Handling and Assessment of Human Primary Prostate Organoid Culture

The following protocol was optimized using human primary prostate epithelial cells harvested from patient tissue at the University of Illinois at Chicago Hospital. All human tissues used for these experiments were acquired via an Institutional Review Board-approved protocol and/or exemption at the University of Illinois at Chicago. While the culture conditions will vary depending on the cell type of interest, the endpoints can be applied to organoids of other tissues.

1. Seeding of Epithelial Cells into Matrix Gel and Changing Media

1. Collect necessary materials: human primary prostate epithelial cells (PrE), matrix gel on ice, 96-well microplate, keratinocyte serum-free media (KSFM) on ice, charcoal stripped fetal bovine serum (FBS), and dihydrotestosterone (DHT).
2. Prepare KSFM-based organoid media by combining 47.5 mL of KSFM with 2.5 mL of charcoal stripped FBS and dihydrotestosterone (DHT) to a final concentration of 10 nM DHT, then reserve on ice.
3. Plate cells into a 3D matrix in a cell culture hood using sterile technique.
   1. Gently mix cold KSFM-based organoid media with ice-cold matrix gel to obtain a 50% matrix gel mixture by slowly pipetting up and down, avoiding bubbles.
      NOTE: Matrix gel should be thawed overnight at 4 °C and kept on ice whenever possible. It solidifies quickly at room temperature (RT).
   2. Pre-wet a pipette tip in cold media and transfer 40–50 μL of 50% matrix gel onto the bottom of each well to be used on a 96-well plate. Coat the bottom of the well to form a base layer.
      NOTE: Do not fully dispense to the second stop on pipette, as this can create bubbles.
   3. Place the 96-well plate into 37 °C incubator for 30–45 min to solidify the base layer.
      NOTE: Do not plate epithelial cells onto the base layer until it has solidified, or cells may fall through the matrix onto the bottom of the plate and grow as a monolayer.
   4. Mix epithelial cells suspended in KSFM-based organoid media with matrix gel to achieve a final concentration of 100–1,000 cells/100 μL and 33% matrix gel. Plate epithelial cells on top of base layers using 100 μL of 33% matrix gel/cell mixture per well.
      ​NOTE: Previous experiments have shown that seeding epithelial cells too densely in 3D will restrict the size of organoid growth, while seeding too sparsely can result in very low yield¹³. It is important to optimize seeding conditions for the cell type of interest, based on the patient-specific organoid formation efficiency.
4. Place the 96-well plate in a 37 °C incubator for 30–45 min to allow the matrix gel to solidify, then gently add 100 μL of KSFM-based organoid media on top of cells by pipetting slowly against the edge of the well.
5. Change the media every 2–3 days. Pipetting against the side of the well with space between the media and matrix gel, carefully remove 50 μL of media and replace with 50 μL of fresh media.
   NOTE: For a long-term culture, the matrix gel needs to be changed every 2–3 weeks. If the matrix gel culture appears unstable and displays translucent wrinkles under the microscope, then the matrix gel has broken down and needs to be replaced.

2. Collection of Organoids from Matrix Gel

NOTE: Collection of organoids is necessary for matrix gel changes, passaging or endpoints of RNA extraction, DNA extraction, protein extraction, and flow cytometry.

1. Warm neutral protease and Hanks' balanced salt solution (HBSS) in a 37 °C water bath.
2. Carefully remove media from wells without disrupting the matrix gel.
3. Add ~200 μL of neutral protease to each well at a ~1:2 ratio of matrix gel:neutral protease and incubate for 20 min at 37 °C.
4. Mechanically dissociate the matrix gel:neutral protease mixture by pipetting up and down.
5. Incubate for 20 min at 37 °C.
6. Transfer the neutral protease-matrix gel-cell mixture to a microcentrifuge tube or conical tube, depending on the volume. Centrifuge at 300 x g for 3–5 min to pellet cells.
   NOTE: If no cell pellet appears, the matrix gel may not be completely dissociated and can be visualized as a translucent phase at the bottom of the tube distinct from the pink supernatant. Remove the supernatant and add more neutral protease at a 1:2 ratio to matrix the gel pellet, incubate for another 20–40 min at 37 °C, and repeat centrifugation at 300 x g.
7. When an organoid pellet is acquired, remove the supernatant. Proceed with passaging (see step 2.7.1), organoid lysis for RNA, DNA, or protein extraction (see step 2.7.2), processing single cells by flow cytometry, and single-cell sequencing (see step 2.7.3).
   1. For long-term culture, gently resuspend intact organoids in 33% fresh matrix gel and plate by following steps described in steps 1.1–1.5. To dissociate organoids and replate as single cells, resuspend the pellet in 1 mL of warm cell-dissociation enzymes and incubate at 37 °C for 10–15 min, wash once with 5 mL of HBSS, and re-plate in the fresh matrix gel following steps 1.1–1.5.
   2. Resuspend the organoid pellet in an appropriate lysis buffer for RNA extraction, DNA extraction, or protein extraction as listed in the Table of Materials and follow the manufacturer’s protocol.
   3. Resuspend organoids in 1 mL of warm cell-dissociation enzymes and incubate at 37 °C for 10–15 min to dissociate the organoids into single cells. Wash once with 5 mL of HBSS and proceed with the protocol for flow cytometry⁵ or single-cell sequencing¹⁸.
      NOTE: To dissociate into single cells at low concentrations of matrix gel, the neutral protease may not be needed, and cell-dissociation enzymes can be applied directly to the well after step 2.2.

