This work was supported by NIH/NICHD HD091068-01 to Dr. Ina Dobrinski.

NOTE: Testes from 1-week-old piglets were obtained from a commercial pig farm as by-product from castration of commercial pigs. Sourcing of testes was approved by the Animal Care Committee at the University of Calgary.
1. Preparation of enzyme solutions for tissue digestion
NOTE: Three different enzymatic solutions are needed, which include two different collagenase IV solutions (solution A, B) and a deoxyribonuclease I (DNase I) solution.
<ol>
	<li>To prepare solution A, dissolve 20 mg of collagenase IV S (Table of Materials) and 40 mg of collagenase IV W (Table of Materials) in 25 mL of high glucose Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM). Then filter sterilize with a 0.22 &#181;m filter. Add 0.4 mL of fetal bovine serum (FBS) to solution A to inhibit trypsin activity.</li>
	<li>To prepare solution B, dissolve 80 mg of collagenase IV W (Table of Materials) in 40 mL of DMEM. Then filter sterilize with a 0.22 &#181;m filter.</li>
	<li>To prepare DNase I solution (7 mg/mL), dissolve 70 mg of DNase I in 10 mL of DMEM. Then filter sterilize with a 0.22 &#181;m filter. 
	NOTE: The enzyme concentration for collagenase IV solutions outlined here, varies from the concentrations (2 mg/mL) stated in the original article14. The current protocol yields cells with slightly higher viability. However, cells isolated by both protocols produce organoids with identical tissue architecture.</li>
</ol>
2. Testis tissue enzymatic digestion
<ol>
	<li>Collect the testes into a sterile beaker and wash with phosphate buffered saline (PBS) containing 1% penicillin/streptomycin (P/S). After washing, transfer the testes to a 100 mm tissue culture dish with PBS containing 1% P/S and remove the tunica vaginalis and epididymis using autoclaved scissors and forceps. Transfer the isolated testes to a new 100 mm dish and wash thoroughly with PBS containing 1% P/S.</li>
	<li>To maintain sterility, use another set of sterile scissors and forceps to peel the testicular parenchyma out of the tunica albuginea. Cut the testes along the longitudinal axis directly under the tunica. Then peel the testes out of the tunica using two forceps and place into a new 100 mm dish containing of 1 mL DMEM with 1% P/S.</li>
	<li>Mince the peeled testes with sterile scissors into 1-2 mm tissue pieces. After chopping, use sterile forceps to remove white fragments of connective tissue.</li>
	<li>Transfer the minced tissue pieces into solution A and top it up to 50 mL with DMEM to obtain a concentration of 0.4 mg/mL for collagenase IV S (Table of Materials) and a concentration of 0.8 mg/mL for collagenase IV W (Table of Materials). Place solution A containing the tissue pieces into a 37 &#176;C water bath for 30 min, gently invert the tubes every 5 min and check visually for the release of DNA.
	<ol>
		<li>Add 500 &#181;L of DNase I if free floating DNA is observed. After 30 min, centrifuge the tube at 90 x g with brakes at 25 &#176;C for 1.5 min and discard the supernatant. 
		NOTE: DNA would appear as a cloudy substance. When the tubes are shaken, the tissue pieces settle down whereas the DNA would stay afloat.</li>
	</ol>
	</li>
	<li>Add solution B to the tube and top up to 50 mL with DMEM to obtain a concentration of 1.2 mg/mL of collagenase IV W (Table of Materials). Place the tube into 37 &#176;C water bath for 30 min and gently invert the tube every 5 min. Add 500 &#181;L of DNase I if free floating DNA is observed.</li>
	<li>After 30 min, centrifuge the tube at 90 x g with brakes at 25 &#176;C for 1.5 min. After that discard the supernatant and wash once with PBS with 1% P/S. 
	NOTE: The discarded supernatants from both solution A and B will primarily contain interstitial cells.</li>
	<li>Top up the tube with PBS up to 50 mL. Carefully collect the tubules from the top and place into a new 50 mL tube. 
	NOTE: The big undigested tissue fragments quickly settle at the bottom, while digested seminiferous tubules will remain in suspension. No big tissue fragments should be collected. This procedure can be repeated several times until the solution in the original tube is almost clear and just undigested tissue fragments are remaining, which can be disposed.</li>
	<li>Centrifuge the tubules at 90 x g with brakes at 25 &#176;C for 1.5 min and discard the supernatant. Add fresh PBS and centrifuge again. Repeat this wash step twice.</li>
