We thank Dr. Stephanie Pangas and Dr. Martin M. Matzuk (M.M.M.) for critical reading and editing of our manuscript. Studies were supported by Eunice Kennedy Shriver National Institute of Child Health and Human Development grants R00-HD096057 (D.M.), R01-HD105800 (D.M.), R01-HD032067 (M.M.M.), and R01-HD110038 (M.M.M.), and by NCI- P30 Cancer Center Support Grant (NCI-CA125123). Diana Monsivais, Ph.D. holds&#160;a Next Gen Pregnancy Award from the Burroughs Wellcome Fund.

Mouse handling and experimental studies were performed under protocols approved by the Institutional Animal Care and Use Committee (IACUC) of Baylor College of Medicine and guidelines established by the NIH Guide for the Care and Use of Laboratory Animals.
1. Isolation of uterine epithelium from mice using enzymatic and mechanical methods
NOTE: This section describes the steps required to establish, passage, freeze, and thaw epithelial endometrial organoids from mice using a gel matrix scaffold. Previous studies have determined that optimal cultures of mouse endometrial organoids are established from mice during the estrus phase4, which can be determined by cytological examination of a vaginal swab15. Adult female WT mice (6-8 weeks old, hybrid C57BL/6J and 129S5/SvEvBrd) were used for all experiments. The mice were humanely euthanized according to IACUC-approved guidelines using isoflurane sedation followed by cervical disarticulation. Once the mice are euthanized, the following steps should be followed. See the Table of Materials for details related to materials and solutions used in this protocol.
<ol>
	<li>Allow the gel matrix to thaw on ice for approximately 1-2 h before use.</li>
	<li>To dissect the mouse, use scissors to make a mid-line incision on the abdomen and gently peel back the skin to expose the underlying peritoneal layer. Use forceps to hold up the peritoneal layer and make lateral incisions with the scissors to expose the abdominal contents.
	<ol>
		<li>Locate the uterine horns by gently moving the abdominal fat pads aside. Dissect them by first holding them at the cervical junction and using scissors to cut along the mesenteric fat. Following the dissection of the mouse uterus, thoroughly remove fat tissue from the uterine horn with small scissors. 
		NOTE: Maintain sterility during the cell isolation by sterilizing all surgery tools before use, spray the mouse abdomen with 70% ethanol, and perform all steps following dissection under a sterile tissue culture hood.</li>
	</ol>
	</li>
	<li>Cut each uterine horn into small fragments, each measuring approximately 4-5 mm.</li>
	<li>Place all the uterine fragments from one uterus into one well of a 24-well plate containing 0.5 mL of 1% Trypsin. Allow the enzymatic solution to enter the uterine lumen, causing enzymatic separation of the endometrial epithelium from the underlying stroma.</li>
	<li>Incubate the 24-well plate in a 37 &#730;C humidified tissue culture incubator for approximately 1 h.</li>
	<li>After the 1 h incubation, transfer the uterine fragments to a 35 mm tissue culture plate containing 1 mL of Dulbecco&#39;s phosphate-buffered solution (DPBS).</li>
	<li>Under a dissection microscope, use fine forceps and a 1 mL pipette to mechanically separate the uterine epithelium from the uterine tube. While holding down one end of a uterine fragment with the forceps, gently run the pipette tip longitudinally across the fragment, squeezing the epithelium out of the other end of the uterine tube. Observe the separated epithelial sheets from the uterine fragment under the dissection microscope.</li>
	<li>Use the 1 mL pipette to collect and gently transfer the epithelial sheets into a 1.5 mL tube.</li>
	<li>Repeat the process for the remaining uterine fragments, transferring all the epithelial sheets to the same collection tube.</li>
	<li>Pellet the dissociated epithelial sheets by centrifugation for 5 min at 375 &#215; g.</li>
	<li>Carefully remove the supernatant to avoid disturbing the cell pellet.</li>
	<li>Resuspend the cell pellet in 0.5 mL of 2.5 mg/mL collagenase + 2 mg/mL DNase solution. Pipette up and down approximately 10x, or until a single-cell suspension is achieved.</li>
	<li>Add 0.5 mL of DMEM/F12 + 10% fetal bovine serum (FBS) + antibiotics, and centrifuge the cells for 5 min at 375 &#215; g. 
	&#8203;NOTE: For additional information on the broad-spectrum antibiotic used for cell culture, refer to the Table of Materials.</li>
