We thank Ben Abrams for technical assistance and core support from the University of California, Santa Cruz (UCSC) Institute for the Biology of Stem Cells (IBSC). We thank Susan Strome and Bill Saxton for the use of their Solamere Spinning Disk Confocal Microscope. This work was supported in part by grants to UCSC from the Howard Hughes Medical Institute through the James H. Gilliam Fellowships for Advanced Study program (S.R.), from the NIH (NIH GM058903) for the initiative for maximizing student development (H.M.) and from the National Science Foundation for a graduate research fellowship (O.C. DGE 1339067) and by a grant (A18-0370) from the UC-Cancer Research Coordinating Committee (LH).

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of the University of California, Santa Cruz.
1. Day 1: Mammary gland digestion
<ol>
	<li>Prepare to harvest the MGs from mature female mice 10-14 weeks of age.
	<ol>
		<li>Perform the harvesting on an open bench under aseptic conditions.</li>
		<li>Sterilize all surgical supplies, cork boards, and pins by autoclaving and soaking in 70% alcohol for 20 min prior to surgery.</li>
		<li>Anesthetize animals with sodium pentobarbital (2X anesthetic dose of 0.06 mg/g body weight) delivered via intraperitoneal injection with a 0.5 mL insulin syringe.</li>
		<li>Monitor the level of anesthesia by pinching the animal&#39;s toes to check for a reflex response and commence the protocol only after the animal is fully anesthetized.</li>
		<li>Place the animal on its back, pin its appendages to the corkboard, and wipe down its abdomen and chest with ethanol.</li>
	</ol>
	</li>
	<li>To harvest the #2, 3, 4, and 5 MGs from one mouse (i.e., 8 MGs, Figure 1A), identify the midline between the two hind legs and make a small incision (1 cm) on the abdominal skin with sharp scissors, then extend the cut up to the neck18.</li>
	<li>Follow by making small cuts laterally towards the legs and arms to allow for the release of the skin using a cotton swab. Pull the skin away and stretch it tight before pinning it down on one side (Figure 1B, step 1)18. Remove the MGs by cutting under them, and remove the lymph nodes from the #4 glands (Figure 1B, steps 2, 3)18. Repeat the procedure on the other side of the body.</li>
	<li>Collect the MG tissue in 50 mL of 4 &#176;C Dulbecco&#39;s Modified Eagle&#39;s Media (DMEM)/Nutrient Mixture F12 (F12) supplemented with 5% fetal bovine serum (FBS) and 1X Antibiotic-Antimycotic (Anti-Anti)18.</li>
	<li>Chop the glands in a 35 mm dish or on a ceramic plate using a razor blade or tissue chopper. Rotate the plate every five manual chops or every round on the tissue chopper until the tissue pieces can fit through a 1 mL micropipette tip with ease (~0.1 mm/fragment, Figure 1C).</li>
	<li>Digest the MGs in Digestion Medium (see Table 1) for 14 h in a 6 well low adhesion dish at 37 &#176;C, 5% CO2.</li>
</ol>
2. Day 2: Isolation of mammary epithelial tissue fragments
<ol>
	<li>After 14 h of digestion, gently mix by pipetting the digested tissue 10X using a 1 mL micropipette to break down any remaining stroma or adipose tissue, ensuring that neither bubbles nor excess mechanical force are generated. 
	NOTE: If there is incomplete digestion after 14 h, this could be due to the accumulation of sheared DNA. In this case, add 1 &#181;L of 1 mg/mL deoxyribonuclease I (DNase I) per 2 mL of Digestion Medium. Incubate for another 30 min at 37 &#176;C, 5% CO2.</li>
	<li>Collect tissue in a 15 mL tube and rinse the well used for digestion with 2-3 mL of tissue culture grade 1X Dulbecco&#39;s Phosphate Buffered Saline (DPBS) free of Ca2+ and Mg2+. Centrifuge at 600 x g for 10 min.</li>
	<li>Evacuate the supernatant containing the lipid layer and medium, and then resuspend the pellet in 5 mL of DPBS and switch to a new 15 mL tube. Centrifuge at 600 x g for 10 min.</li>
	<li>During centrifugation place a 70 &#181;m nylon cell strainer in a 50 mL tube and prewet the strainer using 10 mL of 37 &#176;C DMEM/F12.</li>
	<li>Resuspend tissue fragments from protocol step 2.4 using 5 mL of DPBS and pass the suspension through a prewet 70 &#181;m nylon cell strainer to remove stromal cells and single cells (Figure 1D).</li>
	<li>Collect the tissue fragments on the cell strainer. Rinse 4X with 10 mL of 37 &#176;C DMEM/F12 (Figure 1D). 