	<li>After the last PBS wash, remove the supernatant and resuspend the seminiferous tubules in 5 mL of PBS. Then add 15 mL of 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA) to the tubules. Place the tube in 37 &#176;C water bath and gently invert every 2 min. If a lot of free-floating DNA is observed, add 500 &#181;L of DNase I with 5 mL of DMEM.</li>
	<li>Evaluate the enzymatic digestion of tubules to single cells under the microscope (first after 5 min, then every 2 min). If mostly single cells can be detected, stop the reaction with 5 mL of FBS and filter through a 70 &#181;m mesh and then through a 40 &#181;m mesh. 
	NOTE: Care should be taken that the caps on the 50 mL tubes used for digestions (Solution A, B and 0.25% trypsin-EDTA) are tightened properly. If contamination in the water bath is a concern, paraffin film may be wrapped around the cap to ensure sterility.</li>
	<li>Centrifuge the single cells at 500 x g with brakes at 25 &#176;C for 5 min, resuspend in the enrichment medium (Dulbecco Modified Eagle Medium F/12 (DMEM/F12) containing 5% FBS and 1% P/S) and count the number of viable cells. This is the starting cell population. Fix 100 x 103 cells and perform immunocytochemistry for germ cell marker UCHL1 (Ubiquitin C-Terminal Hydrolase L1) as described12 and determine the percentage of germ cells in this starting cell population. 
	NOTE: Germ cell marker Promyelocytic leukemia zinc finger protein (PLZF)15 could also be used as a suitable alternative for UCHL1. In the starting cell population, the expected cell yield is around 700-800 x 109/g of tissue. The percentage of germ cells in this cell population should be 4-5%.</li>
	<li>Place around 20 x 106 of this starting cell population in two ultra low attachment 100 mm Petri dishes (10 x 106 cells suspended in 10 mL of DMEM/F12 containing 1% P/S in each dish) for 2 days in a tissue culture incubator (37 &#176;C, 5% CO2, 21% oxygen). Perform the germ cell enrichment with the remaining cells. 
	NOTE: To ensure optimum viability and cell quality, while the germ cell enrichment and subsequent quantification takes place, the starting cell population is placed in culture in ultra low attachment tissue culture dishes for 2 days.</li>
</ol>
3. Germ cell enrichment
NOTE: The procedure described above yields primarily Sertoli cells and germ cells. Different adhesion properties allow for the separation of Sertoli cells and germ cells via differential plating.
<ol>
	<li>Seed 20-25 x 106 cells of the starting cell population per 100 mm tissue culture dish in a total volume of 8 mL of enrichment medium (DMEM/F12 with 5% FBS and 1% P/S) for differential plating. Place the cells into an incubator (37 &#176;C, 5% CO2, 21% O2) and after 1.5 h, ensure that the majority of Sertoli cells attach to the plate.</li>
	<li>Collect and combine the supernatant (mainly containing germ cells) of 2 plates to one new 100 mm plate and place it back into the incubator. After 1 h, again combine the supernatant of 2 plates to a new 100 mm plate. Place the plates back into the incubator for overnight. 
	NOTE: The supernatants combined will primarily contain germ cells. The adhered cells that are discarded along with the plates are primarily testicular somatic cells such as Sertoli and peritubular myoid cells.</li>
	<li>Collect the enriched germ cells in two fractions as follows. 
	NOTE: Enriched germ cells adhere differently, non adherent fraction and slightly adhered fraction. Both these fractions together form the enriched germ cell population
	<ol>
		<li>Non adherent fraction: collect the supernatant.</li>
		<li>Slightly adherent fraction: wash the plates gently with PBS, treat with 2 mL of 1:20 dilution of 0.25% trypsin-EDTA for 5 min at room temperature, stop the reaction with 2 mL of enrichment medium and collect in the same tube as non adherent fraction. 
		NOTE: 0.25% Trypsin-EDTA needs to be diluted with PBS to produce a 1:20 trypsin solution.</li>
	</ol>
	</li>
	<li>Count the total number of cells using a cell counter, fix 100 x 103 cells and perform immunocytochemistry for germ cell marker UCHL1 as described12 and determine the percentage of germ cells in this enriched cell population. 
	NOTE: The percentage of germ cells in this cell population should be at least 60-70%. Since the immunocytochemistry for quantification would require 1 day, the enriched cells should be placed in an ultra low attachment 100 mm Petri dish with DMEM/F12 supplemented with 1% P/S for the duration to ensure optimum cell quality and viability.</li>
