PDC and JMG are supported by the Ruth L. Kirschstein National Research Service Award GM007185. JAD is supported by the National Institute of General Medical Sciences of the National Institutes of Health (R25GM055052) awarded to T. Hasson and the Saul Martinez Scholarship. ASG is supported by the Spitzer Family Foundation Fund and the Gill Endowment. This work was supported by the American Cancer Society (RSG-17-068-01-TBG), Department of Defense (W81XWH-13-1-0470), Margaret E. Early Medical Research Trust, NIH/NCI (P50CA092131/UCLA SPORE in Prostate Cancer), Rose Hills Foundation, and support from UCLA&#39;s Jonsson Comprehensive Cancer Center, Broad Stem Cell Research Center, Clinical and Translational Science Institute, and Institute of Urologic Oncology.

All methods described here have been approved by the Institutional Review Board at the University of California, Los Angeles.
NOTE: A schematic illustrating the approaches described in the paper is provided in Figure 1.
1. Isolating Mouse Basal and Luminal Prostate Epithelial Cells Using Fluorescence-activated Cell Sorting (FACS) &#8212; TIMING: 30 Min
NOTE: Perform steps 1.3-1.5 in the dark.
<ol>
	<li>After dissociating cells from total mouse prostate as described in Lawson et al.16, transfer the cells to FACS tubes and resuspend 0.1-5 x 106 cells in 100 &#181;L of dissociation media (Table 1).</li>
	<li>Add the appropriate volume of the following directly-conjugated primary antibodies: CD45, CD31, Ter-119, EpCAM and CD49f.</li>
	<li>Incubate on ice, protected from light, for 20 min. 
	NOTE: It is recommended to utilize 10% of the total dissociated cells for unstained and single-stained controls. These controls are necessary to set the correct compensation and voltage for sorting.</li>
	<li>Quench antibody cocktail by adding 1 mL of dissociation media to each sample. Pellet the cells by centrifugation at 800 x g for 5 min at room temperature (RT) and remove the supernatant by aspirating.</li>
	<li>Resuspend the cells in appropriate volume (250 &#181;L per 1 x 106 cells) of dissociation media containing 1 &#181;g/mL 4&#39;,6-diamidino-2-phenylindole (DAPI). Proceed to FACS. Flow cytometry plots demonstrating isolation of mouse basal and luminal prostate epithelial cells are illustrated in Figure 2.</li>
</ol>
2. Plating Sorted Prostate Epithelial Cells into Primary Mouse Organoid Culture &#8212; TIMING: 2-3 H (Excluding Poly-HEMA-coated Plate Preparation)
NOTE: Plates are coated with Poly-HEMA to prevent 2D colony formation on the surface of the well beneath the matrix gel. Prepare Poly-HEMA-coated plates 1 day prior to plating sorted basal or luminal prostate epithelial cells into mouse organoid culture. Thaw 1 mL aliquots of reduced growth factor matrix gel, hereafter referred to as matrix gel, on ice 2 h prior to step 2.1. Y-27632 (ROCK inhibitor) should be added to mouse organoid media immediately prior to step 2.1. Perform steps 2.1-2.8 on ice.
<ol>
	<li>Pellet the cells in 5 mL round-bottom tubes by centrifugation at 800 x g for 5 min at 4 &#176;C and aspirate the supernatant.</li>
	<li>Wash the cell pellet in 500 &#181;L of mouse organoid media (Table 2)14.</li>
	<li>Pellet the cells by centrifugation at 800 x g for 5 min at 4 &#176;C and aspirate the supernatant.</li>
	<li>Resuspend in mouse organoid media at a cell density of 1,000 cells/&#181;L.</li>
	<li>To prepare master mixes, mix epithelial cells suspended in mouse organoid media with matrix gel to generate a final mixture that contains 25% cells/media and 75% matrix gel. Basal cells are typically plated at a concentration of 100-2,000 cells/80 &#181;L, whereas luminal cells are typically plated at a concentration of 2,000-10,000 cells/80 &#181;L. The density of cells plated varies depending upon the day of anticipated material collection, and the desired downstream application. 
	NOTE: Chill appropriately sized tube(s) for expected master mix volume 5 min prior to master mix preparation. To ensure the matrix gel does not harden while handling, it is critical to chill the pipette tip by pipetting the matrix gel 3-4 times prior to transferring it to a new tube.</li>
	<li>Add 80 &#181;L of the matrix gel/cell mixture per well of a 24-well plate. Pipetting a droplet onto the lower half of the wall of the well, while avoiding direct contact with the Poly-HEMA coating is recommended. After adding the matrix gel, swirl the plate to allow the matrix gel/cell mixture to form a ring around the rim of the well.</li>
	<li>Place the 24-well plate into a 37 &#176;C 5% CO2 incubator right-side up for 10 min to allow the matrix gel to partially harden. 
	NOTE: Begin warming mouse organoid media at 37 &#176;C immediately after placing the 24-well plate in the incubator.</li>
	<li>After incubating for 10 min, flip the 24-well plate upside-down and incubate for an additional 50 min to allow the matrix gel to completely harden.</li>
	<li>Add 350 &#181;L of pre-warmed mouse organoid media dropwise to the center of each well. 
	NOTE: To maintain the integrity of the matrix gel, it is critical to avoid the matrix gel ring while adding media.</li>
	<li>After adding the media, return the 24-well plate to the 37 &#176;C 5% CO2 incubator.</li>
</ol>
3. Replenishing Mouse Organoid Media &#8212; TIMING: 10-15 Min Per 24-well Plate
NOTE: Existing media should be replaced with fresh media every 48 h. Before each media change, pre-warm mouse organoid media. It is not necessary to add ROCK inhibitor to the media used for replenishing.
<ol>
	<li>Tilt the 24-well plate at a 45&#176; angle and gently remove existing media from the center of each well using a p1000 pipette, while avoiding the matrix gel ring.</li>
	<li>Add 350 &#181;L of pre-warmed mouse organoid media as in step 2.9. It is recommended to add a larger volume of media (up to 1 mL) to organoids cultured for longer than 5 days in order to prevent rapid depletion of key nutrients and growth factors.</li>
</ol>
4. Extracting Protein Lysate from Prostate Organoids for Western Blot Analysis &#8212; TIMING: 2.5-4 H