	<li>Carefully remove the supernatant and resuspend the cells in 1 mL of DMEM/F12 + 10% FBS + antibiotics. Centrifuge the cells for 5 min at 375 &#215; g.</li>
</ol>
2. Processing of the stromal compartment
NOTE: This section outlines the protocols necessary for isolating the stromal compartment of the mouse endometrium. Given the increasing interest in epithelial/stromal co-culture experiments, it is important to be able to process the stromal cell populations in addition to the epithelial cells that will generate organoids.
<ol>
	<li>Once all the epithelium has been enzymatically and mechanically separated from the uterine fragments, the remaining tube-like structures are the &#34;stromal/myometrial&#34; compartments. Collect this tissue in a 2.5 mg/mL collagenase + 2 mg/mL DNase in HBSS solution.</li>
	<li>Incubate the stromal/myometrial sample on a 37 &#730;C shaker for 15 min.</li>
	<li>Following incubation, add 500 &#181;L of DMEM/F12 + 10% FBS + antibiotics per mouse, and filter the non-dissociated fragments through a 40 &#181;m cell filter.</li>
	<li>Pellet the cells by centrifugation for 5 min at 375 &#215; g.</li>
	<li>Carefully remove the supernatant and resuspend the cell pellet in 1 mL of DMEM/F12 + 10% FBS + antibiotics. Add the mixture dropwise to 10 mL of DMEM/F12 + 10% FBS + antibiotics in a 10 cm cell culture plate. Incubate the plate at 37 &#730;C in a humidified cell culture incubator.</li>
	<li>To generate co-cultures of mouse endometrial epithelial and stromal cells, follow the methods described for human endometrial organoids using scaffold-free or collagen matrix systems6,8. 
	&#8203;NOTE: While these techniques have not yet been published for mice, they can be adapted based on the published protocols with human endometrium.</li>
</ol>
3. Encapsulation of uterine epithelium into gel matrix to establish organoids
NOTE: Keep the gel matrix on ice until it is ready to be used.
<ol>
	<li>Remove the supernatant and resuspend the cell pellet in a volume of gel matrix that is 20x that of the cell pellet (i.e., if the cell pellet is 20 &#181;L, resuspend the cells with 400 &#181;L of gel matrix). Resuspend the pellet carefully to avoid introducing bubbles.</li>
	<li>Allow the gel matrix/cell suspension to settle at room temperature for ~10 min.</li>
	<li>Once the gel matrix/cell suspension becomes a semi-solid gel, use a P200 micropipette with a wide bore 200 &#181;L tip to gently aspirate 25 &#181;L of the gel matrix/cell suspension. Dispense three separate 25 &#181;L domes per well of a 12-well plate, and allow the gel matrix to cure for 15 min in a 37 &#730;C humidified tissue culture incubator.</li>
	<li>After the gel matrix has cured, add 750 &#181;L of organoid medium to each well containing gel matrix domes. Incubate at 37 &#730;C in a humidified tissue culture incubator. 
	&#8203;NOTE: Organoid media formulation is noted in the Table of Materials. Organoids typically form within 4 days of initial culture.</li>
</ol>
4. Gene expression analysis of endometrial organoids following treatment with estradiol
NOTE: This section describes the methods used to profile the gene expression of endometrial epithelial organoids using real-time qPCR following treatment with estradiol (E2; see Table 1). Because the endometrium is under the cyclical control of the ovarian hormone E2, testing the responsiveness of the organoids to E2 is an important measure of physiological function. We have obtained high-quality RNA and generated sufficient mRNA to profile gene expression using qPCR and/or RNA-sequencing from our endometrial epithelial organoids. This section describes how to collect organoids and process them for downstream analysis of gene expression. The selected treatment medium reflects the one used to treat cultured endometrial cells. However, it should be noted that this treatment medium can be optimized accordingly, as done for the treating of human endometrial 3D cultures with hormones8,16,17.
<ol>
	<li>Culture the endometrial organoids as described above.</li>
	<li>Remove the organoid medium and replace it with 750 &#181;L of starvation medium four days after seeding. Incubate overnight.</li>
	<li>The following morning, remove the starvation medium. Add 750 &#181;L of treatment medium containing either vehicle or 10 nM E2. Incubate for 48 h.</li>
	<li>Proceed with RNA isolation following the kit manufacturer&#39;s protocol.</li>
</ol>
5. Histological analysis of endometrial organoids
