We thank the UIC Biorespository members, Dr. Klara Valyi-Nagy, and Alex Susma, as well as the urologists, Drs. Michael Abern, Daniel Moreira, and Simone Crivallero, for facilitation of tissue acquisition for the primary cell cultures. We thank the UIC Urology patients for donating their tissue to research. This work was funded, in part, by the Department of Defense Prostate Cancer Research Program Health Disparities Idea Award PC121923 (Nonn) and the UIC Center for Clinical and Translation Science Pre-doctoral Education for Clinical and Translational Scientists (PECTS) Program (McCray and Richards) and by the National Institutes of Health&#39;s National Cancer Institute, Grant Numbers U54CA202995, U54CA202997, and U54CA203000, known as the Chicago Health Equity Collaborative (Nonn and Richards). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or the Department of Defense.

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The authors have nothing to disclose.

Organotypic culture is an exciting new method for recapitulating tissue with the convenience of an in vitro environment. Currently, labs grow organoids from many types of tissues for various endpoints. The methods described in this paper summarize useful endpoints and highlight new techniques to fully characterize 3D primary prostate cell cultures.
There are a variety of media recipes for cultivating human primary prostate cell organoids5,6,13. While all achieve viable and comparable results, KSFM-based media13 uses minimal additives so it is described here. Additionally, papers have been published using multiple concentrations for matrix gel, from 10% to 75% to culture prostate organoids5,6,13. Because matrix gel is an expensive reagent, and 33% matrix gel has been shown to be sufficient to cultivate long-term (2-3 weeks), viable, unclumped organoids13, this is the recommended concentration. However, matrix gel can vary in protein concentration between lot numbers, so this should be taken into account when plating. It is also recommended to purchase matrix gel in bulk for use across multiple experiments to reduce inconsistency between lots. Other labs have published different formats for plating matrix gel, such as droplets instead of coating a 96-well plate well5. Both methods form viable organoids, but the format described here allows for cells to be plated more sparsely, which has been shown to promote expansion of cells into larger organoids13. However, plating density should be optimized based off patient-specific formation efficiency. Matrix gel is available in many formats including growth factor-reduced, phenol red-free, high concentration, etc. Growth factor-reduced is recommended for defined culture conditions, and phenol red-free is recommended when working with cells that express GFP or in experiments that involve z-stack image capture. It is critical that any matrix gel plating steps be performed with ice-cold reagents, as matrix gel solidifies quickly at room temperature.
Organoids grown under different experimental treatments may result in changes to shape, so bright-field imaging is widely used to study observed morphological phenotypes. Nevertheless, recording and quantifying area or shape is a challenge for two reasons: 1) selection bias during image capture and 2) applying a two-dimensional parameter such as area to a three-dimensional sample. One strategy of image capture is to record a random field and measure a predetermined number of organoids in that field, but this can create bias during selection, and all organoids in the selected field may not be in focus. Whole-well imaging optimized for the 96-well microplate format eliminates this sampling bias by collecting the entire organoid population of interest. However, depending on the microscope objectives on hand, multiple fields may need to be collected and tiled in order to obtain a whole-well image. Some labs have described measuring area from a single z-plane12, but to capture all organoids in focus, it is recommended to collect and stack multiple planes into a single EDF-image13. In some cases, a meniscus in the matrix gel may cause vignetting in the image, and background correction methods may need to be applied using image analysis software. Exact volume is ideally the most appropriate measurement for organoid size; however, this is difficult to precisely obtain, even with multiple z-stack images. Other useful morphological dimensions, such as circularity and maximum/minimum ratio, can easily be calculated from all the organoids in a whole-well EDF image. Together, these methods overcome sampling bias challenges and enable measurement of 2D parameters from 3D objects in a 3D space.
Formalin fixation and paraffin embedding of organoids is a common method to obtain hematoxylin and eosin (H&#38;E), immunohistochemical (IHC), and immunofluorescent (IF) staining for visualization and analysis6,11,20. However, publications that have described this technique lack comprehensive details on the embedding process. Additionally, locating organoids within the paraffin block can be challenging. Some labs pre-stain organoids with trypan blue prior to embedding to aid in location during sectioning11. The method described here incorporates the use of a histological dye for orientation of the organoid/histology gelplug during embedding to promote efficiency of sectioning. H&#38;E slides facilitate examination of necrosis, nuclear texture, and proliferation, and thus it should be a mandatory endpoint for organoid culture to ensure that cells are healthy throughout the sphere. A strength of organoid culture is that samples are comprised of a heterogeneous mixture of cells that better resemble in vivo patient tissue. In the prostate, for example, both basal and luminal cells are present in prostate organoids5, while cells grown in 2D lack luminal differentiation8. To assess organoid differentiation, it is recommended that researchers assess basal markers such as CK5 and p63 and luminal markers such as androgen receptors and CK88. Any other experimental proteins of interest can also be studied using these staining techniques.
Although FFPE sections allow visualization of cells residing in the inner compartment via histologic methods (H&#38;E, IF, IHC), images are limited to cross sections, and organoid shape may be altered by the embedding process. Whole-mount staining enables an organoid to be stained and observed in situ, preserving morphological phenotypes and permitting images of protein localization. Whole-mounting is a tool utilized to stain whole-tissue or whole-animal specimens, such as the zebrafish or mouse embryo, and is easily adapted for organoids. The technique described here was modified from the procedure by Mah&#233;et al.11, which details the initial growth and culture of gastrointestinal organoids directly on a chamber slide for staining, and requires fixation of the entire parent culture. The method outlined here involves the transfer of a single organoid (or organoids) of interest onto a chamber slide at the time of staining. This enables the selection of individual organoids for whole-mount analysis in an ongoing experiment without fixation of the parent culture. When performing whole-mount staining, it is necessary to optimize permeabilization and the incubation time for primary and secondary antibodies to ensure penetration throughout the specimen (anywhere from one or more days is recommended). Once imaged via confocal microscopy, 3D renderings can be produced to enable visualization and localization of a specific protein and calculate the number of positively stained cells present in a sample. Flow cytometry is a useful endpoint to quantify populations of basal, luminal, or stem cells5,14. To do this, cells are recovered from the matrix gel and gently dissociated for staining. Users should carefully select appropriate markers for separation based on the current literature. For human primary prostate epithelial cells, CD26 and CD49f are suitable luminal and basal markers, respectively5,21.
In summary, this protocol details human primary prostate organoid growth, collection, and experimental endpoints. Of note, the mounting technique described can be applied to a variety of other assays that are normally employed for 2D cells, such as fluorescent probe-based experiments looking at proliferation19, apoptosis, and subcellular organelle stains. Additionally, the collection and dissociation method described here could be utilized in preparation for single-cell RNA sequencing18. Collectively, this demonstrates the possibility for a variety of novel, high-fidelity endpoints that researchers can optimize and explore in the future.