NOTE: Prior to collecting organoids for protein lysate extraction, prepare and pre-warm dispase-containing media (Table 1).
<ol>
	<li>Remove the media from each well as in step 3.1.</li>
	<li>To collect organoids, repeatedly blast the matrix gel by pipetting 1 mL of dispase-containing media directly onto the matrix gel ring until the entire ring is dislodged, and transfer to a 1.5 mL microcentrifuge tube. 
	NOTE: It is critical to avoid direct contact with the Poly-HEMA-coated wells. Direct contact may cause contamination of the collected material with Poly-HEMA, which could negatively impact cell survival.</li>
	<li>Place the 1.5 mL microcentrifuge tube(s) into a 37 &#176;C 5% CO2 incubator for 30 min to 1 h to allow complete digestion of the matrix gel by dispase.</li>
	<li>Pellet organoids by centrifugation at 800 x g for 5 min at RT and remove the supernatant using a micropipette.</li>
	<li>Add phosphate-buffered saline (PBS) to the organoid pellet and resuspend by gently flicking. 
	NOTE: Failure to sufficiently resuspend the organoid pellet may result in the contamination of organoid material with residual dispase or matrix gel.</li>
	<li>Pellet the organoids by centrifugation at 800 x g for 5 min at RT and remove the supernatant using a micropipette.</li>
	<li>Fast freeze the organoid pellets by placing each tube into a solution containing dry ice and methanol. Store the tube(s) until future use at -80 &#176;C. Alternatively, extract protein lysate immediately following step 4.6.</li>
	<li>Resuspend the organoid pellets in 100 &#181;L of protein lysis buffer (Table 1) per 10 &#181;L of packed cell volume. Flick to resuspend. 
	NOTE: If resuming after fast-freezing, ensure protein lysis buffer is thawed prior to removing samples from -80 &#176;C, as lysis buffer must be added to samples immediately in order to prevent phosphatase and protease activity.</li>
	<li>Incubate the samples in protein lysis buffer on ice for at least 45 min. 
	NOTE: It is recommended to sonicate prior to incubation on ice to increase the efficiency of nuclear protein recovery; however, sonication is not required. If sonication is not performed, proceed to step 4.10.
	<ol>
		<li>To sonicate, submerge tubes in wet ice and gently apply the tip of the sonic dismembrator to the outside of the microcentrifuge tube. Sonicate for 40 s at 20 kHz.</li>
	</ol>
	</li>
	<li>Proceed to Western blot following established protocols.</li>
</ol>
5. Fixing and Staining Prostate Organoids for Immunohistochemical Analysis by Whole-mount Confocal Microscopy
<ol>
	<li>Collecting prostate organoids from 24-well plates &#8212; TIMING: 45-60 min 
	NOTE: When collecting prostate organoids to process for confocal microscopy, it is critical to handle them with care in order to maintain their structure. The collection protocol below is designed to reduce disruption of organoid structure during isolation.
	<ol>
		<li>Remove the media from each well as in step 3.1.</li>
		<li>Digest the matrix gel by incubating with 500 &#181;L of dispase-containing media (Table 1) for 30 min in a 37 &#176;C 5% CO2 incubator.</li>
		<li>Collect digested organoid suspension in a microcentrifuge tube and pellet the organoids by centrifugation at 800 x g for 3 min at RT. Remove the supernatant.</li>
	</ol>
	</li>
	<li>Whole-mount immunofluorescent staining of prostate organoids &#8212; TIMING: 3-4 days (1-5 h/day)
	<ol>
		<li>Add 500 &#181;L of 4% paraformaldehyde in PBS and incubate for 2 h at RT with gentle shaking.</li>
		<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT, remove the supernatant, and wash the pellet with 1 mL of PBS for 15 min with gentle shaking.</li>
		<li>Wash the pellet as in step 5.2.2 for additional two times.</li>
		<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant. Add 1 &#181;g/mL DAPI in blocking solution (Table 1). Incubate for 2 h at RT or alternatively overnight at 4 &#176;C with gentle shaking.</li>
		<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant. Add primary antibody (rabbit anti-p63, mouse anti-cytokeratin 8) in blocking solution and incubate overnight at 4 &#176;C with gentle shaking.</li>
		<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant. Wash the pellet with 1 mL of PBS for 15 min with gentle shaking.</li>
		<li>Wash the pellet as in step 5.2.6 for additional two times.</li>
		<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant. Add secondary antibody (goat anti-rabbit IgG-Alexa Fluor 594, goat anti-mouse IgG-Alexa Fluor 488) in blocking solution and incubate overnight at 4 &#176;C with gentle shaking.</li>
		<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT, remove the supernatant, and wash the pellet with 1 mL of PBS for 15 min with gentle shaking.</li>
		<li>Wash the pellet as in step 5.2.9 for additional two times.</li>
	</ol>
	</li>
</ol>
6. Tissue Clearing and Mounting of the Stained Prostate Organoids for Whole-mount Confocal Microscopy &#8212; TIMING: 7 H
<ol>
	<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant.</li>
	<li>Add 1 mL of 30% sucrose in PBS with 1% Triton X-100 and incubate for 2 h at RT with gentle shaking.</li>
	<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant.</li>
	<li>Add 1 mL of 45% sucrose in PBS with 1% Triton X-100 and incubate for 2 h at RT with gentle shaking.</li>
	<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove the supernatant.</li>
	<li>Add 1 mL of 60% sucrose in PBS with 1% Triton X-100 and incubate for 2 h at RT with gentle shaking.</li>
	<li>Pellet the organoids by centrifugation at 800 x g for 3 min at RT and remove 95% of the supernatant. 
	NOTE: The pellet becomes looser as the concentration of sucrose becomes higher. Observing the DAPI-stained organoids under the UV light to confirm that they were not lost during removal of the supernatant is recommended.</li>
	<li>Transfer a 10-20 &#181;L droplet of the remaining suspension to a chambered coverslip and proceed to confocal microscopy. 
	NOTE: Coverslip fragments can be placed on either side of the droplet to be used as spacers (Figure 4C). These prevent organoids from collapsing when a coverslip is placed over the droplet.</li>
</ol>