</ol>
4. Preparation of cells for seeding
<ol>
	<li>To harvest the starting cell preparation, collect the supernatant in a 50 mL tube from the ultra low attachment 100 mm dishes. Wash the plates vigorously with PBS to collect the adhered cells. No enzymatic digestion is needed. 
	NOTE: Some of the testicular cells, primarily Sertoli cells, can slightly adhere to the ultra low attachment plates.</li>
	<li>Combine the starting cell preparation and enriched germ cells to obtain a working cell preparation containing 25% germ cells. Centrifuge at 500 x g with brakes at 25 &#176;C for 5 min, discard the supernatant and resuspend the cells in organoid formation medium (DMEM/F12 supplemented with insulin 10 &#181;g/mL, transferrin 5.5 &#181;g/mL, selenium 6.7 ng/mL, 20 ng/mL epidermal growth factor, 1% P/S). Adjust the cell density to 2.4 x 106 per mL. 
	NOTE: Each well in the microwell plates used13 in this protocol contained 1,200 microwells.</li>
	<li>Use the formula below to calculate the number of cells per well as described below. 
	Number of cells per well = number of microwells x number of cells to be clustered for each organoid.</li>
	<li>To generate organoids from 1, 000 cells each, seed each with (1,200 x 1,000 cells each=) 1.2 x 106 cells. Adjust the cell density in the cell suspension in a way that the cells are seeded in a volume of 0.5 mL of medium. For organoids formed from 1,000 cells each, use a density of 2.4 x 106 cells per mL (1.2 x 106 cells in 0.5 mL).</li>
</ol>
5. Preparation of microwells to receive cells
NOTE: To ensure that the cells do not adhere to the microwell surface, treat with a surfactant rinsing solution that is available for purchase by the manufacturer.
<ol>
	<li>Add 0.5 mL of rinsing solution to each well. Ensure that no air bubbles are trapped in the well. To remove air bubbles, if any, centrifuge the plate at 2,000 x g with brakes at 25 &#176;C for 2 min.</li>
	<li>Observe the plate under a low-magnification inverted microscope, to verify that the bubbles have been removed from the microwells. If trapped bubbles are observed, centrifuge again at 2,000 x g with brakes at 25 &#176;C for 2 min to remove any remaining bubbles.</li>
	<li>Cover the plate with the lid and incubate for 30 min at room temperature. After the treatment is complete, remove the rinsing solution and immediately wash the plate with sterile water or PBS. Ensure that the plate do not dry out after the treatment with the rinsing solution.</li>
</ol>
6. Generation of testicular organoids
<ol>
	<li>Add 0.5 mL of organoid formation medium (without any cells) to each well and centrifuge at 2,000 x g with brakes at 25 &#176;C for 2 min to remove any trapped air bubbles. Observe the well under a low-magnification inverted microscope to verify that the air bubbles have been removed. If trapped bubbles are observed, centrifuge again at 2,000 x g with brakes at 25 &#176;C for 2 min to remove any remaining bubbles.</li>
	<li>Add 0.5 mL of the working cell suspension and gently mix by pipetting up and down. Centrifuge at 500 x g with brakes at 25 &#176;C for 5 min. Use an inverted microscope to verify that the cells have clustered within the microwells.</li>
	<li>Transfer the plate into a cell culture incubator and culture for 5 days. Change half of the medium every other day.
	<ol>
		<li>To change medium without losing any organoids, touch the pipette tip to the wall of the well and lower gently to touch the meniscus and slowly draw the medium. Make sure to follow the meniscus as it drops down. To add fresh medium, touch the pipette tip to the wall of the well on the same side as the medium withdrawal and add fresh medium gently against the well wall, allowing it to slowly flow down the wall.</li>
	</ol>
	</li>
	<li>To recover the organoids, gently pipette the medium up and down using a wide mouth pipette. This would allow the organoids to come out of the microwells.</li>
	<li>Collect the organoids, wash with PBS, fix and perform immunocytochemistry for germ cell marker (UCH-L1), Sertoli cell marker (GATA Binding Protein 4-GATA4), peritubular myoid cell marker (&#945;-Smooth Muscle Actin-&#945;SMA) and Leydig cell marker (Cytochrome P450-CYP450) as described12 and visualize under a confocal microscope.</li>
</ol>