Organoids are a valuable tool to study organogenesis and disease. They provide a less expensive alternative to animal models, and patient-derived organoids are evolving as a strategy for personalized medicine1,2,3. This three-dimensional (3D) culture system involves seeding stem or progenitor cells (harvested from tissue or induced pluripotent stem cells) into a gel of extracellular matrix components (matrix gel)4. The cells proliferate and differentiate, resulting in organotypic structures that recapitulate the cellular hierarchy and morphology of the organ of interest&#39;s functional unit. Organoids have been grown using cells originating from a variety of organs, including the salivary gland, stomach, intestine, liver, prostate, lung, and brain4. Although there are numerous protocols describing steps to establish prostate organoids5,6, it is challenging to find sufficiently detailed methods on how to achieve quantitative endpoints from organoids. This paper summarizes methods developed for human primary prostate cells and details a series of suggested endpoints to assess the organoid phenotypes. These techniques were optimized for prostate organoids and may be applicable to other 3D cell cultures.
Prostate organoid cultures have recently emerged as a valuable in vitro model that lacks the limitations of monolayer cultures using established immortalized cell lines. Benign prostate epithelium and prostate cancer is challenging to model in vitro using immortalized cell lines. The number of benign cell lines is limited, and all have undergone transformation with oncogenes7. Primary prostate epithelial cells in monolayers do not differentiate into luminal cells and lack androgen receptors8. The majority of prostate cancer cell lines do not have functional androgen receptors, a significant mediator of early disease state, and lack key genetic changes that have been found in patient tumors9. Prostate organoid cultures can be grown easily from benign epithelium and are valuable in studying stem cell properties, progenitor cells, differentiation, and effects of experimental changes in the microenvironment4,5. Prostate cancer organoids can be grown as part of a precision medicine approach to modeling patient disease and response to therapies10.
This protocol has been compiled from existing protocols that use various cell types, but it has been optimized here for use in human primary prostate cells. It compliments protocols outlined by Sawyers, Clevers, and Shen5,6 that describe growth, passaging, bright-field imaging, cryopreservation, and RNA and DNA isolation of prostate mouse and human organoids. The whole-mount protocol is modified from Mah&#233;&#160;et al.11, who used gastrointestinal epithelial organoids and describes live imaging and frozen and paraffin embedding. Borten et al.12 described the analysis of breast, colon, and colorectal cancer organoid morphology from bright-field imaging. Additionally, Richards et al.13 used a method for the morphologic assessment of prostate organoids. Finally, Hu et al.14 described a method of adhering prostate organoids to a chamber slide overnight prior to the immunofluorescent staining to observe single cells and dispersion of spheres for flow cytometry analysis.
The goal of this protocol is to demonstrate in sufficient detail these technically challenging methods as one protocol, including cell seeding; media and matrix gel maintenance; collection of cells for flow cytometry; RNA, DNA, and protein extraction; morphologic assessment from bright-field z-stack analysis; embedding, processing and sectioning for histological staining; and whole-mounting for immunofluorescence or fluorescent probe assays. The biological relevance and interpretation of these various endpoints will vary among experimental design and antibodies used for analysis. Through utilization of this protocol, users should feel prepared to set up an experiment with a toolkit of endpoints.
Human primary prostate epithelial (PrE) cells were established in the lab by the protocol described by Peehl15,16 and maintained to a limited number of passages (up to four) as previously described17, but they are also available from commercial vendors. Hepatocyte serum-free media-based6, KSFM-based13, and R-spondin 1-conditioned5 media are all published as having successfully produced organoids. KSFM-based requires the fewest number of additives, so it is described here.