	CAUTION: Incomplete rinsing will result in cultures contaminated with non-epithelial cells.</li>
	<li>Release the tissue fragments by holding the strainer tab with gloved fingers, inverting the strainer over a 60 mm tissue culture dish and passing 1 mL aliquots of Maintenance Medium (see Table 1) through the bottom of the strainer 4X (Figure 1E).</li>
	<li>Check the strainer for tissue fragment remnants, which will be visible by the naked eye, and rinse the strainer 1X more with 1 mL of Maintenance Medium if any fragments are still adhering to the strainer. The rinsed tissue fragments should now be free of stromal cells.</li>
	<li>Quickly examine the 60 mm dish containing the MG fragments from protocol step 2.9 under an inverted microscope (4X or 10X objective, Figure 1F). A typical preparation of eight MGs yields ~500 fragments. Look for single cells or fat droplets and whether there are contaminating cells. 
	NOTE: If there are contaminating cells, repeat the filtration step by collecting the medium and fragments from the 60 mm dish using a 5 mL pipette and filtering the fragment again through a fresh 70 &#181;m strainer, repeating protocol steps 2.6, 2.8-2.10.</li>
	<li>Incubate 24 h at 37 &#176;C, 5% CO2, allowing the tissue fragment to adhere and generate bilayered fragments (Figure 2A). 
	NOTE:If the fragments have not settled by 24 h, continue to incubate until adhered. If the fragments do not adhere well, the separation of the cell compartments will not work. If researchers are concerned about adhesion, the tissue culture plates can be treated to promote fragment attachment (e.g., poly-L-lysine).</li>
</ol>
3. Day 3: Differential trypsinization of myoepithelial and luminal epithelial cells
<ol>
	<li>To separate MyoECs from LECs, empty the media from the dish, rinse 1x with 1 mL of DPBS and add 1 mL of fresh trypsin-EDTA (0.05%), and carefully monitor the digestion under an inverted microscope (10X or 20X objective, Figure 2B,C,F). The detachment of the outer MyoEC layer will require 3-6 min, depending on trypsin-EDTA (0.05%) strength. 
	NOTE: Please see Figure 2 for representative images showing this process. Under brightfield illumination, the MyoECs appear rounded and have a brighter appearance in comparison to the LECs, which remain adhered in the center and appear darker.</li>
	<li>Collect the MyoEC fraction in a 15 mL tube containing 2 mL 10% FBS/DPBS. Without disturbing the LECs, gently rinse the 60 mm dish with 2 mL of DPBS and then dispose of the DPBS (Figure 2H-I). 
	NOTE: The usual recovery for MyoECs is within a range of (3.5 x 106-1.5 x 106) depending on the size of the MGs.</li>
	<li>To remove the LEC fraction, add 1 mL of trypsin-EDTA (0.05%) to the dish again and incubate 7-15 min, monitoring carefully to prevent overdigestion. Quench the trypsin-EDTA (0.05%) on the dish with 2 mL of 10% FBS/DPBS. Collect the LEC fraction in a new 15 mL tube. 
	NOTE: The usual recovery for LECs is within a range (2 x 106-4.2 x 106) depending on the size of the MGs. Routinely, the purity of both fractions as assayed by immunohistochemistry is ~90%19 (Figure 2E).</li>
	<li>Centrifuge each fraction for 5 min at 300 x g to remove the trypsin-EDTA (0.05%). Resuspend the pellet in 250 &#181;L of Maintenance Medium and count each cell population using a hemocytometer or automated cell counter. Place the cells on ice while counting. 
	NOTE: If genetic manipulation of the cell fractions is desired, primary cells can be grown on low adhesion dishes and infected with lentivirus20.</li>
</ol>
4. Day 3: Combining and embedding cell fractions in an extracellular matrix
NOTE: Once the MyoEC and LEC fractions have been collected and counted, they can be combined. The typical MyoEC/LEC ratio is 1:3 (Figure 3A)19. Different studies can be performed. For example, to perform mosaic studies, fractions can be generated from wild type (WT) and mutant (Mut) mice and combined (MyoEC/LEC: WT/WT; WT/Mut; Mut/WT; Mut/Mut)21, or fractions can be combined using different ratios of MyoECs/LECs19.