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The authors have nothing to disclose.

Prostate epithelial cell differentiation has been implicated in both normal prostate biology2,3,4,5,6,7 and disease biology8,10,11,12; however, the master regulators of this process remain undefined. Identifying key regulators of prostate epithelial cell differentiation has been difficult in part due to the absence of well-established contexts to model it. While 2D monolayer culture can be used to model differentiation11,12, this context fails to recapitulate the complex prostate microenvironment. Furthermore, in vivo contexts to model differentiation do not lend themselves to mechanistic studies, as they are challenging to manipulate. Therefore, the identification of an easy to manipulate, yet physiologically-relevant context, to study differentiation is critical.
The prostate organoid model represents an elegant ex vivo context where basal to luminal differentiation is reported to occur. Methods to establish prostate organoids are well established14; however, further optimization of these methods is necessary. Furthermore, approaches to harvest and prepare prostate organoids for analysis are not clearly described. This paper describes an approach to plate prostate epithelial cells isolated from mouse prostate into organoid culture. This approach allows researchers to (1) prevent the occurrence of 2D colonies during organoid formation, (2) reduce the risk of disruption to the matrix gel during media replenishment, and (3) count organoids more effectively. In addition, this manuscript outlines approaches to harvest organoids for preparation for Western blot analysis, or whole-mount confocal microscopy. Importantly, the approach utilized to prepare organoids for confocal microscopy maintains the intact structure of organoids through its duration, which reduces organoid damage prior to image acquisition. Altogether, the approaches described expand the capabilities of the prostate organoid assay.
Notably, the organoid-forming capacity of basal and luminal cells can be altered both by methods used to isolate the respective populations, and by culture conditions. The organoid culture conditions used in this assay were first described by Karthaus et al.13. Whereas Karthaus et al. have reported that basal cells have a higher organoid forming capacity (15%) than luminal cells (1%)13, Chua et al., using distinct isolation methods and culture conditions, have reported that luminal cells (0.2-0.3%) have a higher organoid-forming capacity than basal cells (0.03%)20. Overall, methods described by Karthaus et al. lead to higher organoid-forming rates for both basal and luminal cells, likely reflecting differences in the approach used to isolate basal and luminal cells13, as opposed to culture conditions that bias against organoid formation from luminal cells. It remains unclear whether the protocol described in this manuscript favors luminal organoid formation from multipotent luminal progenitors, or committed-luminal progenitors9. Though timely and cost-prohibitive, in vivo lineage tracing studies can be used to validate progenitor features associated with distinct prostate epithelial lineages elucidated in the organoid assay.
Processes such as development, differentiation and transformation are not only relevant to prostate biology, but also relevant to the biology of other tissues including the brain, lung, intestine, pancreas and liver. The methods described facilitate the utilization of the organoid model to study these processes in not only the prostate, but also a wide range of tissues.