<table><tbody><tr><td>Cells of interest</td><td></td><td></td><td></td></tr><tr><td>Keratinocyte-SFM (KSFM)</td><td>ThermoFisher Scientific</td><td>17005042</td><td></td></tr><tr><td>Fetal Bovine Serum, charcoal stripped, USDA-approved regions (FBS)</td><td>ThermoFisher Scientific</td><td>12676029</td><td></td></tr><tr><td>5&amp;alpha;-Dihydrotestosterone (DHT)</td><td>Sigma-Aldrich</td><td>D-073-1ML</td><td></td></tr><tr><td>Flat bottom, polystyrene 96-well cell culture treated plate</td><td>ThermoFisher Scientific</td><td>161093</td><td></td></tr><tr><td>Matrigel (matrix gel)</td><td>Corning</td><td>Various</td><td>Matrix-gel: growth Factor Reduced, Phenol-red free, etc. depending on application</td></tr><tr><td>Ice Bucket</td><td></td><td></td><td></td></tr><tr><td>Cell Culture Hood</td><td></td><td></td><td></td></tr><tr><td>1.5 mL micro-centrifuge tubes or 15 mL conical</td><td>ThermoFisher Scientific</td><td>05-408-130, 339650</td><td></td></tr><tr><td>Centrifuge</td><td></td><td></td><td></td></tr><tr><td>Dispase (neutral protease)</td><td>STEMCELL Technologies</td><td>7923</td><td>neutral protease</td></tr><tr><td>Hanks&#039;&amp;nbsp;balanced salt solution&amp;nbsp;(HBSS)</td><td>ThermoFisher Scientific</td><td>14025076</td><td></td></tr><tr><td>TrypLE Express (cell dissociation enzymes)</td><td>ThermoFisher Scientific</td><td>12605036</td><td>Cell dissociation enzymes (trypsin may also work, depending on cell type, but TrypLE is more gentle and recommended for primary cells)</td></tr><tr><td>TRIzol</td><td>ThermoFisher Scientific</td><td>15596018</td><td>suggested RNA extraction solution</td></tr><tr><td>RIPA Lysis and Extraction Buffer</td><td>ThermoFisher Scientific</td><td>89900</td><td>suggested protein extraction solution</td></tr><tr><td>DNAzol</td><td>ThermoFisher Scientific</td><td>10503027</td><td>suggested DNA extraction solution</td></tr><tr><td>Organoids</td><td></td><td></td><td></td></tr><tr><td>Flat bottom, polystyrene 96-well cell culture treated plate</td><td>ThermoFisher Scientific</td><td>161093</td><td></td></tr><tr><td>Brightfield microscope with camera capabilities</td><td></td><td></td><td>ex. &amp;lrm;EVOS FL Auto Imaging System</td></tr><tr><td>Photo analysis software</td><td></td><td></td><td>ex. Photoshop, CELLESTE, ImageJ, MorphoLibJ, CellProfiler</td></tr><tr><td>Graphing software</td><td></td><td></td><td>ex. Graphpad, Excel, etc</td></tr><tr><td>Organoids</td><td></td><td></td><td></td></tr><tr><td>Ice pack</td><td></td><td></td><td></td></tr><tr><td>Masking tape</td><td></td><td></td><td></td></tr><tr><td>Pipette tips (1000 &amp;mu;L)</td><td></td><td></td><td></td></tr><tr><td>Razor blade</td><td></td><td></td><td></td></tr><tr><td>Dispase</td><td>STEMCELL Technologies</td><td>7923</td><td></td></tr><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A9045</td><td></td></tr><tr><td>HistoGel (histology-gel)</td><td>ThermoFisher Scientific</td><td>HG-4000-012</td><td></td></tr><tr><td>Pencil</td><td></td><td></td><td></td></tr><tr><td>&quot;Plunger&quot; from 1 cc insulin syringe</td><td></td><td></td><td></td></tr><tr><td>Tissue Casette</td><td>Thomas Scientific</td><td>1202D72</td><td></td></tr><tr><td>Container to hold fixative</td><td></td><td></td><td></td></tr><tr><td>10% neutral buffered formalin (NBF)</td><td>Sigma-Aldrich</td><td>HT501128</td><td></td></tr><tr><td>Histology pen, xylene and EtOH-resistant</td><td>Sigma-Aldrich</td><td>Z648191-12EA</td><td>STATMARK pen</td></tr><tr><td>Ethanol (histologic grade)</td><td>Fisher Scientific</td><td>A405P-4</td><td>For fixation and graded dilutions during processing</td></tr><tr><td>Xylenes</td><td>Sigma-Aldrich</td><td>214736-1L</td><td></td></tr><tr><td>Deionized water</td><td></td><td></td><td></td></tr><tr><td>Paraffin</td><td>Leica</td><td>various</td><td></td></tr><tr><td>Tissue processor</td><td></td><td></td><td></td></tr><tr><td>Embedding workstation</td><td></td><td></td><td></td></tr><tr><td>Economy Tissue Float Bath</td><td>Daigger</td><td>EF4575E XH-1001</td><td></td></tr><tr><td>Microtome</td><td></td><td></td><td></td></tr><tr><td>Microtome blades</td><td>Ted Pella</td><td>27243</td><td></td></tr><tr><td>Positively charged microscope slides</td><td>Thomas Scientific</td><td>1158B91</td><td></td></tr><tr><td>Laboratory Oven</td><td>ThermoFisher Scientific</td><td>PR305225G</td><td></td></tr><tr><td>Hematoxylin and Eosin Stain Kit</td><td>Vector Laboratories</td><td>H-3502</td><td>Suggested H&amp;amp;E staining kit</td></tr><tr><td>ABC Peroxidase Standard Staining Kit</td><td>ThermoFisher Scientific</td><td>32020</td><td>Suggested immunohistochemistry staining kit</td></tr><tr><td>Androgen Receptor Primary Antibody (AR)</td><td>Cell Signaling Technology</td><td>5153S</td><td>Suggested primary antibody for IHC</td></tr><tr><td>FFPE, sectioned organoid sample baked on a slide</td><td></td><td></td><td></td></tr><tr><td>Ethanol (histologic grade)</td><td>Fisher Scientific</td><td>A405P-4</td><td>dilutions should be performed using deinoized water</td></tr><tr><td>Xylenes</td><td>Sigma-Aldrich</td><td>214736-1L</td><td></td></tr><tr><td>Deionized water (DI H&lt;sub&gt;2&lt;/sub&gt;O)</td><td></td><td></td><td></td></tr><tr><td>Antigen retrieval solution</td><td>Sigma-Aldrich, Abcam</td><td>C9999, ab93684</td><td></td></tr><tr><td>1x Phosphate Buffered Saline - PBS</td><td>ThermoFisher Scientific</td><td>10010023</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100-1L</td><td></td></tr><tr><td>Normal Horse Serum</td><td>thermoFisher Scientific</td><td>31874</td><td></td></tr><tr><td>Bovin Serum Albumin</td><td>Sigma-Aldrich</td><td>A2058</td><td></td></tr><tr><td>Counterstain (DAPI, Hoescht)</td><td>Sigma-Aldrich</td><td>D9542</td><td></td></tr><tr><td>Sodium azide</td><td>Sigma-Aldrich</td><td>S2002</td><td>suggested for storage but not required</td></tr><tr><td>Confocal microscope</td><td></td><td></td><td></td></tr><tr><td>Coplin</td><td>Fisher