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Dr. Ungrin has a financial interest in the AggreWell technology as an inventor.

We have established a simple method that allows the consistent, repeatable generation of large numbers of testicular organoids with tissue architecture that is similar to testis in vivo12. While the approach was developed using porcine testis cells, it is more broadly applicable also to mouse, non-human primate and human testis12. A number of different methods have been reported for producing testicular organoids8,9...

In recent years, there has been a resurgence of interest in three-dimensional (3D) organoids. Different organs such as intestine1, stomach2, pancreas3,4, liver5, and brain6 have been successfully derived into 3D organoid systems. These organoids have architectural and functional similarities to the organs in vivo and are more biologically relevant for study of tissue microenvironment than monolayer culture systems7. As a result, testicular organoids have started to garner interest as well8,9,10,11,12. The majority of methods reported so far are complex, non-high throughput10 and require the addition of ECM proteins8,10. This complexity also leads to issues with reproducibility. A simple and reproducible method is needed that allows for the generation of testicular organoids with cell-associations that are like testis in vivo.
We have recently reported a system to address these requirements12. Using the pig as a model, we employed a centrifugal forced aggregation approach in the microwell system. In the microwell system, each well contains a large number of identical smaller microwells13. This allows for the generation of numerous spheroids of uniform size. The microwell system enabled generation of large numbers of uniform organoids with a testis-specific architecture. The system is simple and does not require addition of ECM proteins.

<table><tbody><tr><td>100 mm ultra low attachment tissue culture dish</td><td>Corning</td><td>#CLS3262</td><td></td></tr><tr><td>100 mm tissue culture dish</td><td>Corning</td><td>#353803</td><td></td></tr><tr><td>Aggrwell 400</td><td>Stemcell Technologies</td><td>#34411</td><td></td></tr><tr><td>Anti-Adherence Rinsing Solution</td><td>Stemcell Technologies</td><td>#07010</td><td></td></tr><tr><td>Collagenase type IV from Clostridium histolyticum</td><td>Sigma-Aldrich</td><td>#C5138</td><td>referred as Collagenase IV S</td></tr><tr><td>Collagenase type IV Worthington</td><td>Worthington-Biochem</td><td>#LS004189</td><td>referred as Collagenase IV W</td></tr><tr><td>Deoxyribonuclease I from bovine pancreas</td><td>Sigma-Aldrich</td><td>#DN25</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Modified Eagle&amp;rsquo;s Medium/F12</td><td>Gibco</td><td>#11330-032</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Modified Eagle&amp;rsquo;s Medium - high glucose</td><td>Sigma-Aldrich</td><td>#D6429</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered Saline</td><td>Sigma-Aldrich</td><td>#D8537</td><td></td></tr><tr><td>Epidermal Growth Factor</td><td>R&amp;amp;D Systems</td><td>#236-EG</td><td></td></tr><tr><td>Falcon Cell Strainers 70 &amp;micro;m</td><td>FisherScientific</td><td>#352350</td><td></td></tr><tr><td>Falcon Cell Strainers 40 &amp;micro;m</td><td>FisherScientific</td><td>#352340</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>ThermoFisher Scientific</td><td>#12483-020</td><td></td></tr><tr><td>Insulin-Transferrin-Selenium</td><td>Gibco</td><td>#41400-045</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma-Aldrich</td><td>#P4333</td><td></td></tr><tr><td>Porcine testicular tissue</td><td>Sunterra Farms Ltd (Alberta, Canada)</td><td></td><td></td></tr><tr><td>Steriflip-GP Sterile Centrifuge Tube Top Filter Unit</td><td>Millipore</td><td>#SCGP00525</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Sigma</td><td>#T4049</td><td></td></tr></tbody></table>