NOTE: Imaging the morphological features of endometrial organoids is critical to evaluating the cellular effect of growth factors, genetic manipulations, or small molecule inhibitors. This section describes the techniques used to fix, process, and image endometrial epithelial organoids using histological stains and antibody immunofluorescent staining.
<ol>
	<li>Prepare 1.5 mL microcentrifuge tubes containing 1 mL of 4% paraformaldehyde in 1x PBS. Place them on ice.</li>
	<li>Aspirate the medium from the wells of the 12-well plate containing the organoids.</li>
	<li>Using a 1 mL pipette tip with a cut tip, transfer 500 &#181;L of 4% paraformaldehyde to each well and gently detach the gel matrix domes from the bottom of the plate.</li>
	<li>Gently aspirate the entire gel matrix domes into the pipette tip and transfer to the 1.5 mL microcentrifuge tube.</li>
	<li>Fix the organoids by placing them on a rotator at 4 &#730;C overnight.</li>
	<li>The following morning, centrifuge the tube at 600 &#215; g for 5 min to pellet the organoids. Gently remove the 4% paraformaldehyde solution with a pipette and discard. Wash the organoids 2x with 70% ethanol.</li>
	<li>After the last wash, remove all but 50-100 &#181;L of ethanol from the tube. Set the tube aside.</li>
	<li>Place a tube of specimen processing gel in a water bath and microwave for ~30 s to melt the gel. Ensure that the specimen processing gel does not boil over by monitoring the consistency of the gel. 
	NOTE: Once the specimen processing gel is melted, but not boiling hot, work quickly to encapsulate the organoids.</li>
	<li>Transfer enough specimen processing gel to cover the entire surface of the mold (approximately 250 &#181;L).</li>
	<li>While the specimen processing gel is still molten, quickly transfer the 50 &#181;L of 70% ethanol solution containing the organoids. Ensure that the organoids are sunken or pushed to the bottom portion of the mold.</li>
	<li>Place the histology mold on a bucket of ice and allow the specimen processing gel to cool and solidify.</li>
	<li>Once the specimen processing gel is completely dry, carefully transfer the specimen processing gel square to a specimen bag, keeping track of the plane where the organoids are located.</li>
	<li>Place the bag in a histology cassette and process using standard methods used for formalin fixation and paraffin embedding of tissues18.</li>
	<li>Following fixation and embedding in paraffin, section into 5 &#181;m sections using a microtome19. Proceed with standard hematoxylin and eosin (H&#38;E) staining or immunostaining procedures, as outlined below.</li>
</ol>
6. Hematoxylin &#38; eosin staining
<ol>
	<li>Deparaffinize the sections as follows: xylene, 2 x 10 min; 100% ethanol, 2 x 3 min; 80% ethanol, 3 min; 60% ethanol, 3 min; dH2O, 2 x 3 min; 1 min in hematoxylin; tap water (3 x 5 s); 1 min in eosin.</li>
	<li>Dehydrate the sections as follows: 60% ethanol, 3 min; 80% ethanol, 3 min; 95% ethanol, 3 min; 100% ethanol, 2 x 3 min; xylene, 2 x 15 min.</li>
	<li>Mount using mounting medium.</li>
</ol>
7. Immunofluorescence staining
<ol>
	<li>Deparaffinize the sections as described in step 6.1.</li>
	<li>Perform antigen retrieval
	<ol>
		<li>Submerge the slides in antigen retrieval solution in a microwave-safe container.</li>
		<li>Microwave at high heat for 20 min, using 5 min intervals to ensure that the solution does not boil over.</li>
	</ol>
	</li>
	<li>After the 20 min antigen retrieval step is complete, allow the slides to cool on ice for 40 min while still submerged in antigen retrieval buffer.</li>
	<li>Wash the slides with 1x TBST for 3 min</li>
	<li>Block the slides by incubating in 3% BSA in TBST for 1 h at room temperature.</li>
	<li>Primary antibody incubation
	<ol>
		<li>Dilute the antibody in 3% BSA in TBST (1:50-1:1,000, depending on Ab).</li>
		<li>Incubate overnight at 4 &#176;C in a humidified chamber.</li>
		<li>Wash 3 x 5 min with TBST.</li>
	</ol>
	</li>
	<li>Secondary antibody incubation
	<ol>
		<li>Dilute the antibody in 3% BSA (in TBST) (1:250), or in 5% normal donkey serum.</li>
		<li>Incubate for 1 h at room temperature (RT) in the dark, since the antibody is conjugated to a fluorophore.</li>
	</ol>
	</li>
	<li>Nuclear staining
	<ol>
		<li>Dilute 4&#8242;,6-diamidino-2-phenylindole (DAPI) 1:1,000 in TBST.</li>
		<li>Incubate for 5 min at RT.</li>
		<li>Wash 2 x 5 min with TBST.</li>
	</ol>
	</li>
	<li>Mounting
	<ol>
		<li>Use one drop of mounting medium to mount the coverslip.</li>
		<li>Seal the coverslip using nail polish the following day.</li>
	</ol>
	</li>
</ol>