3. Whole-well Image Acquisition and Analysis of Organoid Morphology

1. Acquire whole-well z-stack images using a transmitted light inverted microscope with a motorized X/Y scanning stage (workflow illustrated in Figure 1A).
   1. With the focal position adjusted to the bottom of the well, begin focusing upward until the first organoid is encountered, which will be in the center of the well due to the meniscus from the base layer. Set the bottom z-plane at this position.
   2. Continue focusing upward until the last organoid is in focus, which will be in the periphery of the well. Set the top z-plane at this position.
      NOTE: After setting the top and bottom of the z-stack, ensure that these positions capture the organoids within all remaining wells and adjust the top/bottom as needed. This allows for the z-range to be used for a single scan that includes every organoid within each well.
   3. Capture 15–35 z-planes per well, using a larger number for greater resolution. Capture the entire well image, merge and collect quadrants or subsections if needed.
      NOTE: It is recommended to take multiple z-planes as opposed to a single z-plane, as illustrated in Figure 1B in which organoids are out of focus when capturing only one z-plane.
   4. Save z-stack images for downstream analysis.
2. Analyze images using image software with freeform drawing capabilities.
   1. Open a single z-plane image set from step 3.1.4.
   2. If needed, tile the images taken at each individual z-plane into a single, whole-well image. Save the new image as a separate file. Repeat this process for each z-plane acquired.
   3. Using image software, open the whole-well image for every z-plane from a single well and create an extended depth of field (EDF) image from the stack of z-planes.
      NOTE: This type of image will select the best focus region of each z-plane to create a single in-focus image of the well.
   4. Set the pixel scale of the software to the scale bar of the image that is being measured.
   5. Using the freeform drawing tool of the image software, trace the perimeter of every organoid in the well.
   6. Obtain the measurement metrics for traced organoid perimeters from the image software.
      NOTE: Recommended parameters are area, circularity, and maximum/minimum radius ratio. Representative results illustrating the application of these metrics are shown in Figure 1C. Area is calculated from the polygon defined by the organoid’s perimeter. Circularity is the ratio of the area of the organoid to the area of a circle with a diameter equal the organoid’s maximum ferret, where 0 is less circular and 1 is more circular. Maximum/minimum radius ratio is the ratio between the maximum radius and minimum radius of the traced organoid.
      Circularity =
      Radius ratio =