Basal and luminal cells comprise the majority of the prostate epithelium1. Lineage tracing studies have revealed that these cell types are predominantly self-sustained by distinct progenitors in the adult mouse2; however, luminal differentiation from basal progenitors has been observed in several contexts including development3,4, tissue regeneration5, inflammation6,7 and prostate cancer initiation2,8. Furthermore, emerging data supports the existence of multipotent luminal progenitors as well as luminal-committed progenitors9. In metastatic prostate cancer, differentiation from an AR-dependent luminal lineage to an AR-indifferent lineage with basal and neuroendocrine features represents an increasingly appreciated mechanism of resistance to androgen pathway inhibitors10,11,12. Therefore, as differentiation is implicated in normal physiology, cancer initiation and resistance to therapy, elucidating key molecular regulators of prostate epithelial cell differentiation is critical.
The mouse prostate organoid model has emerged as an elegant ex vivo context to study prostate epithelial cell differentiation9,13,14. In this assay, individual epithelial cells are plated into a 3D matrix where they generate glandular structures containing both basal and luminal cells within 1 week. While existing approaches for plating cells into organoid culture can be used to efficiently generate organoids, these approaches require further optimization14. Notable challenges associated with culturing prostate organoids include (1) excluding two-dimensional (2D) colonies that form beneath the Matrigel (matrix gel) from analysis, (2) maintaining the integrity of the matrix gel during media changes, and (3) counting organoids accurately. This paper outlines an approach to generate organoids from epithelial cells isolated from mouse prostate. The approach described entails coating plates with poly(2-hydroxyethyl methacrylate) (Poly-HEMA) to prevent the occurrence of 2D colonies. Furthermore, cells are plated into a matrix gel ring, rather than a matrix gel disc, which makes changing the media and counting organoids less challenging. These techniques allow researchers to more easily investigate how genetic alterations or small molecules introduced prior to, or during, organoid formation alter key processes such as differentiation.
Harvesting of prostate organoids for Western blot or immunohistochemical analysis by whole-mount confocal microscopy can provide valuable mechanistic insight into differentiation13, yet well-established protocols to prepare organoids for such techniques are lacking. This manuscript describes approaches to harvest organoids for (1) collection of protein lysate, or (2) fixation and staining for confocal microscopy. Importantly, the approach described for fixing and staining prostate organoids is considerably improved in relation to existing methods. While these rely on sectioning organoids15, the method described in this manuscript utilizes intact organoids, which helps protect against organoid damage during sample preparation. When used in combination, Western blot and confocal microscopy can provide valuable insight into the molecular regulators of differentiation. Alternatively, these approaches can be used to model other processes such as development and transformation.