The authors have no conflicts of interest to disclose.

Here, we describe methods to generate endometrial epithelial organoids from mouse endometrium and the protocols routinely used for their downstream analysis. Endometrial organoids are a powerful tool to study the mechanisms that control endometrial-related diseases, such as endometriosis, endometrial cancer, and implantation failure. Landmark studies published in 2017 reported the conditions to culture long-term and renewable cultures of endometrial organoids from mouse and human epithelium4,...

The endometrium - the inner lining mucosal tissue of the uterine cavity - is a unique and highly dynamic tissue that plays critical roles in a woman&#39;s reproductive health. During the reproductive lifespan, the endometrium holds the potential to undergo hundreds of cycles of proliferation, differentiation, and breakdown, coordinated by the concerted action of the ovarian hormones - estrogen and progesterone. Studies of genetically engineered mice have uncovered basic biological mechanisms underpinning the endometrial response to hormones and control of embryo implantation, stromal cell decidualization, and pregnancy1. In vitro studies, however, have been limited due to difficulties in maintaining non-transformed primary mouse endometrial tissues in traditional 2D cell cultures2,3. Recent advances in the culture of endometrial tissues as 3D organ systems, or organoids, present a novel opportunity to investigate biological pathways that control endometrial cell regeneration and differentiation. Mouse and human endometrial organoid systems have been developed from pure endometrial epithelium encapsulated in various matrices4,5, while human endometrium has been cultured as scaffold-free epithelial/stromal co-cultures6,7, and more recently as collagen-encapsulated epithelial/stromal assembloids8. The growth and regenerative potential of epithelial organoid cultures is supported by a defined cocktail of growth factors and small molecule inhibitors that have been empirically determined to maximize growth and regeneration of the organoids4,5,9. Furthermore, the ability to freeze and thaw endometrial organoids permits the long-term banking of endometrial organoids from mice and humans for future studies.
Genetically engineered mice have revealed the complex signaling pathways that control early pregnancy and decidualization, and have been used as models of pregnancy loss, endometrial cancer, and endometriosis. These genetic studies have been largely achieved with cell-specific deletion of loxP flanked alleles (&#34;floxed&#34;) using cre recombinases that are specifically active in female reproductive tissues. These mouse models include the widely used progesterone receptor-cre10, which has strong recombinase activity in the endometrial epithelial and stromal tissues, lactoferrin i-cre, which induces endometrial epithelial recombination in adult mice11, or Wnt7a-cre, which triggers epithelial-specific deletion in M&#252;llerian-derived tissues12. Culturing endometrial tissues from genetically engineered mouse models as 3D organoids has provided an excellent opportunity to investigate endometrial biology and facilitate the identification of growth factors and signaling pathways that control endometrial cell renewal and differentiation13,14. Methods for the isolation and culture of mouse endometrial tissue are described in the literature and report the use of various enzymatic strategies for the isolation of uterine epithelium for subsequent culturing of endometrial epithelial organoids4. While previous literature provides a critical framework for endometrial epithelial organoid culture protocols4,5,6, this paper provides a clear, comprehensive method for generating, maintaining, processing, and analyzing these organoids. Standardization of these techniques is important for accelerating advancements in the field of women&#39;s reproductive biology. Here, we report a detailed methodology for the enzymatic and mechanical purification of mouse endometrial epithelial tissue for the subsequent culture of endometrial organoids in a gel matrix scaffold. We also describe the methodologies for downstream histological and molecular analyses of the gel matrix-encapsulated mouse endometrial epithelial organoids.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="4">Organoid Media Formulation</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>Corning Matrigel Growth Factor Reduced (GFR) Basement Membrane Matrix, *LDEV-free</td>
