A High-Throughput Platform for Culture and 3D Imaging of Organoids

This paper presents a fabrication protocol for a new type of culture substrate with hundreds of microcontainers per mm², in which organoids can be cultured and observed using high-resolution microscopy. The cell seeding and immunostaining protocols are also detailed.

This protocol demonstrate a microtexture cell culture dish engineered for the growth of hundreds of organoids with knowledge of material and fully compatible with microscopy. Our technique efficiently allow to generate thousands of organoids, fully compatible with long-term imaging and long-term cell culture. This protocol is of strong interest for biomedical applications such as drug screening pipeline and cancer research domains.

The technique presented in this protocol is easy to master after a few attempts by anybody who has experience with laboratory procedures and optical microscopy. To begin, cut small portions of the PDMS mold to the dimension required for the final device. Cut them parallel to the XY directions of the array.

Place one of the prepared PDMS mold dice face down on the flat PDMS section. Then using a pipette, add approximately 0.1 to 0.2 milliliters of UV curable adhesive to one side of the mold. Follow the progression of the liquid inside the cavity using an inverted optical microscope at 10X magnification.

Expose the adhesive to UV light to cure it. Adjust the time of exposure depending on the power density of the UV source as described in the text manuscript. Using the excess amount of adhesive at one edge, hold the cured adhesive on the flat PDMS substrate by gently pressing it with a finger.

Meanwhile, using tweezers, pinch one corner of the mold next to the same edge being held down and slowly peel off the mold, while ensuring that the textured film is not lifted. Trim the excess adhesive and PDMS substrate using a razor blade to leave the cured, textured, and adhesive layer flat on the PDMS with the excess substrate on one edge only. Treat a clean coverslip with a short oxygen plasma process to improve its hydrophilicity.

After the plasma activation, spin coat the thin layer of UV curable adhesive by placing the coverslip on the vacuum chuck of a standard spin coater and pouring a small drop of adhesive at the center of the coverslip. Run the spin coating process as described in the text manuscript. If spin coating is unavailable, drop approximately 0.1 milliliters of UV curable adhesive on a clean coverslip using a pipette.

Take a second coverslip and place it on top of the first to make the liquid adhesive spread evenly between the two coverslips. Once the adhesive has reached the edge of the coverslips, gently separate them by sliding one over the other. Once separated, both coverslips will be fully coated with a thin layer of liquid adhesive.

After spin coating, pre-cure the adhesive by exposure to UV.Adjust the time of exposure depending on the power density of the UV source used. Take one of the textured films and place it in contact with the adhesive coated coverslip. Make sure the contact between the partially cured adhesive and the textured film is as uniform as possible.

At this stage, the adhesive on the coverslip should be solid enough to avoid re-flowing and the contact can be optimized by gently pressing on the coverslip. Expose the coverslips to UV light until the coated adhesive layer is fully cured. This will seal the textured film on the coverslip and provide leak-proof isolation between the perimeter cavities.

Then using tweezers, pinch the PDMS at the edge where the excess material was left after trimming and peel off the PDMS flat substrate. This step enables the textured film layer to adhere to the coverslip with open access at the top for cell seeding. Before cell seeding, on a device passivated with biometric copolymer as described in the manuscript, dispense one milliliter of sterile PBS into the 35 millimeter cell culture Petri dishes.

Degas the dish with sterile PBS using the vacuum chamber for 10 minutes, followed by several rounds of pipetting to remove all the bubbles. Replace the PBS with sterile culture medium and sterilize the plate with UV light for 30 minutes under a cell culture hood. Prepare a cell suspension by following the trypsinization or cell suspension preparation recommendations for the cells of interest as described in the text manuscript.

Count and adjust the cell concentration to 500, 000 cells per milliliter in the recommended cell culture medium. Remove the PBS buffer from the 35 millimeter cell culture dish and then dispense one milliliter of the adjusted cell suspension. Place the cell culture dish back into the cell incubator for 10 minutes.

Approximately 80 to 100 cells will enter each perimeter cavity. After approximately 10 minutes of incubation, recover the cell culture dish from the incubator and gently aspirate the cellular suspension to remove untrapped cells. Add one milliliter of culture medium to the culture dish and place it back into the cell incubator.

Optionally, to retrieve the organoids after they grew in the wells, pinch the textured adhesive layer on the cut edge using tweezers and gently peel it off from the glass coverslip, but keep it immersed in the medium. Increasing the concentration of biometric copolymer increased the coating thickness. Complete degassing of the cell culture dishes was essential to remove air bubbles trapped in the perimeter cavities.

Organoids were formed after days two and 15 of culturing. The spinning disc confocal microscopy showed representative images of the organoids and a 3D reconstruction. When preparing the culture dish, it's important to pay attention to the assembly of the mold or substrate stack and to ensure good contact between the textured film and the coverslip.

The containment provided by our micro cavities and the absence of material loss has attracted the attention of space biology researchers interested in studying the effect of microgravity on organoid homeostasis.