<table><tbody><tr><td>&amp;micro;-Dish 35 mm, high</td><td>ibidi</td><td>81156</td><td></td></tr><tr><td>16% Paraformaldehyde</td><td>Thermo Fisher Scientific</td><td>50-980-487</td><td></td></tr><tr><td>4&amp;rsquo;,6-diamidino-2-phenylindole (DAPI)</td><td>Thermo Fisher Scientific</td><td>D1306</td><td></td></tr><tr><td>A83-01</td><td>Tocris</td><td>2939</td><td></td></tr><tr><td>Advanced DMEM/F-12</td><td>Thermo Fisher Scientific</td><td>12634010</td><td></td></tr><tr><td>APC/Cy7 anti-mouse CD326 (Ep-CAM) Antibody, 100 &amp;mu;g</td><td>BioLegend</td><td>118218</td><td></td></tr><tr><td>B-27 Supplement (50x), Serum Free</td><td>Thermo Fisher Scientific</td><td>17504044</td><td></td></tr><tr><td>Complete Protease Inhibitor Cocktail</td><td>Sigma</td><td>11836145001</td><td></td></tr><tr><td>(DiHydro)testosterone (5&amp;alpha;-Androstan-17&amp;beta;-ol-3-one)</td><td>Sigma</td><td>A-8380</td><td></td></tr><tr><td>Dispase II, Powder</td><td>Thermo Fisher Scientific</td><td>17-105-041</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Sigma</td><td>F8667</td><td></td></tr><tr><td>FITC anti-mouse CD31 Antibody (0.5 mg/mL, 50 &amp;mu;g)</td><td>BioLegend</td><td>102405</td><td></td></tr><tr><td>FITC anti-mouse CD45 Antibody (0.5 mg/mL, 50 &amp;mu;g)</td><td>BioLegend</td><td>103107</td><td></td></tr><tr><td>FITC anti-mouse TER-119/Erythroid Cells Antibody (0.5 mg/mL, 50 &amp;mu;g)</td><td>BioLegend</td><td>116205</td><td></td></tr><tr><td>Goat anti-mouse IgG-Alexa Fluor 488</td><td>Invitrogen</td><td>A28175</td><td></td></tr><tr><td>Goat anti-rabbit IgG-Alexa Fluor 594</td><td>Invitrogen</td><td>A11012</td><td></td></tr><tr><td>GlutaMAX</td><td>Thermo Fisher Scientific</td><td>35050061</td><td></td></tr><tr><td>Halt Phosphatase Inhibitor</td><td>Thermo Fisher Scientific</td><td>78428</td><td></td></tr><tr><td>Matrigel GFR Membrane Matrix</td><td>Corning</td><td>CB-40230C</td><td></td></tr><tr><td>Mouse anti-cytokeratin 8</td><td>BioLegend</td><td>904804</td><td></td></tr><tr><td>N-acetyl-L-cysteine</td><td>Sigma</td><td>A9165</td><td></td></tr><tr><td>Normocin</td><td>Thermo Fisher Scientific</td><td>ant-nr-1</td><td></td></tr><tr><td>PE anti-human/mouse CD49f Antibody</td><td>BioLegend</td><td>313612</td><td></td></tr><tr><td>Penicillin-Streptomycin (10,000 U/mL)</td><td>Thermo Fisher Scientific</td><td>15-140-122</td><td></td></tr><tr><td>Poly(2-hydroxyethyl methacrylate) (Poly-HEMA)</td><td>Sigma</td><td>P3932-25G</td><td></td></tr><tr><td>Rabbit anti-p63</td><td>BioLegend</td><td>619002</td><td></td></tr><tr><td>Radioimmunoprecipitation assay (RIPA)</td><td>Thermo Fisher Scientific</td><td>PI89901</td><td></td></tr><tr><td>Recombinant Human EGF, Animal-Free</td><td>PeproTech</td><td>AF-100-15</td><td></td></tr><tr><td>Recombinant Human Noggin</td><td>PeproTech</td><td>120-10C</td><td></td></tr><tr><td>RPMI 1640 Medium, HEPES (cs of 10)</td><td>Thermo Fisher Scientific</td><td>22400105</td><td></td></tr><tr><td>Sonic Dismembrator</td><td>Thermo Fisher Scientific</td><td>FB120</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S0389-500G</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>X100-5ML</td><td></td></tr><tr><td>Y-27632 dihydrochloride (ROCK inhibitor)</td><td>Selleck Chemical</td><td>S1049-50MG</td><td></td></tr></tbody></table>

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.