			<td>Corning</td>
			<td>354230</td>
			<td>100%</td>
		</tr>
		<tr>
			<td>Trypsin from Bovine Pancreas</td>
			<td>Sigma Aldrich</td>
			<td>T1426-1G</td>
			<td>1%</td>
		</tr>
		<tr>
			<td>Advanced DMEM/F12</td>
			<td>Life Technologies</td>
			<td>12634010</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>N2 supplement</td>
			<td>Life Technologies</td>
			<td>17502048</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>B-27&#8482; Supplement (50X), minus vitamin A</td>
			<td>Life Technologies</td>
			<td>12587010</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>Primocin</td>
			<td>Invivogen</td>
			<td>ant-pm-1</td>
			<td>100 &#181;g/mL</td>
		</tr>
		<tr>
			<td>N-Acetyl-L-cysteine</td>
			<td>Sigma Aldrich</td>
			<td>A9165-5G</td>
			<td>1.25 mM</td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>Life Technologies</td>
			<td>25030024</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>Nicotinamide</td>
			<td>Sigma Aldrich</td>
			<td>N0636-100G</td>
			<td>10 nM</td>
		</tr>
		<tr>
			<td>ALK-4, -5, -7 inhibitor, A83-01</td>
			<td>Tocris</td>
			<td>2939</td>
			<td>500 nM</td>
		</tr>
		<tr>
			<td>Recombinant human EGF</td>
			<td>Peprotech</td>
			<td>AF-100-15</td>
			<td>50 ng/mL</td>
		</tr>
		<tr>
			<td>Recombinant human Noggin</td>
			<td>Peprotech</td>
			<td>120-10C</td>
			<td>100 ng/mL</td>
		</tr>
		<tr>
			<td>Recombinant human Rspondin-1</td>
			<td>Peprotech</td>
			<td>120-38</td>
			<td>500 ng/mL</td>
		</tr>
		<tr>
			<td>Recombinant human FGF-10</td>
			<td>Peprotech</td>
			<td>100-26</td>
			<td>100 ng/mL</td>
		</tr>
		<tr>
			<td>Recombinant human HGF</td>
			<td>Peprotech</td>
			<td>100-39</td>
			<td>50 ng/mL</td>
		</tr>
		<tr>
			<td>WNT3a</td>
			<td>R&#38;D systems</td>
			<td>5036-WN</td>
			<td>200 ng/mL</td>
		</tr>
		<tr>
			<td>Other supplies and reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>Collagenase from Clostridium histolyticum</td>
			<td>Sigma Aldrich</td>
			<td>C0130-1G</td>
			<td>5 mg/mL</td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancreas</td>
			<td>Sigma Aldrich</td>
			<td>DN25-100MG</td>
			<td>2 mg/mL</td>
		</tr>
		<tr>
			<td>DPBS, no calcium, no magnesium</td>
			<td>ThermoFisher</td>
			<td>14190-250</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>HBSS, no calcium, no magnesium</td>
			<td>ThermoFisher</td>
			<td>14170112</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>Falcon&#160;Polystyrene Microplates (24-Well)</td>
			<td>Fisher Scientific</td>
			<td>#08-772-51</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon&#160;Polystyrene Microplates (12-Well)</td>
			<td>Fisher Scientific</td>
			<td>#0877229</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon Cell Strainers, 40 &#181;m</td>
			<td>Fisher Scientific</td>
			<td>#08-771-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Direct-zol RNA MiniPrep (50 &#181;g)</td>
			<td>Genesee Scientific</td>
			<td>11-331</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizol reagent</td>
			<td>Invitrogen</td>
			<td>15596026</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM/F-12, HEPES, no phenol red</td>
			<td>ThermoFisher</td>
			<td>11039021</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum, Charcoal stripped</td>
			<td>Sigma Aldrich</td>
			<td>F6765-500ML</td>
			<td>2%</td>
		</tr>
		<tr>
			<td>Estratiol (E2)</td>
			<td>Sigma Aldrich</td>
			<td>E1024-1G</td>
			<td>10 nM</td>
		</tr>
		<tr>
			<td>Formaldehyde 16% in aqueous solution, EM Grade</td>
			<td>VWR</td>
			<td>15710</td>
			<td>4%</td>
		</tr>
		<tr>
			<td>Epredia Cassette 1 Slotted Tissue Cassettes</td>
			<td>Fisher Scientific</td>
			<td>1000961</td>
			<td></td>
		</tr>
		<tr>
			<td>Epredia Stainless-Steel Embedding Base Molds</td>
			<td>Fisher Scientific</td>
			<td>64-010-15&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol, 200 proof (100%)</td>
			<td>Fisher Scientific</td>