Scientific</td><td>19-4</td><td></td></tr><tr><td>Staining rack</td><td>IHC World</td><td>M905-12DGY</td><td></td></tr><tr><td>Staining dish</td><td>IHC World</td><td>M900-12B</td><td></td></tr><tr><td>Decloaking chamber</td><td>Biocare Medical</td><td>DC2012</td><td></td></tr><tr><td>Humidity chamber</td><td>Thomas Scientific</td><td>1219D68</td><td></td></tr><tr><td>Hydrophobic barrier pen</td><td>Vector Laboratories</td><td>H-4000</td><td></td></tr><tr><td>Cytokeratin 8/18 primary antibody (CK8)</td><td>ARP American Research Products</td><td>03-GP11</td><td>suggested primary antibody for IF</td></tr><tr><td>p63-alpha antibody (p63)</td><td>Cell Signaling Technology</td><td>4892S</td><td>suggested primary antibody for IF</td></tr><tr><td>Keratin 5 Polyclonal Antibody (CK5)</td><td>Biolegend</td><td>905501</td><td>suggested primary antibody for IF</td></tr><tr><td>Goat anti-rabbit secondary</td><td>ThermoFisher Scientific</td><td>A21245</td><td>suggested secondary antibody for p63 or CK5 detection (do not use at same time)</td></tr><tr><td>Goat anti-guinea pig secondary</td><td>ThermoFisher Scientific</td><td>A-11075</td><td>suggested secondary antibody for CK8 detection</td></tr><tr><td>Microscope cover glass</td><td>Globe Scientific</td><td>1414-10</td><td></td></tr><tr><td>Anti-fade mounting media</td><td>ThermoFisher Scientific</td><td>S36972</td><td></td></tr><tr><td>Organoids</td><td></td><td></td><td></td></tr><tr><td>Pipette tips (1000 &amp;mu;L)</td><td></td><td></td><td></td></tr><tr><td>Pipette tips (200 &amp;mu;L)</td><td></td><td></td><td></td></tr><tr><td>8-well chamber slide</td><td>ThermoFisher Scientific</td><td>154534PK</td><td>It is also possible to use a confocal dish, depends on preference of user</td></tr><tr><td>Cell-Tak</td><td>Corning</td><td>CB40240</td><td>optional adherent reagent</td></tr><tr><td>1x Phosphate Buffered Saline (PBS)</td><td>ThermoFisher Scientific</td><td>10010023</td><td></td></tr><tr><td>4% paraformaldehye (PFA)</td><td>Biotium</td><td>22023</td><td>other fixatives such as methanol or formalin can be used</td></tr><tr><td>50mM NH&lt;sub&gt;4&lt;/sub&gt;Cl</td><td>sigma-Aldrich</td><td>254134</td><td></td></tr><tr><td>Triton&amp;trade; X-100 (non-ionic detergent)</td><td>sigma-Aldrich</td><td>X100-1L</td><td></td></tr><tr><td>Normal Horse Serum (NHS)</td><td>thermoFisher Scientific</td><td>31874</td><td></td></tr><tr><td>Bovin Serum Albumin (BSA)</td><td>sigma-Aldrich</td><td>A2058</td><td></td></tr><tr><td>Counterstain (DAPI, Hoescht)</td><td>Sigma-Aldrich</td><td>D9542</td><td></td></tr><tr><td>Counterstain (phalloidin)</td><td>thermoFisher Scientific</td><td>A22287</td><td></td></tr><tr><td>Sodium azide</td><td>sigma-Aldrich</td><td>S2002</td><td>suggested for storage but not required</td></tr><tr><td>Confocal microscope</td><td></td><td></td><td></td></tr><tr><td>Cytokeratin 8/18 primary antibody (CK8)</td><td>ARP American Research Products</td><td>03-GP11</td><td>suggested primary antibody for IF</td></tr><tr><td>p63-alpha antibody (p63)</td><td>Cell Signaling Technology</td><td>4892S</td><td>suggested primary antibody for IF</td></tr><tr><td>Keratin 5 Polyclonal Antibody (CK5)</td><td>Biolegend</td><td>905501</td><td>suggested primary antibody for IF</td></tr><tr><td>Goat anti-rabbit secondary</td><td>ThermoFisher Scientific</td><td>A21245</td><td>suggested secondary antibody for p63 or CK5 detection (do not use at same time)</td></tr><tr><td>Goat anti-guinea pig secondary</td><td>ThermoFisher Scientific</td><td>A-11075</td><td>suggested secondary antibody for CK8 detection</td></tr><tr><td>Visikol HISTO-M</td><td>Visikol</td><td>various</td><td>optional clearing agent</td></tr><tr><td>Organoids</td><td></td><td></td><td></td></tr><tr><td>Pipette tips (1000 &amp;mu;L)</td><td></td><td></td><td></td></tr><tr><td>Pipette tips (200 &amp;mu;L)</td><td></td><td></td><td></td></tr><tr><td>8-well chamber slide</td><td>ThermoFisher Scientific</td><td>154534PK</td><td>It is also possible to use a confocal dish, depends on preference of user</td></tr><tr><td>Cell-Tak</td><td>Corning</td><td>CB40240</td><td></td></tr><tr><td>1x Phosphate Buffered Saline - PBS</td><td>ThermoFisher Scientific</td><td>10010023</td><td></td></tr><tr><td>Cell assay of interest</td><td>Various</td><td>Various</td><td>Click-iT EdU Alexa Fluor proliferation assay (fluorescently-labelled EdU proliferation kit), Image-iT Lipid Peroxidation Kit, etc) and fluorescent probes/dyes (ex. HCS mitochondrial Health Kit, CellMask, LIVE/DEAD Viability assays, CellROX reagents, etc)</td></tr><tr><td>Organoids</td><td></td><td></td><td></td></tr><tr><td>Ice bucket</td><td></td><td></td><td></td></tr><tr><td>Cell culture hood</td><td></td><td></td><td></td></tr><tr><td>1.5 mL eppendorf tubes or 15 mL conical</td><td></td><td></td><td></td></tr><tr><td>Microcentrifuge</td><td></td><td></td><td></td></tr><tr><td>Dispase</td><td>STEMCELL Technologies</td><td>7923</td><td></td></tr><tr><td>TrypLE Express (cell dissociation enzymes)</td><td>ThermoFisher Scientific</td><td>12605036</td><td>Cell dissociation enzymes (trypsin may also work, depending on cell type, but TrypLE is more gentle and recommended for primary cells)</td></tr><tr><td>Hanks&#039;&amp;nbsp;balanced salt solution&amp;nbsp;(HBSS)</td><td>ThermoFisher Scientific</td><td>14175079</td><td></td></tr><tr><td>Flow Tube with Cell Strainer Snap Cap</td><td>Fisher Scientific</td><td>08-771-23</td><td></td></tr><tr><td>Cytokeratin 5 Antibody - FITC</td><td>Millipore Sigma</td><td>FCMAB291F</td><td>Suggested flow antibody</td></tr><tr><td>Cytokeratin 8 Antibody - Alexafluor 405</td><td>Abcam</td><td>ab210139</td><td>Suggested flow antibody</td></tr><tr><td>CD49f - Alexafluor 647</td><td>BioLegend</td><td>313609</td><td>Suggested flow antibody</td></tr><tr><td>CD26 - PE</td><td>BioLegend</td><td>320576</td><td>Suggested flow antibody</td></tr><tr><td>Flow cytometer</td><td></td><td></td><td></td></tr></tbody></table>