4. Formalin Fixation and Paraffin Embedding of Organoids for Histological Endpoints

1. Prepare embedding supplies: HBSS, 2% agar solution, histological marking dye, histology gel, pipette tip mold, tape, plunger from a 1 cc insulin syringe, ice pack, cassettes, 10% neutral buffered formalin (NBF), pencil, and 75% histological grade ethanol (EtOH).
   1. Prepare two microcentrifuge tubes of 2% agar solution. Incubate in a water bath or on a heat block at 100–110 °C to maintain agar in liquid form. Add 1–2 drops of histologic marking dye to one of the 2% agar solutions, which will denote the top of the agar plug and assist in maintaining orientation during embedding. Mix well.
   2. Warm the histology gel in a water bath or heat block at 55–65 °C.
   3. Using a 1000 μL pipette tip, create molds by cutting out a small section of the pipette tip to make a cylinder that holds ~50 μL. Create a second, wider mold that fits around the first mold.
   4. Create a cold block by wrapping an ice pack with tape (adhesive side up) and adhering molds to the tape.
   5. Create a plunger by removing the plunger from a 1 cc insulin syringe.
2. Embed the organoids in a histology gel plug for processing, following the workflow depicted in Figure 2A.
   1. Dissociate the matrix gel and pellet organoids by following steps 2.1–2.7.
   2. Wash the organoid pellet once with 1 mL of HBSS and re-pellet by spinning for 3–5 min at 300 x g and removing the supernatant.
   3. Re-suspend the organoid pellet in 30–50 μL of liquid histology gel. Mix gently by pipetting up and down slowly. Transfer the histology gel/organoid mixture into a mold on the cold block and allow the specimen to cool and solidify into a plug.
   4. Use a plunger to push the plug out of the small mold and into the larger mold. Pipette clear 2% agar around the plug to fill the larger mold.
   5. Pipette histologic dye-tinted 2% agar on top of the plug to denote the top of the sample.
   6. Once cooled, use a plunger to transfer the plug into a histology cassette. Label the cassette using a pencil and place cassette into 10% NBF for fixation overnight.
   7. Transfer the cassette from 10% NBF to 70% histological grade EtOH.
3. Process the histology gel plug using a pre-set biopsy protocol on a processor, if available. If preset is not available, input the following sequence and lengths: 70% EtOH for 6 min, 70% EtOH for 10 min, 80% EtOH for 10 min, 95% EtOH 2 times for 10 min, 100% EtOH 3 times for 10 min, xylene 3 times for 15 min, and paraffin 3 times for 15 min. Deviating from the recommended processing sequence can result in ruptured organoids (Figure 3A).
4. Embed the histology gel plug with the colored portion facing up, as this will allow the discolored portion containing the organoids to be sectioned into first.
5. Section the FFPE histology gel blocks:
   1. Collect necessary supplies: FFPE histology gel blocks, histology pen, tissue float water bath, ice bucket with ice, microtome, microtome blades, embedding workstation, positively charged microscope slides, and laboratory oven.
   2. Fill water bath until very full and set to 40 °C, then pre-warm laboratory oven to 45–50 °C.
   3. Prepare an ice bucket, then fill with ice and add water to create an ice-water slurry. Chill the blocks on ice until they are very cold.
      NOTE: Blocks can be set on ice for up to 1 day, but if left overnight, the blocks will become hydrated and will need to be reprocessed.
   4. Insert a block into microtome cassette clamp, add the blade to the knife holder, and unlock any protective locking measures on the machine.
   5. Begin by trimming the face of the paraffin block (facing off) at a 10 µm thickness. Once a section emerges that contains the plug area, stop trimming, lock the microtome rotor, and transfer the block back onto the ice to chill. If several blocks are to be sectioned, face-off each block before moving to step 4.5.6.
   6. Move the blade to a new, unused portion and transfer the block back to the clamp. Unlock the machine and begin trimming at 5 μm per section.
      NOTE: 5 μm is recommended for best results, but 3–10 μm is also acceptable.
   7. Using tweezers, transfer the section to the 40 °C water bath and allow the section to spread out. Transfer the section onto a slide by dipping vertically into the water.
      NOTE: Dipping at an angle can transfer bubbles from the bottom of the bath to the section and cause distortion of the sample.
   8. Visualize the section under microscope (Figure 2B,C).
      NOTE: If an organoid present, it will appear similar to the red arrow in Figure 2B. Bubbles (Figure 2B, blue arrow) can appear similar to organoids and are easier to discern on a fresh-slide as dry organoids appear translucent (Figure 2C). If no organoids are present, section further into the block.
6. When slides containing organoids have been acquired, circle the area around the section on the back of slide with a histology pen and bake overnight at 45–50 °C.
7. Collect slides and proceed to H&E (see step 4.9.1), immunohistochemical (see step 4.9.2), or immunofluorescent staining (see step 5).
   NOTE: Representative results are shown in Figure 3B.
   1. To perform H&E staining, obtain a kit from the materials list and follow the manufacturer’s specifications.
   2. To perform immunohistochemical staining, obtain a kit and primary antibody from the materials list and follow the manufacturer’s specifications.