			<td>22-032-601&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Histoclear</td>
			<td>Fisher Scientific</td>
			<td>50-899-90147</td>
			<td></td>
		</tr>
		<tr>
			<td>Permount Mounting Medium</td>
			<td>Fisher Scientific</td>
			<td>50-277-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Epredia Nylon Biopsy Bags</td>
			<td>Fisher Scientific</td>
			<td>6774010</td>
			<td></td>
		</tr>
		<tr>
			<td>HistoGel Specimen Processing Gel</td>
			<td>VWR</td>
			<td>83009-992</td>
			<td></td>
		</tr>
		<tr>
			<td>Hematoxylin solution Premium</td>
			<td>VWR</td>
			<td>95057-844</td>
			<td></td>
		</tr>
		<tr>
			<td>Eosin Y (yellowish) solution Premium</td>
			<td>VWR</td>
			<td>95057-848</td>
			<td></td>
		</tr>
		<tr>
			<td>TBS Buffer, 20X, pH 7.4</td>
			<td>GenDEPORT</td>
			<td>T8054</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>TBST (10X), pH 7.4</td>
			<td>GenDEPORT</td>
			<td>T8056</td>
			<td>1X</td>
		</tr>
		<tr>
			<td>Citric acid&#160;</td>
			<td>Sigma Aldrich</td>
			<td>C0759-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium citrate tribasic dihydrate</td>
			<td>Sigma Aldrich</td>
			<td>S4641-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween20</td>
			<td>Fisher Scientific</td>
			<td>BP337-500&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma Aldrich</td>
			<td>A2153-100G</td>
			<td>3%</td>
		</tr>
		<tr>
			<td>DAPI Solution (1 mg/mL)</td>
			<td>ThermoFisher</td>
			<td>62248</td>
			<td>1:1000 dilution</td>
		</tr>
		<tr>
			<td>VECTASHIELD Antifade Mounting Medium</td>
			<td>Vector Labs</td>
			<td>H-1000-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Clear Nail Polish</td>
			<td>Fisher Scientific</td>
			<td>NC1849418</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Superfrost Plus Microscope Slides</td>
			<td>Fisher Scientific</td>
			<td>22037246</td>
			<td></td>
		</tr>
		<tr>
			<td>VWR Micro Cover Glasses</td>
			<td>VWR</td>
			<td>48393-106</td>
			<td></td>
		</tr>
		<tr>
			<td>SuperScript VILO Master Mix</td>
			<td>ThermoFisher</td>
			<td>11755050</td>
			<td></td>
		</tr>
		<tr>
			<td>SYBR Green PCR Master Mix</td>
			<td>ThermoFisher</td>
			<td>4364346</td>
			<td></td>
		</tr>
		<tr>
			<td>Krt8 Antibody (TROMA-I)&#160;</td>
			<td>DSHB</td>
			<td>TROMA-I&#160;</td>
			<td>1:50 dilution</td>
		</tr>
		<tr>
			<td>Vimentin Antobody</td>
			<td>Cell Signaling</td>
			<td>5741S</td>
			<td>1:200 dilution</td>
		</tr>
		<tr>
			<td>Donkey anti-Rat IgG (H+L) Highly Cross-Adsorbed Secondary 
			Antibody, Alexa Fluor 594</td>
			<td>ThermoFisher</td>
			<td>A-21209</td>
			<td>1:250 dilution</td>
		</tr>
		<tr>
			<td>Donkey anti-Rabbin IgG (H+L) Highly Cross-Adsorbed Secondary 
			Antibody, Alexa Fluor 488</td>
			<td>ThermoFisher</td>
			<td>A-21206</td>
			<td>1:250 dilution</td>
		</tr>
		<tr>
			<td>ZEISS Stemi 508 Stereo Microscope</td>
			<td>ZEISS</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ZEISS Axio Vert.A1 Inverted Routine Microscope with digital camera</td>
			<td>ZEISS</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Primer Sequence</td>
			<td>Forward (5&#39;-3&#39;)</td>
			<td>Reverse (5&#39;-3&#39;)</td>
			<td>_</td>
		</tr>
		<tr>
			<td>Lipocalin 2 (Lcn2)</td>
			<td>GCAGGTGGTACGTTGTGGG</td>
			<td>CTCTTGTAGCTCATAGATGGTGC</td>
			<td></td>
		</tr>
		<tr>
			<td>Lactoferrin (Ltf)</td>
			<td>TGAGGCCCTTGGACTCTGT</td>
			<td>ACCCACTTTTCTCATCTCGTTC</td>
			<td></td>
		</tr>
		<tr>
			<td>Progesterone (Pgr)</td>
			<td>CCCACAGGAGTTTGTCAAGCTC</td>
			<td>TAACTTCAGACATCATTTCCGG</td>
			<td></td>
		</tr>
		<tr>
			<td>Glyceraldehyde 3 phosphate dehydrogenase (Gapdh)</td>
			<td>CAATGTGTCCGTCGTGGATCT</td>
			<td>GCCTGCTTCACCACCTTCTT</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Phase contrast images of mouse endometrial organoids 
We established organoids from WT mouse endometrial epithelium, as described in the attached protocol (see diagram in Figure 1). Following enzymatic dissociation of the mouse endometrial epithelium, epithelial sheets were mechanically separated from the uterine stromal cells and further dissociated with collagenase to generate a single-cell suspension. If performed correctly, this method of epithelial and stromal cell...