Prostate epithelial cells are plated into mouse organoid culture where they form organoids, which are harvested prior to preparation for downstream analysis (Figure 1).
Basal and luminal epithelial cells are isolated using FACS. After excluding DAPI+ cells and depleting Lin+ cells (CD45, CD31, Ter119), basal and luminal cells are distinguished based on differential expression of EpCAM and CD49f (Figure 2). The approach described to plate prostate basal and luminal cells into organoid culture entails: (1) plating cells into matrix gel rings, and (2) coating wells with Poly-HEMA. Plating into rings has been previously described in Agarwal et al9. Utilizing this approach (Figure 3A) allows researchers to more easily avoid the matrix gel while replenishing the media (Step 3), and more easily count organoids by following the circumference of the well. Coating wells with Poly-HEMA has been shown to prevent 2D colony formation in retinal organoids17; however, this approach has not been utilized in the prostate organoid model. Importantly, coating wells with Poly-HEMA (Table 3) eliminates the occurrence of 2D colonies without interfering with organoid formation (Figure 3B). These modifications expand the capabilities of the prostate organoid assay.
Basal and luminal cells form organoids with distinct morphologies (Figure 4A). While most basal-derived organoids are similar in size (100-300 &#181;m diameter) after 7 days in culture, luminal-derived organoids exhibit significant heterogeneity (30-450 &#181;m diameter). Furthermore, most basal-derived organoids contain lumens surrounded by multi-layered epithelium (Figure 4A, top), whereas luminal-derived organoids range in morphology from hollow, with single-layered epithelium to solid, with multi-layered cords of cells that do not canalize (Figure 4A, bottom). The approaches described above to prepare organoids for downstream analysis (Steps 4, 5), were used to investigate whether these phenotypic differences are reflective of differences in lineage marker expression. Western blot analysis revealed that basal and luminal-derived organoids retain features associated with basal and luminal primary cells. Basal-derived organoids express higher levels of the basal marker cytokeratin 5 (K5), whereas luminal-derived organoids express higher levels of the luminal marker cytokeratin 8 (K8) (Figure 4B). Both basal and luminal markers were detected in basal and luminal-derived organoids in the bulk population, perhaps suggestive of differentiation (Figure 4B).
We sought to characterize lineage marker expression in basal-derived organoids and determine whether morphologically distinct luminal-derived organoids exhibit differences in marker expression by staining intact organoids and performing confocal microscopy (Figure 4D). Basal-derived organoids contained multi-layered epithelium with outer layers expressing high levels of the basal marker p63 and moderate levels of the luminal marker K8 (p63hi, K8mid), and inner layers without detectable levels of p63 and high levels of K8 (p63lo, K8hi) (Figure 4D, top). While all cells in single-layered luminal-derived organoids stained positively for K8, only select cells contained nuclear p63 (Figure 4D, bottom). These data validate the approaches to harvest and prepare organoids for analysis by Western blot or confocal microscopy and thereby expand the capability of the organoid assay to study key cellular processes, including differentiation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60223/60223fig1.jpg" /> 
Figure 1: Schematic illustrating workflow to generate prostate organoids for collection and analysis. Total mouse prostate is dissociated and basal and luminal prostate epithelial cells are isolated by fluorescence-activated cell sorting via established protocols18,19. Basal or luminal cells suspended in a mixture of mouse organoid media and matrix gel are plated into matrix gel rings. After 5 to 7 days of culture, organoids are harvested for analysis by Western blot or confocal microscopy. <a href="https://www.jove.com/files/ftp_upload/60223/60223fig1large.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/60223/60223fig2.jpg" /> 
Figure 2: Isolation of mouse basal and luminal prostate epithelial cells using fluorescence-activated cell sorting (FACS). Dissociated cells from mouse prostate are stained with DAPI, to distinguish live from dead cells, and surface antibodies, to distinguish basal from luminal cells, prior to FACS. Left = Gated on DAPI- cells. FSC-A = forward-scatter. Center = Gated on Lin- cells (CD45lo, CD31lo, Ter119lo). SSC-A = side-scatter. Right = Basal cells (Bas) (EpCAMhi, CD49fhi), Luminal cells (Lum) (EpCAMhi, CD49fmid). <a href="https://www.jove.com/files/ftp_upload/60223/60223fig2large.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/60223/60223fig3.jpg" /> 