5. Immunofluorescent Staining of FFPE and Sectioned Organoids

1. Collect supplies: baked slide from step 4, xylene, 100% EtOH, 95% EtOH, 70% EtOH, deionized water (DI H₂O), coplin jars, 1x antigen retrieval solution (sodium citrate buffer or EDTA-Tris buffer), phosphate-buffered saline (PBS), 5% normal horse serum (NHS) with 0.1% non-ionic detergent in PBS, 1% bovine serum albumin (BSA) with 0.3% non-ionic detergent in PBS, staining rack, staining dish, decloaking chamber, hydrophobic barrier pen, humidity chamber, 1° and 2° antibodies of interest, and counterstains of interest.
2. Deparaffinize and rehydrate the slides by dipping into coplins filled with the following solutions: xylene (3 dips, 5 min each), 100% EtOH (2 dips, 3 min each), 95% EtOH (2 dips, 3 min each), 70% EtOH (2 dips, 3 min each), and DI H₂O (2 dips, 3 min each).
3. Fill the staining dish with antigen retrieval solution and pre-heat decloaking chamber to 100 °C.
4. Perform antigen retrieval by immersing slides into the pre-heated staining dish and incubate for 5–10 min at 100°C. Once the cycle is complete, allow the decloaking chamber to sit for 20 min before opening the lid.
5. Once the slide dish has cooled to RT, place the staining dish under running DI H₂O to rinse the slides for 5 min.
6. Remove the slides from the staining dish, draw a circle around the sample with a hydrophobic barrier pen, and lay the slide on the humidity chamber.
7. Block any non-specific antibody staining by applying 5% NHS with 0.1% non-ionic detergent in PBS to the hydrophobic circle around the sample (about 30 μL is needed to cover the sample).
8. Wash slides in coplins filled with PBS 3 times for 5 min each.
9. Add the 1° antibodies (Table of Materials) diluted in 1% BSA with 0.3% non-ionic detergent in PBS to the hydrophobic circle around the sample (about 30 μL should cover the section), then incubate for 1 h at RT or overnight at 4 °C in humidity chamber.
10. Wash the slides in coplins filled with PBS 3 times for 5 min each.
11. Add the 2° antibody diluted in 1% BSA with 0.3% non-ionic detergent in PBS to the hydrophobic circle around the sample (about 30 μL will cover the area), then incubate for 1 h at RT in humidity chamber.
12. Wash the slides in coplin filled with PBS 3 times for 5 min each.
13. Add DAPI, diluted to the manufacturer’s specifications, in 1% BSA with 0.3% non-ionic detergent in PBS to the hydrophobic circle around the sample, then incubate for 2–5 min at RT in the dark (30 μL).
14. Wash the slides in coplins filled with PBS 3 times for 5 min each.
15. Mount the slides with antifade mounting media and coverslip, then visualize the results under a microscope.