Watch this Scientific Journal Video about Establishing 3D Endometrial Organoids from the Mouse Uterus at JoVE.com

Establishing 3D Endometrial Organoids from the Mouse Uterus

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 ...

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5.6K Views.  Baylor College of Medicine. This protocol describes methodologies to establish mouse endometrial epithelial organoids for gene expression and histological analyses.

Video: Establishing 3D Endometrial Organoids from the Mouse Uterus

Intravitreous Injection for Establishing Ocular Diseases Model

Intravitreous injection is a widely used technique in visual sciences research. It can be used to establish animal models with ocular diseases or as direct application of local treatment. This video introduces how to use simple and inexpensive tools to finish the intravitreous injection procedure. Use of a 1 ml syringe, instead of a hemilton syringe, is used. Practical tips for how to make appropriate injection needles using glass pipettes with perfect tips, and how to easily connect the syringe needle with the glass pipette tightly together, are given.
To conduct a good intravitreous injection, there are three aspects to be observed: 1) injection site should not disrupt retina structure; 2) bleeding should be avoided to reduce the risk of infection; 3) lens should be untouched to avoid traumatic cataract. In brief, the most important point is to reduce the interruption of normal ocular structure. To avoid interruption of retina, the superior nasal region of rat eye was chosen. Also, the puncture point of the needle was at the par planar, which was about 1.5 mm from the limbal region of the rat eye. A small amount of vitreous is gently pushed out through the puncture hole to reduce the intraocular pressure before injection. With the 45&deg; injection angle, it is less likely to cause traumatic cataract in the rat eye, thus avoiding related complications and influence from lenticular factors. In this operation, there was no cutting of the conjunctiva and ocular muscle, no bleeding. With quick and minor injury, a successful intravitreous injection can be done in minutes. 
The injection set outlined in this particular protocol is specific for intravitreous injection. However, the methods and materials presented here can also be used for other injection procedures in drug delivery to the brain, spinal cord or other organs in small mammals.

Intravitreous injection is a widely used technique in visual sciences research for ocular diseases or as direct application of local treatment. This video demonstrated a protocol for intravitreous injection using a 1ml syringe with glass pipette. Useful tips about avoiding massive bleeding and lens damage are given.

Intravitreous injection is a widely used technique in visual sciences research. It can be used to establish animal models with ocular diseases or as ...

Establishing Primary Adult Fibroblast Cultures From Rodents

The importance of using primary cells, rather than cancer cell lines, for biological studies is becoming widely recognized. Primary cells are preferred in studies of cell cycle control, apoptosis, and DNA repair, as cancer cells carry mutations in genes involved in these processes. Primary cells cannot be cultured indefinitely due to the onset of replicative senescence or aneuploidization. Hence, new cultures need to be established regularly. The procedure for isolation of rodent embryonic fibroblasts is well established, but isolating adult fibroblast cultures often presents a challenge. Adult rodent fibroblasts isolated from mouse models of human disease may be a preferred control when comparing them to fibroblasts from human patients. Furthermore, adult fibroblasts are the only available material when working with wild rodents where pregnant females cannot be easily obtained. Here we provide a protocol for isolation and culture of adult fibroblasts from rodent skin and lungs. We used this procedure successfully to isolate fibroblasts from over twenty rodent species from laboratory mice and rats to wild rodents such as beaver, porcupine, and squirrel.

This article describes a protocol for isolation and maintenance of primary fibroblast cultures from skin and lung tissue of wild rodents.

The importance of using primary cells, rather than cancer cell lines, for biological studies is becoming widely recognized.  Primary cells are preferred ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...

Ex Vivo Method for Assessing the Mouse Reproductive Tract Spontaneous Motility and a MATLAB-based Uterus Motion Tracking Algorithm for Data Analysis

Dysmenorrhea, or painful cramping, is the most common symptom associated with menses in females and its severity can hinder women&#39;s everyday lives. Here, we present an easy and inexpensive method that would be instrumental for testing new drugs decreasing uterine contractility. This method utilizes the unique ability of the entire mouse reproductive tract to exhibit spontaneous motility when maintained ex vivo in a Petri dish containing oxygenated Krebs buffer. This spontaneous motility resembles the wave-like myometrial activity of the human uterus, referred to as endometrial waves. To demonstrate the effectiveness of the method, we employed a well-known uterine relaxant drug, epinephrine. We demonstrate that the spontaneous motility of the entire mouse reproductive tract can be quickly and reversibly inhibited by 1 &#181;M epinephrine in this Petri dish model. Documenting the changes of uterine motility can be easily done using an ordinary smart phone or a sophisticated digital camera. We developed a MATLAB-based algorithm allowing motion tracking to quantify spontaneous uterine motility changes by measuring the rate of uterine horn movements. A major advantage of this ex vivo approach is that the reproductive tract remains intact throughout the entire experiment, preserving all intrinsic intrauterine cellular interactions. The major limitation of this approach is that up to 10-20% of uteri may exhibit no spontaneous motility. Thus far, this is the first quantitative ex vivo method for assessing spontaneous uterine motility in a Petri dish model.

Uterine contractions are important for the well-being of females. However, pathologically increased contractility may result in dysmenorrhea, especially in younger females. Here, we describe a simple ex vivo preparation allowing quick assessment of the efficacy of smooth muscle relaxants that may be used for treating dysmenorrhea.

Dysmenorrhea, or painful cramping, is the most common symptom associated with menses in females and its severity can hinder women&#39;s everyday lives. ...

