magnesium. Using disposable scalpels and scissors, fragment the tissue in the Petri dish into small pieces.

Collect the tissue in cryogenic tubes. Then transfer these tubes into a container filled with liquid nitrogen for shock freezing. Next, collect tissue measuring two to three millimeters into a histological specimen container, and then fix the tissue in a formalin container for 24 hours.

Use a scalpel to homogenize the tissue before enzymatic digestion to maximize mechanical dissociation. Then incubate in a water bath at 37 degrees Celsius for 1.5 hours. Post-incubation, introduce 15 milliliters of cold basal culture medium to the tube.

Centrifuge the tube for five minutes at 300 G, remove the supernatant, and add five milliliters of new basal culture medium. Then strain the cell suspension using a 400-micrometer filter into a fresh 15-milliliter tube. To eliminate erythrocytes, add five milliliters of red blood cell lysing buffer to the cell pellet, and incubate in a water bath for five minutes at 37 degrees Celsius.

Post-incubation, add five milliliters of basal medium for lysis inactivation. Pellet the cells by centrifugation, and add three milliliters of basal culture medium to the cell pellet. Now transfer the required number of cells suspended in the basil culture medium into a new tube.

Centrifuge the cells, and after discarding the supernatant, mix the cell pellet with cold BME 2 matrix thoroughly on ice. Seed the cells in droplets of BME 2 matrix onto a prewarmed empty 48-well plate, while pipetting in between to ensure even distribution. Then add 250 microliters of each ovarian cancer medium into the corresponding wells.

To start a 2D culture from the remaining isolated cells, centrifuge the cells for five minutes at 300 g and add the cell pellet to the 2D medium. Use a phase contrast microscope for high-quality images of both 2D and 3D cultures, as well as direct-seeding plates. Image the plate under 4x objective for an overview of the wells and under 10x and 20x objectives for documenting organoid morphology.

Organize and store the pictures in folders dedicated to individual lines. Approximately 14 to 21 days after isolation, evaluate the organoid-forming potential, including quantity, size, and cellular phenotype. Look for specific features, such as the absence of vacuoles, membrane blebbing, loss of adhesion, and rounding up of cells.

Organoid lines were generated successfully from different histological types and stages of ovarian cancer, including primary high-grade serous, post neoadjuvant interval surgeries, and recurrent disease. To begin, take the ovarian cancer organoids cultured in a 24 or 48-well plate. Add ice cold basal culture medium, and pipette or scrape the bottom of the plate to disrupt each BME 2 matrix droplet.

Then transfer the cell suspension into a 15-milliliter tube and keep it on ice. Add one milliliter of ice cold basal culture medium. After combining two to three technical replicate wells for uniform expansion, centrifuge the tube for five minutes at 300 g.

Check the suspension against light to confirm the absence of the BME 2 gel. For cryopreservation of organoids, prepare the cell pellet as demonstrated earlier, and resuspend the cell pellet in one milliliter of ice cold cryopreservation medium. Transfer the cell suspension into pre-labeled 1.8-milliliter cryogenic tubes.

To begin, take the cultured ovarian cancer organoids and collect them. Add three milliliters of 4%paraformaldehyde in PBS to the organoids and incubate for one hour at room temperature. Wash the organoids two times with five milliliters of PBS.

Centrifuge for three minutes at 300 g. After removing the supernatant, add four milliliters of fresh PBS to the pellet. Mix it, and store at four degrees Celsius until embedding.

Now heat the histological gel at 65 degrees Celsius to liquefy it. Then identify fixed organoids settled at the bottom of the tube and remove the supernatant. Resuspend the pellet in 100 microliters of warm gel by pipetting.

Transfer the droplet to a piece of sealing film and allow it to solidify for approximately 15 minutes at room temperature. Move the solidified gel droplet to a tissue paraffin cassette, and follow standard tissue embedding protocols. Using a micro-sectioning cutting tool, slice the embedded tissue into five to 10-micrometer thick sections.

Place these sections onto histology slides. Dry the slides for one hour at 65 degrees Celsius, and then store them in a dry location.