6. Whole-mounting of Organoids for Immunofluorescent Staining

1. Collect supplies: chamber slide or confocal dish, phosphate-buffered saline (PBS), fixative, 50 mM NH₄Cl in PBS, 0.1% non-ionic detergent in PBS, 5% NHS with 0.1% non-ionic detergent in PBS, 1% BSA with 0.3% non-ionic detergent in PBS, 1° and 2° antibodies of interest, and counterstains of interest.
2. Carefully remove as much media as possible without disrupting the organoid culture by pipetting against the side of well, leaving space between media and matrix gel.
3. Using a trimmed, pre-wet with media or PBS, 1,000 μL pipette tip, draw up one drop (25–50 μL) of matrix gel that contains the organoid(s) of interest and dispense onto the middle of a chamber slide well or confocal dish.
   NOTE: 33% matrix gel is not a solid. It is a gelatinous fluid that handles similar to a Jello that has not fully set. A trimmed pipette tip with a large opening will prevent organoids from breaking during transfers.
4. If organoids remain in the parent culture, replace media with warm KSFM-based media.
5. With the naked eye or dissecting scope, aspirate the excess media and matrix gel from the chamber slide with a fine-tip pipette (10–200 μL).
   NOTE: It is optional to pretreat the chamber slide with an adherence reagent.
6. Plate an equal volume of matrix gel into an empty well of the chamber slide as an optional control to observe autofluorescence of leftover matrix.
7. Place the chamber slide in a 37 °C incubator for 30–45 min to promote the organoid to adhere to the glass and prevent loss of samples during wash steps.
8. Pipetting slowly to prevent detachment from slide, wash organoids with 200 μL of 1x PBS for 5 min at RT while gently shaking.
9. Remove the 1x PBS and fix with 200 μL of 4% paraformaldehyde for 30 min at RT while gently shaking. Ensure that the matrix gel appears clear after this step.
   NOTE: Methanol or formalin fixation are also possible. Fixation method should be optimized for specific antibodies as certain fixation methods may crosslink the antigens of interest or quench fluorescent-labeling-proteins of interest.
10. Remove the fixative and wash twice with 200 μL of 1x PBS for 5 min while gently shaking.
    NOTE: The length of washing steps should be optimized depending on organoid size. Five minutes is a sufficient starting point, but the washing step can be lengthened as needed.
11. Quench autofluorescence with 200 μL of 50 mM NH₄Cl in 1x PBS for 30 min at RT while gently shaking.
    NOTE: This step will quench autofluorescence from debris in the lumen of the organoid and from fluorescent protein expression (such as GFP) that may present from experimental design. It should be included if it is desired to use a fluorophore in the 488 channel but can be skipped if desired to preserve GFP signal.
12. Remove NH₄Cl and wash twice with 200 μL of 1x PBS for 5 min while gently shaking.
13. Permeabilize organoids in 200 μL of 0.1% non-ionic detergent-100 in 1x PBS for 30 min at RT while gently shaking.
14. Remove the 0.1% non-ionic detergent-100 in 1x PBS and wash twice with 200 μL of 1x PBS for 5 min while gently shaking.
15. Block with 5% NHS with 200 μL of 0.1% non-ionic detergent X-100 in PBS and incubate for 60 min at RT while gently shaking.
16. Remove the blocking buffer and wash twice with 200 μL of 1x PBS for 5 min while gently shaking.
17. Dilute the 1° antibody in 1% BSA with 0.3% Triton X-100 in 1x PBS and add to 200 μL of the chamber slide, then incubate for 1–3 days at 4 °C.
18. Remove the 1° antibody and wash twice with 200 μL of 1x PBS for 5 min while gently shaking.
19. Dilute the 2° antibody in 1% BSA with 0.3% Triton X-100 in 1x PBS and add to 200 μL of the chamber slide, then incubate for 1–3 days at 4 °C in the dark.
    NOTE: The length of incubation times should be optimized for organoid size and antibody. Some will require more time to penetrate through the organoid. Antibody concentration will also need to be optimized. In general, 2x the concentration used for FFPE sections is appropriate for 1° antibodies and the same concentration for FFPE sections is appropriate for 2° antibodies.
20. Remove the 2° antibody and wash twice with 200 μL of 1x PBS for 5 min while gently shaking.
21. Counterstain the actin with phalloidin and the nucleus with DAPI or Hoescht, diluted according to the manufacturer’s recommendations in 1% BSA with 0.3% Triton-X in 1x PBS. Add 200 μL to chamber slide, then incubate at RT for 40 min in the dark.
22. Mount in 200 μL of fresh 1x PBS or mounting media and image immediately (fluorescence should be preserved for up to 5 days, if not longer). Representative results are shown in Figure 3C.
    NOTE: Sodium azide can be added to samples to prevent contamination if confocal imaging is not readily available. Adding a clearing agent can improve imaging.

7. Whole-mounting of Organoids for Assays and Fluorescent Probes

NOTE: There are commercially available assays and fluorescent probes/dyes (examples are included on the materials list) amendable for use on whole-mounted organoids to observe a variety of useful endpoints¹⁹. The following protocol is for the fluorescently-labelled EdU proliferation kit, however any kit can be modified for use with a whole-mounted sample.

1. Carefully remove 50 μL of media and replace with 50 μL of 20 μM 2x working EdU solution by pipetting against the side of well, leaving space between media and matrix gel.
2. Incubate at 4 °C for 1–3 days (the desired length of time for EdU incorporation should be optimized for cell type of choice).
3. Follow steps 6.2–6.8 to transfer the organoids of interest onto a chamber slide or confocal dish.
4. Remove PBS and fix with 200 μL of 3.7% paraformaldehyde for 30 min at RT while gently shaking. Ensure that the matrix gel appear clear after this step.
5. Remove the fixative and wash twice with 200 μL of 1x PBS for 5 min while gently shaking. Remove PBS and add 200 μL of 0.5% non-ionic detergent-100 in 1x PBS for 30 min at RT while gently shaking.
6. Prepare a 1x fluorescent reaction cocktail according to the manufacturer’s specifications.
7. Remove 0.5% non-ionic detergent-100 and wash twice with 200 μL of 3% BSA in 1x PBS for 5 min while gently shaking.
8. Add reaction cocktail and incubate for 30 min at RT in the dark.
9. Remove the fluorescent reaction cocktail and wash once with 200 μL of 3% BSA in 1x PBS for 5 min while gently shaking. Counterstain and mount by following steps 6.21–6.22 (Figure 3D).