handling and assessment of human primary prostate organoid culture

This paper describes a detailed protocol for three-dimensional (3D) culturing, handling, and evaluation of human primary prostate organoids. The process involves seeding of epithelial cells sparsely in a 3D matrix gel on a 96-well microplate with media changes to cultivate expansion into organoids. Morphology is then assessed by whole-well capturing of z-stack images. Compression of z-stacks creates a single in-focus image from which organoids are measured to quantify a variety of outputs, including circularity, roundness, and area.DNA, RNA, and protein can be collected from organoids recovered from the matrix gel. Cell populations of interest can be assessed by organoid dissociation and flow cytometry. Formalin-fixation-paraffin-embedding (FFPE) followed by sectioning is used for the histological assessment and antibody staining. Whole-mount immunofluorescent staining preserves organoid morphology and facilitates observation of protein localization in organoids in situ. Commercial assays that are traditionally used for 2D monolayer cells can be modified for 3D organoids. Used together, the techniques in this protocol provide a robust toolbox to quantify prostate organoid growth, morphologic characteristics, and expression of differentiation markers.

Upon successful culture of human primary prostate organoids, morphology and differentiation can be assessed using bright-field image analysis and FFPE and whole-mount staining techniques.
The process of bright-field image capture is illustrated in Figure 1A. Organoids are grown in a 3D matrix and dispersed across different focal planes. To observe organoids in focus across many planes, it is suggested to use an EDF image (Figure 1B, right) instead of a single z-plane (Figure 1B, left). In the lab, organoids grown under different experimental conditions may result in changes in morphology. Using area, circularity (defined by how similar the organoid is to a circle) and max/min radius (measure of length) give users an indication of organoid size and shape and provide a quantifiable readout of morphology. To emphasize the usefulness of these measures, representative images of organoids with similar area but differing morphology are shown in Figure 1C, in which a long organoid will be less circular and have a larger maximum/minimum ratio than a spherical organoid.
Embedding organoids for histology is a useful endpoint to observe the interior of samples and ensure that necrosis is not present within the core of an organoid. A workflow for this process and representative results of the unstained, sectioned samples under a bright-field microscope are provided in Figure 2A,B,C. Figure 3A shows a mishandled organoid (left) compared to properly handed organoids in Figure 3A (right) and Figure 3B. To determine the differentiation-state of the sample, it is recommended to look at basal cell markers (cytokeratin 5 and p63)8 along with luminal cell markers (cytokeratin 8)8. If it is desired to stain organoids in situ, whole-mount staining is a convenient alternative, and these representative results are provided in Figure 3C,D. 
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59051/59051fig1v2.jpg" /> 
Figure 1: Morphology assessment of organoids in 3D culture. (A) Image collection and compression workflow for human primary prostate organoids acquired by a transmitted light inverted microscope with a motorized X/Y scanning stage and companion software. (B) Representative images of a single z-stack (left) vs. an EDF image (right) of a whole-well sample of human primary prostate organoids, showing that more organoids are in focus when multiple z-stacks are combined into a single projected image (scale bar = 1000 &#181;m). (C) Representative images for area, circularity, and max/min ratio of morphologically dissimilar human primary prostate organoids which have the same area (scale bar = 200 &#181;m). <a href="https://www.jove.com/files/ftp_upload/59051/59051fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59051/59051fig2.jpg" /> 
Figure 2: Organoid embedding workflow and representative sectioning results. (A) Human primary prostate organoid embedding workflow. (B) Representative image of an unstained, fresh slide containing human primary prostate organoids under a bright field microscope depicting agar (green arrow), histology gel (black arrow), bubble (blue arrow), organoids (red arrows) (scale bar = 1000 &#181;m). (C) Unstained freshly cut slide (left) vs. unstained dry slide (right), organoids (red arrows) appear translucent when dry (scale bar = 500 &#181;m) and are harder to discern under a bright-field microscope. <a href="https://www.jove.com/files/ftp_upload/59051/59051fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59051/59051fig3.jpg" /> 
Figure 3: Histological staining on sectioned and whole-mounted organoids. (A) H&#38;E staining of a broken human primary prostate organoid from mishandling (aggressive pipetting, wrong processing protocol, left) next to a well-handled organoid (right) (scale bar = 100 &#181;m). (B) Human primary prostate organoid that has been formalin-fixed, paraffin-embedded, sectioned, and stained with basal cytokeratin 5 or luminal cytokeratin 8 (top) or basal p63 and luminal cytokeratin 8 (bottom) and imaged with confocal microscope (scale bar = 100 &#181;m). (C) A whole-mounted human primary prostate organoid stained with basal cytokeratin 5 or luminal cytokeratin 8 and counterstained with phalloidin and DAPI, imaged with confocal microscope (scale bar = 100 &#181;m). (D) A whole-mounted human primary prostate cell organoid stained with fluorescently labeled EdU and counter stained with Hoescht, imaged with confocal microscope (scale bar = 500 &#181;m) <a href="https://www.jove.com/files/ftp_upload/59051/59051fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Handling and Assessment of Human Primary Prostate Organoid Culture at JoVE.com