generation of porcine testicular organoids with testis specific architecture using microwell culture

Organoids are three dimensional structures composed of multiple cell types that are capable of recapitulating tissue architecture and functions of organs in vivo. Formation of organoids has opened up different avenues of basic and translational research. In recent years, testicular organoids have garnered interest in the field of male reproductive biology. Testicular organoids allow for the study of cell-cell interactions, tissue development, and the germ cell niche microenvironment and facilitate high throughput drug and toxicity screening. A method is needed to reliably and reproducibly generate testicular organoids with testis specific tissue architecture. The microwell culture system contains a dense array of pyramid-shaped microwells. Testicular cells derived from pre-pubertal testes are centrifuged into these microwells and cultured to generate testicular organoids with testis-specific tissue architecture and cell associations. Thousands of homogeneous organoids can be generated via this process. The protocol reported here will be of broad interest to researchers studying male reproduction.

Isolated cells from 1-week old porcine testes that were cultured in the microwells self-organized into spheroids (Figure 1A, Figure 2), with delineated and distinct exterior (seminiferous epithelium) and interior compartments (interstitium) (Figure 1B, Figure 2). The two compartments were separated by a collagen IV+ve basement...

Watch this Scientific Journal Video about Generation of Porcine Testicular Organoids with Testis Specific Architecture using Microwell Culture at JoVE.com

Generation of Porcine Testicular Organoids with Testis Specific Architecture using Microwell Culture

Here, we present a protocol for the reproducible generation of porcine testicular organoids with testis specific tissue architecture using the commercially available microwell culture system.

Organoids are three dimensional structures composed of multiple cell types that are capable of recapitulating tissue architecture and functions of organs ...

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8.8K Views.  University of Calgary. Here, we present a protocol for the reproducible generation of porcine testicular organoids with testis specific tissue architecture using the commercially available microwell culture system.

Video: Generation of Porcine Testicular Organoids with Testis Specific Architecture using Microwell Culture

Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

Investigating organogenesis in utero is a technically challenging process in placental mammals due to inaccessibility of reagents to embryos that develop within the uterus. A newly developed ex vivo upright droplet culture method provides an attractive alternative to studies performed in utero. The ex vivo droplet culture provides the ability to examine and manipulate cellular interactions and diverse signaling pathways through use of various blocking and activating compounds; additionally, the effects of various pharmacological reagents on the development of specific organs can be studied without unwanted side effects of systemic drug delivery in utero. As compared to other in vitro systems, the droplet culture not only allows for the ability to study three-dimensional morphogenesis and cell-cell interactions, which cannot be reproduced in mammalian cell lines, but also requires significantly less reagents than other ex vivo and in vitro protocols. This paper demonstrates proper mouse fetal organ dissection and upright droplet culture techniques, followed by whole organ immunofluorescence to demonstrate the effectiveness of the method. The ex vivo droplet culture method allows the formation of organ architecture comparable to what is observed in vivo and can be utilized to study otherwise difficult-to-study processes due to embryonic lethality in in vivo models. As a model application system, a small-molecule inhibitor will be utilized to probe the role of vascularization in testicular morphogenesis. This ex vivo droplet culture method is expandable to other fetal organ systems, such as lung and potentially others, although each organ must be extensively studied to determine any organ-specific modifications to the protocol. This organ culture system provides flexibility in experimentation with fetal organs, and results obtained using this technique will help researchers gain insights into fetal development.

Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

The&#160;ex vivo&#160;upright droplet culture is an alternative to current&#160;in vitro&#160;and&#160;in vivo&#160;experimental techniques. This protocol is easy to perform and requires smaller amounts of reagent, while permitting the ability to manipulate and study fetal vascularization, morphogenesis, and organogenesis.

Investigating organogenesis in utero is a technically challenging process in placental mammals due to inaccessibility of reagents to embryos that develop ...

Developmental Biology

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

Generation of Mosaic Mammary Organoids by Differential Trypsinization

Organoids offer self-organizing, three-dimensional tissue structures that recapitulate physiological processes in the convenience of a dish. The murine mammary gland is composed of two distinct epithelial cell compartments, serving different functions: the outer, contractile myoepithelial compartment and the inner, secretory luminal compartment. Here, we describe a method by which the cells comprising these compartments are isolated and then combined to investigate their individual lineage contributions to mammary gland morphogenesis and differentiation. The method is simple and efficient and does not require sophisticated separation technologies such as fluorescence activated cell sorting. Instead, we harvest and enzymatically digest the tissue, seed the epithelium on adherent tissue culture dishes, and then use differential trypsinization to separate myoepithelial from luminal cells with ~90% purity. The cells are then plated in an extracellular matrix where they organize into bilayered, three-dimensional (3D) organoids that can be differentiated to produce milk after 10 days in culture. To test the effects of genetic mutations, cells can be harvested from wild type or genetically engineered mouse models, or they can be genetically manipulated prior to 3D culture. This technique can be used to generate mosaic organoids that allow investigation of gene function specifically in the luminal or myoepithelial compartment.

The mammary gland is a bilayered structure, comprising outer myoepithelial and inner luminal epithelial cells. Presented is a protocol to prepare organoids using differential trypsinization. This efficient method allows researchers to separately manipulate these two cell types to explore questions concerning their roles in mammary gland form and function.

Organoids offer self-organizing, three-dimensional tissue structures that recapitulate physiological processes in the convenience of a dish. The murine ...

Extra Cellular Matrix-Based and Extra Cellular Matrix-Free Generation of Murine Testicular Organoids

Testicular organoids provide a tool for studying testicular development, spermatogenesis, and endocrinology in vitro. Several methods have been developed in order to create testicular organoids. Many of these methods rely upon extracellular matrix (ECM) to promote de novo tissue assembly, however, there are differences between methods in terms of biomimetic morphology and function of tissues. Moreover, there are few direct comparisons of published methods. Here, a direct comparison is made by studying differences in organoid generation protocols, with provided outcomes. Four archetypal generation methods: (1) 2D ECM-free, (2) 2D ECM, (3) 3D ECM-free, and (4) 3D ECM culture are described. Three primary benchmarks were used to assess the testicular organoid generation. These are cellular self-assembly, inclusion of major cell types (Sertoli, Leydig, germ, and peritubular cells), and appropriately compartmentalized tissue architecture. Of the four environments tested, 2D ECM and 3D ECM-free cultures generated organoids with internal morphologies most similar to native testes, including the de novo compartmentalization of tubular versus interstitial cell types, the development of tubule-like-structures, and an established long-term endocrine function. All methods studied utilized unsorted, primary murine testicular cell suspensions and used commonly accessible culture resources. These testicular organoid generation techniques provide a highly accessible and reproducible toolkit for research initiatives into testicular organogenesis and physiology in vitro.

Here, four methods for generating testicular organoids from primary neonatal murine testicular cells are described i.e., extracellular matrix (ECM) and ECM-free 2D and 3D culture environments. These techniques have multiple research applications and are especially useful for studying testicular development and physiology in vitro.