<ol>
	<li>Based on cell counts, calculate the number of wells (8 well chamber, see Table of Materials) that need to be prepared for 12,000 cells/well (e.g. 3,000 MyoECs:9,000 LECs).</li>
	<li>Establish the base layer for 3D culture by adding 90 &#181;L of 50% ECM (50% ECM/50% DMEM/F12, without phenol red - see the Table of Materials) to each well. Ensure there are no bubbles and the wells are coated evenly. To solidify the base layer, incubate the slides at 37 &#176;C, 5% CO2 for 30 min. 
	NOTE: The ECM needs to stay ice-cold until this step, otherwise it will polymerize prematurely and lead to uneven base coating and polymerization. Using a base ECM enhances organoid growth in a single plane, which aids image capture. This also obtains faster organoid growth and uses less ECM22.</li>
	<li>During polymerization, prepare the cell mixes. Pellet the MyoEC and LEC fractions at 300 x g for 5 min and resuspend each cell fraction in 10% ECM/90% Growth Medium so each well has 100 &#181;L (see Table 1 for how to make Growth Medium). 
	NOTE: For ease of preparation, replicate wells using the same cell mixes. These can be combined and prepared in one tube (e.g., four wells of the same cell mix can be prepared in 400 &#181;L of 10% ECM/90% Growth Medium).</li>
	<li>Add 100 &#181;L of each cell mix in 10% ECM/90% Growth Medium to each well and allow organoids to settle for 20 min at 37 &#176;C, 5% CO2.</li>
	<li>Once the cells have settled, gently add 100 &#181;L of Growth Medium by gently pipetting down the chamber wall of each well. Incubate the slides at 37 &#176;C, 5% CO2 (see Figure 3A for the total composition of each well). 
	NOTE: The Rho Kinase inhibitor, R-spondin, and Nrg1 are factors that have been identified as important for the long-term culture of organoids grown from both primary murine mammary cells and human breast cancer cells13,14. In addition, the stem cell factors B27 and N2 extend the time that organoids can be cultured.</li>
	<li>Image cells every 24 h to track their growth and gently renew the Growth Medium every 2-3 days using 100 &#181;L/well (Figure 3B-E). 
	NOTE: Use extreme care when renewing the medium. Tilt the chamber slides to collect the medium at one corner of the wells. Remove &#8804;100 &#181;L of the medium and replenish with care to leave the ECM layer undisturbed.</li>
	<li>If researchers are interested in investigating lactation/alveologenesis, switch to Alveologenesis Medium (see Table 1) on day 5 and continue to renew the medium every 2-3 days until day 10 (Figure 3F) or beyond by passaging using recovery solution (see the Table of Materials)13. 
	NOTE: At this point, organoids can be isolated from the ECM by incubating at 4 &#176;C in 400 &#181;L of 4 &#176;C recovery solution (see Table of Materials for protocol and reagent info).</li>
</ol>
5. Day 5 or 10: Fixing and immunostaining organoids
<ol>
	<li>Remove the medium carefully by gently pipetting off the media (a bulb pipette works best). Rinse each well using 200 &#181;L of 1x Dulbecco&#39;s Phosphate Buffered Saline (DPBS, see recipes).</li>
	<li>Fix the organoids using cold (4 &#176;C) 4% (w/v) paraformaldehyde (PFA, see recipes) for 10 min at room temperature. 
	CAUTION: PFA is hazardous. Wear personal protective equipment (lab coat, gloves, and safety glasses). This step should be performed inside a fume hood. 
	NOTE: The ECM is dissolved by the PFA treatment. Incomplete ECM removal can lead to background staining when the organoids are analyzed by immunofluorescence.</li>
	<li>Remove the 4% PFA and add 200 &#181;L of 0.2% (w/v) glycine/DPBS (see Table 1) to each well. Incubate the slides at room temperature for 30 min or 4 &#176;C overnight on a rocking surface set to a slow setting. 
	NOTE: The organoids can be stored for 1-3 days in DPBS at 4 &#176;C prior to the next step.</li>
	<li>Permeabilize the organoids using DPBS + 0.25% Triton X-100 (PBST, see Table 1) for 10 min at room temperature.</li>
	<li>Block the organoids using 5% donkey serum (DS) (or other serum matching the species of the secondary antibody) in DPBS for 1 h on a rocking surface. 