Handling and Assessment of Human Primary Prostate Organoid Culture

Here, we present a protocol to guide human primary prostate organoid handling then suggest endpoints to assess phenotype. Seeding, culture maintenance, recovery from matrix gel, morphologic quantification, embedding and sectioning, FFPE sectioning, whole-mount staining, and application of commercial assays are described.

This paper describes a detailed protocol for three-dimensional (3D) culturing, handling, and evaluation of human primary prostate organoids. The process ...

handling-and-assessment-of-human-primary-prostate-organoid-culture

Research

JoVE Journal

Bioengineering

17.1K Views.  University of Illinois at Chicago. Prostaatorganoïden zijn een spannend in vitro systeem voor het bestuderen van ontwikkeling en ziekte. Ze overwinnen uitdagingen van vereeuwigde cellijnen en bieden een goedkoop alternatief voor diermodellen. Ze kunnen worden gekweekt uit menselijke prostaatcellen, zoals we hebben aangetoond in dit protocol, evenals muis prostaatcellen, als een ziekte-relevant pre-klinisch modelsysteem.Veel laboratoria hebben beschreven nuttige cultuur methoden en eindpunten. Maar we hopen deze details...

Video: Handling and Assessment of Human Primary Prostate Organoid Culture

A Rapid Filter Insert-based 3D Culture System for Primary Prostate Cell Differentiation

Conditionally reprogrammed cells (CRCs) provide a sustainable method for primary cell culture and the ability to develop extensive &#34;living biobanks&#34; of patient derived cell lines. For many types of epithelial cells, various three dimensional (3D) culture approaches have been described that support an improved differentiated state. While CRCs retain their lineage commitment to the tissue from which they are isolated, they fail to express many of the differentiation markers associated with the tissue of origin when grown under normal two dimensional (2D) culture conditions. To enhance the application of patient-derived CRCs for prostate cancer research, a 3D culture format has been defined that enables a rapid (2 weeks total) luminal cell differentiation in both normal and tumor-derived prostate epithelial cells. Herein, a filter insert-based format is described for the culturing and differentiation of both normal and malignant prostate CRCs. A detailed description of the procedures required for cell collection and processing for immunohistochemical and immunofluorescent staining are provided. Collectively the 3D culture format described, combined with the primary CRC lines, provides an important medium- to high- throughput model system for biospecimen-based prostate research.

Here, we present a method for the establishment of a rapid in vitro system that supports the three dimensional culturing and subsequent luminal differentiation of primary prostate epithelial cells.

Conditionally reprogrammed cells (CRCs) provide a sustainable method for primary cell culture and the ability to develop extensive &#34;living ...

Cancer Research

Rotating Cell Culture Systems for Human Cell Culture: Human Trophoblast Cells as a Model

The field of human trophoblast research aids in understanding the complex environment established during placentation. Due to the nature of these studies, human in vivo experimentation is impossible. A combination of primary cultures, explant cultures and trophoblast cell lines1 support our understanding of invasion of the uterine wall2 and remodeling of uterine spiral arteries3,4 by extravillous trophoblast cells (EVTs), which is required for successful establishment of pregnancy. Despite the wealth of knowledge gleaned from such models, it is accepted that in vitro cell culture models using EVT-like cell lines display altered cellular properties when compared to their in vivo counterparts5,6. Cells cultured in the rotating cell culture system (RCCS) display morphological, phenotypic, and functional properties of EVT-like cell lines that more closely mimic differentiating in utero EVTs, with increased expression of genes mediating invasion (e.g. matrix metalloproteinases (MMPs)) and trophoblast differentiation7,8,9. The Saint Georges Hospital Placental cell Line-4 (SGHPL-4) (kindly donated by Dr. Guy Whitley and Dr. Judith Cartwright) is an EVT-like cell line that was used for testing in the RCCS.
The design of the RCCS culture vessel is based on the principle that organs and tissues function in a three-dimensional (3-D) environment. Due to the dynamic culture conditions in the vessel, including conditions of physiologically relevant shear, cells grown in three dimensions form aggregates based on natural cellular affinities and differentiate into organotypic tissue-like assemblies10,11,12 . The maintenance of a fluid orbit provides a low-shear, low-turbulence environment similar to conditions found in vivo. Sedimentation of the cultured cells is countered by adjusting the rotation speed of the RCCS to ensure a constant free-fall of cells. Gas exchange occurs through a permeable hydrophobic membrane located on the back of the bioreactor. Like their parental tissue in vivo, RCCS-grown cells are able to respond to chemical and molecular gradients in three dimensions (i.e. at their apical, basal, and lateral surfaces) because they are cultured on the surface of porous microcarrier beads. When grown as two-dimensional monolayers on impermeable surfaces like plastic, cells are deprived of this important communication at their basal surface. Consequently, the spatial constraints imposed by the environment profoundly affect how cells sense and decode signals from the surrounding microenvironment, thus implying an important role for the 3-D milieu13.
We have used the RCCS to engineer biologically meaningful 3-D models of various human epithelial tissues7,14,15,16. Indeed, many previous reports have demonstrated that cells cultured in the RCCS can assume physiologically relevant phenotypes that have not been possible with other models10,17-21. In summary, culture in the RCCS represents an easy, reproducible, high-throughput platform that provides large numbers of differentiated cells that are amenable to a variety of experimental manipulations. 
In the following protocol, using EVTs as an example, we clearly describe the steps required to three-dimensionally culture adherent cells in the RCCS.

