Prostate organoids are an exciting in vitro system for studying development and disease. They overcome challenges of immortalized cell lines and offer an inexpensive alternative to animal models. They can be grown from human prostate cells, as we've shown in this protocol, as well as mouse prostate cells, as a disease-relevant pre-clinical model system.
Many labs have described useful culture methods and endpoints. But we hope to bring those details together in one protocol and demonstrate the particularly challenging steps for new users. As the techniques improve, prostate cancer organoids can be grown from patient samples as a precision medicine approach to model disease and response to therapies.
The culture conditions are specific to prostate organoids, however many of the endpoints have been adapted from other tissue types. A user could adapt portions of this protocol to their cell type of interest. Start this experiment by growing epithelial cells in matrix gel-coded 96-well plates as described in the manuscript.
To collect organoids from the plates after 12 days, remove medium from the wells without disturbing the matrix gel. Then immediately add approximately 200 microliters of neutral protease to each well at about a one-to-two ratio of matrix gel to neutral protease. Incubate the plate for 20 minutes at 37 degrees Celsius.
Then pipette up and down to mechanically dissociate the mixture, and incubate again for 20 minutes at 37 degrees Celsius. After the incubation, transfer the neutral protease matrix gel-cell mixture from a few wells to a microcentrifuge tube. Pellet the cells by centrifugation at 300 times g for three to five minutes.
Remove the supernatant, and save the organoid pellet for subsequent analyses. To create molds for embedding, cut out a small section of a 1, 000-microliter pipette tip to make a cylinder that holds 50 microliters. Then from the wider part of the same tip, make a second wider mold that fits around the first mold.
To create a cold block, wrap an icepack with tape with the adhesive side up. Then adhere the molds perpendicularly to the tape. To embed the organoids in a histology gel plug for processing, first wash the previously-obtained organoid pellet with one milliliter of Hank's Balanced Salt Solution.
After centrifuging at 300 times g for three to five minutes, remove the supernatant. Then add 30 to 50 microliters of liquid histology gel to the organoid pellet, and mix gently by slowly pipetting up and down to resuspend the pellet. Transfer this mixture into a mold on the cold block, and allow the specimen to cool and solidify into a plug.
Next, use a plunger to push the plug out of the small mold and into the center of the larger mold, and fill it by pipetting clear 2%agar around the plug. To mark the top of the sample, pipette histologic dye-tinted 2%agar on top of the plug. Once the agar has cooled, use a plunger to transfer the plug into a histology cassette.
Label the cassette with a pencil and place it into 10%neutral buffered formalin for overnight fixation. On the following day, transfer the cassette from 10%neutral buffered formalin to 70%histological grade ethanol, and process it further and embed in paraffin. Begin by trimming the base of the block at a 10-micrometer thickness.
Once a section that contains the plug area appears, stop trimming. Lock the microtome rotor, and transfer the block back onto the ice to chill. Move the blade to a new, unused portion.
And transfer the block back to the clamp. Unlock the machine and begin trimming at five micrometers per section. Use tweezers to transfer the section to the 40-degree Celsius water bath, and allow it to spread out.
Dip a slide vertically in the water to transfer the section onto it. Then observe the slide under a bright field microscope to ensure that an organoid is present in the section. Bake overnight at 45 to 50 degrees Celsius, and then proceed with staining as described in the manuscript.
Start this part of the experiment with unused wells of the 96-well matrix gel-coated plate with organoids. By pipetting against the side of the well, leaving space between media and matrix gel, carefully remove as much media as possible without disrupting the organoid culture. Use a trimmed 1, 000-microliter pipette tip, pre-wet with PBS to draw up one drop of matrix gel that contains the organoids of interest.
Dispense the drop onto the middle of a chamber slide. With the naked eye, aspirate the excess media and matrix gel from the chamber slide using a fine-tipped pipette. To promote organoid adherence to the glass and prevent loss of samples during subsequent washing steps.
Place the slide in a 37-degree-Celsius incubator for 30 to 45 minutes. Pipetting slowly to prevent detachment from the dish, add 200 microliters of 1X PBS to wash the organoids for five minutes at room temperature with gentle shaking. Carefully remove the 1X PBS and add 200 microliters of 4%paraformaldehyde.
Fix for 30 minutes at room temperature with gentle shaking. At PFA, wash and proceed with subsequent permeabilization and staining steps as described by this protocol. Mount and image immediately.
Unstained freshly-cut slide clearly shows human primary prostate organoids, whereas in unstained dry slide, organoids appear translucent and are harder to detect under a bright field microscope. Hematoxylin and eosin staining shows a broken human primary prostate organoid from mishandling, such as aggressive pipetting, wrong processing protocol compared to a well-handled organoid. Human primary prostate organoid was formalin-fixed, paraffin-embedded, sectioned, and stained with basal cytokeratin 5 and luminal cytokeratin 8, basal p63 and luminal cytokeratin 8, and imaged with confocal microscope.
A whole-mounted human primary prostate organoid was stained with basal cytokeratin 5 or luminal cytokeratin 8, and counterstained with phalloidin and DAPI, and imaged with confocal microscope. A whole-mounted human primary prostate cell organoid was stained with fluorescently-labeled Edu and counterstained with Hoescht, and imaged with confocal microscope. It's important to visualize your organoids throughout the procedure.
After collecting the sections, make sure to visualize the samples on the slide, and make sure that you observe organoids during whole-mounting. After collecting the organoids from the matrix gel, you can whole-mount them and use commercial assays, or dissociate to single cells for flow cytometry or single-cell sequencing. 3D culture provides researchers with a novel way to study tissue in the setting of a lab dish.
It can be applied to organogenesis, or to understanding the pathology of a patient's disease. When working with patient samples, always exercise caution and handle the materials as potential for blood-borne pathogen exposure.