Testicular organoids provide a tool for studying testicular development, spermatogenesis, and endocrinology in vitro. Several methods have been developed ...

hiPSC-Derived Vascular Organoids for Drug Screening and Disease Modeling

Vascular Organoid Generation from Human-Induced Pluripotent Stem Cells

Vascular organoids derived from human induced pluripotent stem cells (hiPSCs) recapitulate the cell type diversity and complex architecture of human vascular networks. This three-dimensional (3D) model holds substantial potential for vascular pathology modeling and in vitro drug screening. Despite recent advances, a key technical challenge remains in reproducibly generating organoids with consistent quality, which is crucial for downstream assays and applications. Here, a modified protocol is presented that improves both the homogeneity and reproducibility of vascular organoid generation. The modified protocol incorporates the use of microwells and the CEPT cocktail (chroman 1, emricasan, polyamines, and the integrated stress response inhibitor, trans-ISRIB) to improve embryoid body formation and cell survival. Differentiated, mature vascular organoids generated using this protocol are characterized by whole-mount 3D immunofluorescence microscopy to analyze their morphology and complex vasculature. This protocol enables the production of high-quality vascular organoids in a scalable manner, potentially facilitating their use in disease modeling and drug screening applications.

Author Spotlight: Improving Reproducibility in Vascular Organoids Using ROCK Inhibitors and Microwell Confinement

This protocol presents a method for generating vascular organoids using human pluripotent stem cells.

Vascular organoids derived from human induced pluripotent stem cells (hiPSCs) recapitulate the cell type diversity and complex architecture of human ...

Bioengineering

Generation of Three-Dimensional Spheroids/Organoids from Two-Dimensional Cell Cultures Using a Novel Stamp Device

Three-dimensional (3D) cell cultures provide a more accurate representation of the in vivo microenvironment than conventional two-dimensional (2D) cultures, since they promote enhanced interactions among cells and the extracellular matrix.&#160;This study aimed to develop an efficient, cost-effective, and reproducible methodology to generate 3D cell structures (spheroids/organoids) using an innovative stamp-based system to create microwells in agarose molds.A novel stamp was used to produce 663 microwells per well of a 6-well plate, providing an ideal environment for cell aggregation. Primary porcine pancreatic islet cells were seeded into these microwells, where they aggregated to form spheroids/organoids. The cultures were incubated at 37 &#176;C under 5% CO2, and the medium was replaced every 3 days. Spheroid formation was periodically monitored, and samples were collected for characterization.&#160;The method successfully generated uniform and high-quality spheroids, reducing experimental variability, minimizing manipulation, and enhancing cell interactions. The use of agarose-based micropattern molds provided a simplified, controlled environment for 3D cultures, offering a standardized and cost-effective solution.This methodology supports applications for drug testing and tissue engineering, offering a practical and scalable platform for 3D cell culture models that can be easily implemented in various laboratory settings.

This study presents a cost-effective and efficient methodology for generating 3D cell structures using a stamp-based system to create microwells in agarose molds. The system promotes the formation of uniform spheroids/organoids, thus improving cell interactions. This approach reduces experimental variability and supports applications in drug testing and tissue engineering.

Three-dimensional (3D) cell cultures provide a more accurate representation of the in vivo microenvironment than conventional two-dimensional (2D) ...

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

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

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by standard imaging approaches. This protocol describes a way to prepare microfabricated organoid culture chips, which enable multiscale, 3D live imaging on a user-friendly instrument requiring minimal manipulations and capable of up to 300 organoids/h imaging throughput. These culture chips are compatible with both air and immersion objectives (air, water, oil, and silicone) and a wide range of common microscopes (e.g., spinning disk, point scanner confocal, wide field, and brightfield). Moreover, they can be used with light-sheet modalities such as the single-objective, single-plane illumination microscopy (SPIM) technology (soSPIM). 
The protocol described here gives detailed steps for the preparation of the microfabricated culture chips and the culture and staining of organoids. Only a short length of time is required to become familiar with, and consumables and equipment can be easily found in normal biolabs. Here, the 3D imaging capabilities will be demonstrated only with commercial standard microscopes (e.g., spinning disk for 3D reconstruction and wide field microscopy for routine monitoring).

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

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by ...