Figure 3: Establishment of mouse prostate organoids. (A) Schematic illustrating approach to generate a matrix gel ring in a well of a 24-well plate. (B) Representative phase contrast images of organoids (3D growth plane) and two-dimensional colonies (2D growth plane) formed 7 days after plating prostate epithelial cells into un-coated (Poly-HEMA (-)), or coated (Poly-HEMA (+)) 24-well plates. Boxed regions within 2D growth plane are magnified on the right. Scale bars = 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/60223/60223fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60223/60223fig4.jpg" /> 
Figure 4: Analysis of lineage marker expression in prostate organoids by Western blot and whole-mount confocal microscopy. (A) Representative phase contrast images of basal-derived (top), and luminal-derived (bottom) organoids after 7 days of culture. Scale bar = 100 &#181;m. (B) Western blot analysis of basal-derived (Bas) and luminal-derived (Lum) organoids after 5 days of culture. Staining for the basal marker, cytokeratin 5 (K5), and the luminal marker, cytokeratin 8 (K8), and a loading control, histone H3 (HH3). (C) Schematic illustrating chambered coverslip with spacers. (D) Representative differential interference contrast (DIC) and immunofluorescent images of basal-derived (top) and luminal-derived (bottom) organoids after 7 days of culture. Staining for p63 (red), K8 (green) and DAPI (blue) individually and merged. Scale bars = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/60223/60223fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td colspan="2">Recipes</td>
		</tr>
		<tr>
			<td>Dispase-containing media</td>
			<td>1 mg/mL dispase + 10 &#181;M ROCK inhibitor in advanced DMEM F12. Filter sterilize using 0.22 &#181;m filter.</td>
		</tr>
		<tr>
			<td>Dissociation media</td>
			<td>10% FBS + 1x Penicillin-Streptomycin in RPMI 1640. Filter sterilize using 0.22 &#181;m filter.</td>
		</tr>
		<tr>
			<td>Protein lysis buffer</td>
			<td>RIPA buffer + phosphatase inhibitors + protease inhibitors</td>
		</tr>
		<tr>
			<td>Blocking solution</td>
			<td>10% FBS in PBS with 0.2% Triton X-100</td>
		</tr>
	</tbody>
</table>
Table 1: Instructions for the preparation of key solutions.
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>B-27</td>
			<td>1x (dilute from 50x concentrate)</td>
		</tr>
		<tr>
			<td>GlutaMAX</td>
			<td>1x (dilute from 100x concentrate)</td>
		</tr>
		<tr>
			<td>N-acetyl-L-cysteine</td>
			<td>1.25 mM</td>
		</tr>
		<tr>
			<td>Normocin</td>
			<td>50 &#181;g/mL</td>
		</tr>
		<tr>
			<td>Recombinant Human EGF, Animal-Free</td>
			<td>50 ng/mL</td>
		</tr>
		<tr>
			<td>Recombinant Human Noggin</td>
			<td>100 ng/mL</td>
		</tr>
		<tr>
			<td>R-spondin 1-conditioned media</td>
			<td>10% conditioned media</td>
		</tr>
		<tr>
			<td>A83-01</td>
			<td>200 nM</td>
		</tr>
		<tr>
			<td>DHT</td>
			<td>1 nM</td>
		</tr>
		<tr>
			<td>Y-27632 dihydrochloride (ROCK inhibitor)</td>
			<td>10 &#181;M</td>
		</tr>
		<tr>
			<td>Advanced DMEM/F-12</td>
			<td>Base media</td>
		</tr>
		<tr>
			<td colspan="2">R-spondin 1-conditioned media is generated as described in Drost, et al.13. After addition of all components, filter sterilize mouse organoid media using 0.22 &#181;m filter. ROCK inhibitor is only added during establishment of culture and passaging of organoids.</td>
		</tr>
	</tbody>
</table>
Table 2: Instructions for the preparation of mouse organoid media.
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td colspan="2">Protocol for preparing Poly-HEMA-coated plates</td>
		</tr>
		<tr>
			<td>1</td>
			<td>Add 0.25 g Poly-HEMA to 50 mL 98% EtOH. Dissolve Poly-HEMA at 37 &#176;C on a shaker. This process takes at least 4 h.</td>
		</tr>
		<tr>
			<td>2</td>
			<td>Filter sterilize Poly-HEMA using 0.22 &#181;m filter.</td>
		</tr>
		<tr>
			<td>3</td>
			<td>Add 200 &#181;l of Poly-HEMA solution per well of a 24-well plate(s).</td>
		</tr>
		<tr>
			<td>4</td>
			<td>Remove lid(s) from 24-well plate(s) after adding Poly-HEMA and allow solution to evaporate overnight.</td>
		</tr>
		<tr>
			<td>5</td>
			<td>Wash each well twice with PBS and ensure wells are completely dry prior to storage after final wash. NOTE: Disrupting the Poly-HEMA coating during washing could contribute to 2-dimensional growth upon plating epithelial cells into organoid culture. To prevent damage to Poly-HEMA-coated wells, avoid direct contact with the pipette tip while washing. The integrity of the Poly-HEMA-coated wells will remain intact unless the Poly-HEMA is scraped off by the pipette tip.</td>
		</tr>
		<tr>
			<td>6</td>
			<td>Poly-HEMA-coated plates can be stored at 4 &#176;C for up to two weeks. NOTE: Wrapping plates in parafilm prior to storage will reduce the risk of contamination.</td>
		</tr>
	</tbody>
</table>
Table 3: Protocol for preparation of Poly-HEMA-coated plates.