3D Culturing of Organoids from the Intestinal Villi Epithelium Undergoing Dedifferentiation

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is compartmentalized into crypts and villi: the stem and proliferating cells are confined to the crypts, whereas the villi epithelium contains only differentiated cells. Hence, the normal intestinal crypts, but not the villi, can give rise to organoids in 3D cultures. The procedure described here is applicable only to villus epithelium undergoing dedifferentiation leading to stemness. The method described uses the Smad4-loss-of-function:&#946;-catenin gain-of-function (Smad4KO:&#946;-cateninGOF) conditional mutant mouse. The mutation causes the intestinal villi to dedifferentiate and generate stem cells in the villi. Intestinal villi undergoing dedifferentiation are scraped off the intestine using glass slides, placed in a 70 &#181;m strainer and washed several times to filter out any loose cells or crypts prior to plating in BME-R1 matrix to determine their organoid-forming potential. Two main criteria were used to ensure that the resulting organoids were developed from the dedifferentiating villus compartment and not from the crypts: 1) microscopically evaluating&#160;the isolated villi to ensure absence of any tethered crypts, both before and after plating in the 3D matrix, and 2) monitoring the time course of organoid development from the villi. Organoid initiation from the villi occurs only two to five days after plating and appears irregularly shaped, whereas the crypt-derived organoids from the same intestinal epithelium are apparent within sixteen hours of plating and appear spherical. The limitation of the method, however, is that the number of organoids formed, and the time required for organoid initiation from the villi vary depending on the degree of dedifferentiation. Hence, depending upon the specificity of the mutation or the insult causing the dedifferentiation, the optimal stage at which villi can be harvested to assay their organoid forming potential, must be determined empirically.

The procedure describes isolation of the villi from the mouse intestinal epithelium undergoing dedifferentiation to determine their organoid forming potential.

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is ...

Establishing Stable Binary Cultures of Symbiotic Saccharibacteria from the Oral Cavity

Many bacterial species cannot be cultured in the laboratory using standard methods, posing a significant barrier to studying the majority of microbial diversity on earth. Novel approaches are required to culture these uncultured bacteria so that investigators can effectively study their physiology and lifestyle using the powerful tools available in the laboratory. The Candidate Phyla Radiation (CPR) is one of the largest groups of uncultivated bacteria, comprising ~15% of the living diversity on earth. The first isolate of this group was a member of the Saccharibacteria phylum, 'Nanosynbacter lyticus' strain TM7x. TM7x is an unusually small bacterium that lives as a symbiont in direct contact with a bacterial host, Schaalia odontolytica, strain XH001. Taking advantage of the unusually small cell size and its lifestyle as a symbiotic organism, we developed a protocol to rapidly culture Saccharibacteria from dental plaque. This protocol will show how to filter a suspension of dental plaque through a 0.2 &#181;m filter, then concentrate the collected Saccharibacteria cells and infect a culture of host organisms. The resulting coculture can be passaged as any normal bacterial culture and infection is confirmed by PCR. The resulting binary culture can be maintained in the laboratory and used for future experiments. While contamination is a possibility, the binary culture can be purified by either further filtering and reinfection of host, or by plating the binary culture and screening for infected colonies. We hope this protocol can be expanded to other sample types and environments, leading to the cultivation of many more species in the CPR.

We demonstrate a method for isolating difficult-to-grow members of the novel bacterial phylum, Saccharibacteria, by filtering dental plaque and co-culturing with host bacteria.

Many bacterial species cannot be cultured in the laboratory using standard methods, posing a significant barrier to studying the majority of microbial ...

Establishing Human Lung Organoids and Proximal Differentiation to Generate Mature Airway Organoids

The lack of a robust in vitro model of the human respiratory epithelium hinders the understanding of the biology and pathology of the respiratory system. We describe a defined protocol to derive human lung organoids from adult stem cells in the lung tissue and induce proximal differentiation to generate mature airway organoids. The lung organoids are then consecutively expanded for over 1 year with high stability, while the differentiated airway organoids are used to morphologically and functionally simulate human airway epithelium to a near-physiological level. Thus, we establish a robust organoid model of the human airway epithelium. The long-term expansion of lung organoids and differentiated airway organoids generates a stable and renewable source, enabling scientists to reconstruct and expand the human airway epithelial cells in culture dishes. The human lung organoid system provides a unique and physiologically active in vitro model for various applications, including studying virus-host interaction, drug testing, and disease modeling.

The protocol presents a method to derive human lung organoids from primary lung tissues, expand the lung organoids and induce proximal differentiation to generate 3D and 2D airway organoids that faithfully phenocopy the human airway epithelium.

The lack of a robust in vitro model of the human respiratory epithelium hinders the understanding of the biology and pathology of the respiratory system. ...

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, ...