Culture of Piglet Intestinal 3D Organoids from Cryopreserved Epithelial Crypts and Establishment of Cell Monolayers

Intestinal organoids are increasingly being used to study the gut epithelium for digestive disease modeling, or to investigate interactions with drugs, nutrients, metabolites, pathogens, and the microbiota. Methods to culture intestinal organoids are now available for multiple species, including pigs, which is a species of major interest both as a farm animal and as a translational model for humans, for example, to study zoonotic diseases. Here, we give an in-depth description of a procedure used to culture pig intestinal 3D organoids from frozen epithelial crypts. The protocol describes how to cryopreserve epithelial crypts from the pig intestine and the subsequent procedures to culture 3D intestinal organoids. The main advantages of this method are (i) the temporal dissociation of the isolation of crypts from the culture of 3D organoids, (ii) the preparation of large stocks of cryopreserved crypts derived from multiple intestinal segments and from several animals at once, and thus (iii) the reduction in the need to sample fresh tissues from living animals. We also detail a protocol to establish cell monolayers derived from 3D organoids to allow access to the apical side of epithelial cells, which is the site of interactions with nutrients, microbes, or drugs. Overall, the protocols described here is a useful resource for studying the pig intestinal epithelium in veterinary and biomedical research.

Here, we describe a protocol to culture pig intestinal 3D organoids from cryopreserved epithelial crypts. We also describe a method to establish cell monolayers derived from 3D organoids, allowing access to the apical side of epithelial cells.

Intestinal organoids are increasingly being used to study the gut epithelium for digestive disease modeling, or to investigate interactions with drugs, ...

Two-dimensional Porcine Intestinal Organoids Reflecting the Physiological Properties of Native Gut

The gastrointestinal tract (GIT) serves both in the digestion of food and the uptake of nutrients but also as a protective barrier against pathogens. Traditionally, research in this area has relied on animal experiments, but there's a growing demand for alternative methods that adhere to the 3R principles-replace, reduce, and refine. Porcine organoids have emerged as a promising tool, offering a more accurate in vitro replication of the in vivo conditions than traditional cell models. One major challenge with intestinal organoids is their inward-facing apical surface and outward-facing basolateral surface. This limitation can be overcome by creating two-dimensional (2D) organoid layers on transwell inserts (from here on referred to as insert(s)), providing access to both surfaces. In this study, we successfully developed two-dimensional cultures of porcine jejunum and colon organoids. The cultivation process involves two key phases: First, the formation of a cellular monolayer, followed by the differentiation of the cells using tailored media. Cellular growth is tracked by measuring transepithelial electrical resistance, which stabilizes by day 8 for colon organoids and day 16 for jejunum organoids. After a 2-day differentiation phase, the epithelium is ready for analysis. To quantify and track active electrogenic transport processes, such as chloride secretion, we employ the Ussing chamber technique. This method allows for real-time measurement and detailed characterization of epithelial transport processes. This innovative in vitro model, combined with established techniques like the Ussing chamber, provides a robust platform for physiologically characterizing the porcine GIT within the 3R framework. It also opens opportunities for investigating pathophysiological mechanisms and developing potential therapeutic strategies.

This study outlines a protocol for generating 2D monolayers of porcine organoids derived from the small and large intestines. The growth of these monolayers is marked by increasing TEER values, indicating robust epithelial integrity. Additionally, these monolayers exhibit physiological secretory responses in Ussing chamber experiments following the application of forskolin.

The gastrointestinal tract (GIT) serves both in the digestion of food and the uptake of nutrients but also as a protective barrier against pathogens. ...

Generation of Porcine Testicular Organoids with Testis Specific Architecture using Microwell Culture

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Chapters in this video

Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

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

Generation of Mosaic Mammary Organoids by Differential Trypsinization

Extra Cellular Matrix-Based and Extra Cellular Matrix-Free Generation of Murine Testicular Organoids

Vascular Organoid Generation from Human-Induced Pluripotent Stem Cells

Generation of Three-Dimensional Spheroids/Organoids from Two-Dimensional Cell Cultures Using a Novel Stamp Device

Generation and Characterization of Murine Oral Mucosal Organoid Cultures

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

Culture of Piglet Intestinal 3D Organoids from Cryopreserved Epithelial Crypts and Establishment of Cell Monolayers

Two-dimensional Porcine Intestinal Organoids Reflecting the Physiological Properties of Native Gut