Watch this Scientific Journal Video about Evaluating the Differentiation Capacity of Mouse Prostate Epithelial Cells Using Organoid Culture at JoVE.com

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

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 ...

evaluating-differentiation-capacity-mouse-prostate-epithelial-cells

Research

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Cancer Research

8.8K Views.  University of California, Los Angeles. Het 3D-organoïde systeem wordt beschouwd als meer fysiologisch relevant dan 2D-tests, en het kan waardevolle informatie over de biologie die anders zou een uitdaging om te verwerven. Dit is een aanpasbaar systeem waar we farmacologische en genetische manipulaties kunnen testen met minder tijd en aanzienlijk lagere kosten dan met behulp van een in vivo benadering. Matrix trial is erg lastig te hanteren, omdat het stolt wanneer het kamertemperatuur bereikt.Het kan alleen worden getrokke...

Video: Evaluating the Differentiation Capacity of Mouse Prostate Epithelial Cells Using 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 ...

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

Differentiation of Mouse Breast Epithelial HC11 and EpH4 Cells

Cadherins play an important role in the regulation of cell differentiation as well as neoplasia. Here we describe the origins and methods of the induction of differentiation of two mouse breast epithelial cell lines, HC11 and EpH4, and their use to study complementary stages of mammary gland development and neoplastic transformation.
The HC11 mouse breast epithelial cell line originated from the mammary gland of a pregnant Balb/c mouse. It differentiates when grown to confluence attached to a plastic Petri dish surface in medium containing fetal calf serum and Hydrocortisone, Insulin and Prolactin (HIP medium). Under these conditions, HC11 cells produce the milk proteins &#946;-casein and whey acidic protein (WAP), similar to lactating mammary epithelial cells, and form rudimentary mammary gland-like structures termed &#34;domes&#34;.
The EpH4 cell line was derived from spontaneously immortalized mouse mammary gland epithelial cells isolated from a pregnant Balb/c mouse. Unlike HC11, EpH4 cells can fully differentiate into spheroids (also called mammospheres) when cultured under three-dimensional (3D) growth conditions in HIP medium. Cells are trypsinized, suspended in a 20% matrix consisting of a mixture of extracellular matrix proteins produced by Engelbreth-Holm-Swarm (EHS) mouse sarcoma cells, plated on top of a layer of concentrated matrix coating a plastic Petri dish or multiwell plate, and covered with a layer of 10% matrix-containing HIP medium. Under these conditions, EpH4 cells form hollow spheroids that exhibit apical-basal polarity, a hollow lumen, and produce &#946;-casein and WAP.
Using these techniques, our results demonstrated that the intensity of the cadherin/Rac signal is critical for the differentiation of HC11 cells. While Rac1 is necessary for differentiation and low levels of activated RacV12 increase differentiation, high RacV12 levels block differentiation while inducing neoplasia. In contrast, EpH4 cells represent an earlier stage in mammary epithelial differentiation, which is inhibited by even low levels of RacV12.

We describe techniques for differentiation induction of two breast epithelial lines, HC11 and EpH4. While both require fetal calf serum, insulin, and prolactin to produce milk proteins, EpH4 cells can fully differentiate into mammospheres in three-dimensional culture. These complementary models are useful for signal transduction studies of differentiation and neoplasia.

Cadherins play an important role in the regulation of cell differentiation as well as neoplasia. Here we describe the origins and methods of the induction ...

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 ...

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.

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 ...

Bioengineering

Generation and Characterization of Murine Oral Mucosal Organoid Cultures

The mucous lining covering the inside of our mouth, the oral mucosa, is a highly compartmentalized tissue and can be subdivided into the buccal mucosa, gingiva, lips, palate, and tongue. Its uppermost layer, the oral epithelium, is maintained by adult stem cells throughout life. Proliferation and differentiation of adult epithelial stem cells have been intensively studied using in vivo mouse models as well as two-dimensional (2D) feeder-cell based in vitro models. Complementary to these methods is organoid technology, where adult stem cells are embedded into an extracellular matrix (ECM)-rich hydrogel and provided with a culture medium containing a defined cocktail of growth factors. Under these conditions, adult stem cells proliferate and spontaneously form three-dimensional (3D) cell clusters, the so-called organoids. Organoid cultures were initially established from murine small intestinal epithelial stem cells. However, the method has since been adapted for other epithelial stem cell types. Here, we describe a protocol for the generation and characterization of murine oral mucosal organoid cultures. Primary epithelial cells are isolated from murine tongue tissue, embedded into an ECM hydrogel, and cultured in a medium containing: epidermal growth factor (EGF), R-spondin, and fibroblast growth factor (FGF) 10. Within 7 to 14 days of initial seeding, the resulting organoids can be passaged for further expansion and cryopreservation. We additionally present strategies for the characterization of established organoid cultures via 3D whole-mount imaging and gene-expression analysis. This protocol may serve as a tool to investigate oral epithelial stem cell behavior ex vivo in a reductionist manner.

We present a method for the generation and characterization of oral mucosal organoid cultures derived from the tongue epithelium of adult mice.

The mucous lining covering the inside of our mouth, the oral mucosa, is a highly compartmentalized tissue and can be subdivided into the buccal mucosa, ...

Biology

Establishing 3D Endometrial Organoids from the Mouse Uterus

Endometrial tissue lines the inner cavity of the uterus and is under the cyclical control of estrogen and progesterone. It is a tissue that is composed of luminal and glandular epithelium, a stromal compartment, a vascular network, and a complex immune cell population. Mouse models have been a powerful tool to study the endometrium, revealing critical mechanisms that control implantation, placentation, and cancer. The recent development of 3D endometrial organoid cultures presents a state-of-the-art model to dissect the signaling pathways that underlie endometrial biology. Establishing endometrial organoids from genetically engineered mouse models, analyzing their transcriptomes, and visualizing their morphology at a single-cell resolution are crucial tools for the study of endometrial diseases. This paper outlines methods to establish 3D cultures of endometrial epithelium from mice and describes techniques to quantify gene expression and analyze the histology of the organoids. The goal is to provide a resource that can be used to establish, culture, and study the gene expression and morphological characteristics of endometrial epithelial organoids.

This protocol describes methodologies to establish mouse endometrial epithelial organoids for gene expression and histological analyses.

Endometrial tissue lines the inner cavity of the uterus and is under the cyclical control of estrogen and progesterone. It is a tissue that is composed of ...

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 ...

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

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