Traditional, two dimensional cell culture techniques often result in altered characteristics with respect to differentiation markers, cytokines and growth factors. Three-dimensional cell culture in the rotating cell culture system (RCCS) reestablishes expression of many of these factors as shown here with an extravillous trophoblast cell line.

The field of human trophoblast research aids in  understanding the complex environment established during placentation. Due to the  nature of these ...

Identification, Histological Characterization, and Dissection of Mouse Prostate Lobes for In Vitro 3D Spheroid Culture Models

Genetically engineered mouse models (GEMMs) serve as effective pre-clinical models for investigating most types of human cancers, including prostate cancer (PCa). Understanding the anatomy and histology of the mouse prostate is important for the efficient use and proper characterization of such animal models. The mouse prostate has four distinct pairs of lobes, each with their own characteristics. This article demonstrates the proper method of dissection and identification of mouse prostate lobes for disease analysis. Post-dissection, the prostate cells can be further cultured in vitro for mechanistic understanding. Since mouse prostate primary cells tend to lose their normal characteristics when cultured in vitro, we outline here a method for isolating the cells and growing them as 3D spheroid cultures, which is effective for preserving the physiological characteristics of the cells. These 3D cultures can be used for analyzing cell morphology and behavior in near-physiological conditions, investigating altered levels and localizations of key proteins and pathways involved in the development and progression of a disease, and looking at responses to drug treatments.

Genetically engineered mice are useful models for investigating prostate cancer mechanisms. Here we present a protocol to identify and dissect prostate lobes from a mouse urogenital system, differentiate them based on histology, and isolate and culture the primary prostate cells in vitro as spheroids for downstream analyses.

Genetically engineered mouse models (GEMMs) serve as effective pre-clinical models for investigating most types of human cancers, including prostate ...

Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) are a rigorous method for studying mammalian development and disease. Our laboratory has developed several GEMMs of prostate cancer (PCa) that lack expression of one or multiple tumor suppressor genes using the site-specific Cre-loxP recombinase system and a prostate-specific promoter. In this article, we describe our method for necropsy of these PCa GEMMs, primarily focusing on dissection of mouse prostate tumors. New methods developed over the last decade have facilitated the culture of epithelial-derived cells to model organ systems in vitro in three dimensions. We also detail a 3D cell culture method to generate tumor organoids from mouse PCa GEMMs. Pre-clinical cancer research has been dominated by 2D cell culture and cell line-derived or patient-derived xenograft models. These methods lack tumor microenvironment, a limitation of using these techniques in pre-clinical studies. GEMMs are more physiologically-relevant for understanding tumorigenesis and cancer progression. Tumor organoid culture is an in vitro model system that recapitulates tumor architecture and cell lineage characteristics. In addition, 3D cell culture methods allow for growth of normal cells for comparison to tumor cell cultures, rarely possible using 2D cell culture techniques. In combination, use of GEMMs and 3D cell culture in pre-clinical studies has the potential to improve our understanding of cancer biology.&#160;

We show a method for necropsy and dissection of mouse prostate cancer models, focusing on prostate tumor dissection. A step-by-step protocol for generation of mouse prostate tumor organoids is also presented.

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) ...

Genetics

Evaluating the Differentiation Capacity of Mouse Prostate Epithelial Cells Using Organoid Culture

The prostate epithelium is comprised predominantly of basal and luminal cells. In vivo lineage tracing has been utilized to define the differentiation capacity of mouse prostate basal and luminal cells during development, tissue-regeneration and transformation. However, evaluating cell-intrinsic and extrinsic regulators of prostate epithelial differentiation capacity using a lineage tracing approach often requires extensive breeding and can be cost-prohibitive. In the prostate organoid assay, basal and luminal cells generate prostatic epithelium ex vivo. Importantly, primary epithelial cells can be isolated from mice of any genetic background or mice treated with any number of small molecules prior to, or after, plating into three-dimensional (3D) culture. Sufficient material for evaluation of differentiation capacity is generated after 7-10 days. Collection of basal-derived and luminal-derived organoids for (1) protein analysis by Western blot and (2) immunohistochemical analysis of intact organoids by whole-mount confocal microscopy enables researchers to evaluate the ex vivo differentiation capacity of prostate epithelial cells. When used in combination, these two approaches provide complementary information about the differentiation capacity of prostate basal and luminal cells in response to genetic or pharmacological manipulation.

Mouse prostate organoids represent a promising context to evaluate mechanisms that regulate differentiation. This paper describes an improved approach to establish prostate organoids, and introduces methods to (1) collect protein lysate from organoids, and (2) fix and stain organoids for whole-mount confocal microscopy.

The prostate epithelium is comprised predominantly of basal and luminal cells. In vivo lineage tracing has been utilized to define the differentiation ...

Prostate Organoid Cultures as Tools to Translate Genotypes and Mutational Profiles to Pharmacological Responses

Presented here is a protocol to study pharmacodynamics, stem cell potential, and cancer differentiation in prostate epithelial organoids. Prostate organoids are androgen responsive, three-dimensional (3D) cultures grown in a defined medium that resembles the prostatic epithelium. Prostate organoids can be established from wild-type and genetically engineered mouse models, benign human tissue, and advanced prostate cancer. Importantly, patient derived organoids closely resemble tumors in genetics and in vivo tumor biology. Moreover, organoids can be genetically manipulated using CRISPR/Cas9 and shRNA systems. These controlled genetics make the organoid culture attractive as a platform for rapidly testing the effects of genotypes and mutational profiles on pharmacological responses. However, experimental protocols must be specifically adapted to the 3D nature of organoid cultures to obtain reproducible results. Described here are detailed protocols for performing seeding assays to determine organoid formation capacity. Subsequently, this report shows how to perform drug treatments and analyze pharmacological response via viability measurements, protein isolation, and RNA isolation. Finally, the protocol describes how to prepare organoids for xenografting and subsequent in vivo growth assays using subcutaneous grafting. These protocols yield highly reproducible data and are widely applicable to 3D culture systems.

Presented here is a protocol to study pharmacological responses in prostate epithelial organoids. Organoids closely resemble in vivo biology and recapitulate patient genetics, making them attractive model systems. Prostate organoids can be established from wildtype prostates, genetically engineered mouse models, benign human tissue, and advanced prostate cancer.

Presented here is a protocol to study pharmacodynamics, stem cell potential, and cancer differentiation in prostate epithelial organoids. Prostate ...

Establishment and Analysis of Three-Dimensional (3D) Organoids Derived from Patient Prostate Cancer Bone Metastasis Specimens and their Xenografts

Three-dimensional (3D) culture of organoids from tumor specimens of human patients and patient-derived xenograft (PDX) models of prostate cancer, referred to as patient-derived organoids (PDO), are an invaluable resource for studying the mechanism of tumorigenesis and metastasis of prostate cancer. Their main advantage is that they maintain the distinctive genomic and functional heterogeneity of the original tissue compared to conventional cell lines that do not. Furthermore, 3D cultures of PDO can be used to predict the effects of drug treatment on individual patients and are a step towards personalized medicine. Despite these advantages, few groups routinely use this method in part because of the extensive optimization of PDO culture conditions that may be required for different patient samples. We previously demonstrated that our prostate cancer bone metastasis PDX model, PCSD1, recapitulated the resistance of the donor patient&rsquo;s bone metastasis to anti-androgen therapy. We used PCSD1 3D organoids to characterize further the mechanisms of anti-androgen resistance. Following an overview of currently published studies of PDX and PDO models, we describe a step-by-step protocol for 3D culture of PDO using domed or floating basement membrane (e.g., Matrigel) spheres in optimized culture conditions. In vivo stitch imaging and cell processing for histology are also described. This protocol can be further optimized for other applications including western blot, co-culture, etc. and can be used to explore characteristics of 3D cultured PDO pertaining to drug resistance, tumorigenesis, metastasis and therapeutics.

Establishment and Analysis of Three-Dimensional 3D Organoids Derived from Patient Prostate Cancer Bone Metastasis Specimens and their Xenografts

Three-dimensional cultures of patient BMPC specimens and xenografts of bone metastatic prostate cancer maintain the functional heterogeneity of their original tumors resulting in cysts, spheroids and complex, tumor-like organoids. This manuscript provides an optimization strategy and protocol for 3D culture of heterogeneous patient derived samples and their analysis using IFC.

Three-dimensional (3D) culture of organoids from tumor specimens of human patients and patient-derived xenograft (PDX) models of prostate cancer, referred ...

Culture of Bladder Cancer Organoids as Precision Medicine Tools

Current in vitro therapeutic testing platforms lack relevance to tumor pathophysiology, typically employing cancer cell lines established as two-dimensional (2D) cultures on tissue culture plastic. There is a critical need for more representative models of tumor complexity that can accurately predict therapeutic response and sensitivity. The development of three-dimensional (3D) ex vivo culture of patient-derived organoids (PDOs), derived from fresh tumor tissues, aims to address these shortcomings. Organoid cultures can be used as tumor surrogates in parallel to routine clinical management to inform therapeutic decisions by identifying potential effective interventions and indicating therapies that may be futile. Here, this procedure aims to describe strategies and a detailed step-by-step protocol to establish bladder cancer PDOs from fresh, viable clinical tissue. Our well-established, optimized protocols are practical to set up 3D cultures for experiments using limited and diverse starting material directly from patients or patient-derived xenograft (PDX) tumor material. This procedure can also be employed by most laboratories equipped with standard tissue culture equipment. The organoids generated using this protocol can be used as ex vivo surrogates to understand both the molecular mechanisms underpinning urological cancer pathology and to evaluate treatments to inform clinical management.

Patient-derived organoids (PDOs) are a powerful tool in translational cancer research, reflecting both the genetic and phenotypic heterogeneity of the disease and response to personalized anti-cancer therapies. Here, a consolidated protocol to generate human primary bladder cancer PDOs in preparation for the evaluation of phenotypic analyses and drug responses is detailed.

Current in vitro therapeutic testing platforms lack relevance to tumor pathophysiology, typically employing cancer cell lines established as ...

Prostate Organoid Assay: A Matrix Gel Ring-based Ex Vivo Culture Technique to Study the Differentiation Capacity of Prostate Epithelial Cells

Source: Crowell, P. D. et al. <a href="https://www.jove.com/video/60223" target="_blank">Evaluating the Differentiation Capacity of Mouse Prostate Epithelial Cells Using Organoid Culture.</a> J. Vis. Exp. (2019)
In this video, we demonstrate the culturing of isolated mouse prostate epithelial cells using a matrix gel-ring based culture technique. This approach can provide complementary information about the differentiation capacity of prostate epithelial cells to form 3D glandular structures in response to genetic or pharmacological manipulation.

Source: Crowell, P. D. et al. Evaluating the Differentiation Capacity of Mouse Prostate Epithelial Cells Using Organoid Culture. J. Vis. Exp. (2019)
In ...

JoVE Encyclopedia of Experiments

Prostate Cancer

Assays and techniques

Preparing Organoids from Tumors: A 3D In Vitro Tumor Model

Source: Wadosky, K. M., et al. <a href="https://www.jove.com/video/59710/" target="_blank">Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer</a>. J. Vis. Exp. (2019).
Organoids derived from tumors are a type of 3D cell culture model that can maintain the pathological features and organ-specific cell lineage of tumors. In the example protocol, we will see scientists generate organoids from a prostate tumor from a genetically engineered mouse model.

Source: Wadosky, K. M., et al. Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer. J. Vis. Exp. (2019).
Organoids ...

Handling and Assessment of Human Primary Prostate Organoid Culture

Samenvatting

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