	NOTE: This step can be performed overnight at 4 &#176;C on a rocking surface.</li>
	<li>Prepare primary antibodies in 1% DS/DPBS. Use 125-200 &#181;L for each well. Perform immunostaining by incubating the organoids in primary antibody overnight at 4 &#176;C on a rocking surface.</li>
</ol>
6. Day 11: Complete immunofluorescence
<ol>
	<li>Wash each well 2X with 200 &#181;L PBST for 5 min. Add secondary antibody in 1% DS/DPBS using 125-200 &#181;L for each well. Incubate at room temperature on a rocking surface for 45 min.</li>
	<li>Wash each well 2X using 200 &#181;L PBST per well.</li>
	<li>Stain the nuclei using Hoechst DNA dye in DPBS (1:2,000 in DPBS) for 5 min at room temperature.</li>
	<li>Remove all liquid left on the well by gently suctioning with a vacuum.</li>
	<li>Carefully remove the chambers and gasket, place one drop (~30 &#181;L) of mounting media (see Table of Materials) on each well and coverslip, taking care to remove bubbles. Allow the slide to dry in a dark space for 1-2 days. Seal with clear nail polish. Image the organoids on a confocal microscope (Figure 3E-F).</li>
</ol>

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The authors have nothing to disclose.

Here, a method is presented detailing how researchers can generate 3D organoid cultures using primary MG cells. The difference between this and other protocols is that we detail a method to separate the two, distinct MG cell compartments: the outer basal MyoECs and inner LECs. Our method employs a two-step trypsin-EDTA (0.05%) treatment that we call differential trypsinization19. This procedure allows researchers to isolate MyoECs and LECs without using sophisticated flow cytometry and thus can be...

The mammary gland (MG) is a tree-like, tubular epithelial structure embedded within an adipocyte rich stroma. The bilayered ductal epithelium comprises an outer, basal layer of contractile, myoepithelial cells (MyoECs) and an inner layer of luminal, secretory epithelial cells (LECs), encircling a central lumen1. During lactation when the outer MyoECs contract to squeeze milk from the inner alveolar LECs, the MG undergoes numerous changes that are under the control of growth factors (e.g., EGF and FGF) and hormones (e.g. progesterone, insulin, and prolactin). These changes cause the differentiation of specialized structures, alveoli, which synthesize and secrete milk during lactation1. The mammary epithelia can be experimentally manipulated using techniques in which either epithelial tissue fragments, cells, or even a single basal cell are transplanted into host mammary fat pads, precleared of endogenous mammary parenchyma, and allowed to grow out to reconstitute an entire, functional epithelial tree2,3,4,5. Transplantation is a powerful technique, but it is time-consuming and impossible if a mutation results in early embryonic lethality (prior to E14) that prevents the rescue of transplantable mammary anlage. Furthermore, investigators frequently wish to research the roles of the two different compartments, which are derived from lineage-restricted progenitor cells. While Cre-lox technology allows differential genetic manipulation of MyoECs and LECs, this is also a time-consuming and expensive undertaking. Thus, since the 1950s, investigators have used in vitro mammary organoids as a relatively easy and efficient way to address questions concerning mammary tissue structure and function6,7.
In early protocols describing the isolation and culture of primary mammary epithelial cells, investigators found that a basement membrane matrix (BME), composed of a plasma clot and chicken embryo extract, was required for MG fragments grown on a dish6. In the following decades, extracellular matrices (ECMs, collagen, and jellylike protein matrix secreted by Engelbreth-Holm-Swarm murine sarcoma cells) were developed to facilitate 3D culture and better mimic the in vivo environment7,8,9,10. Culturing cells in 3D matrices revealed by multiple criteria (morphology, gene expression, and hormone responsiveness) that such a microenvironment better models in vivo physiological processes9,10,11,12. Research using primary murine cells identified key growth factors and morphogens necessary for the extended maintenance and differentiation of organoids13. These studies have set the stage for the protocol presented here, and for the culture of human breast cells as 3D organoids, which is now a modern clinical tool, allowing for drug discovery and drug testing on patient samples14. Overall, organoid culturing highlights the self-organization capacities of primary cells and their contributions to morphogenesis and differentiation.
Presented here is a protocol to culture murine epithelia that can be differentiated into milk-producing acini. A differential trypsinization technique is used to isolate the MyoECs and LECs that comprise the two distinct MG cell compartments. These separated cell fractions can then be genetically manipulated to overexpress or knockdown gene function. Because lineage-intrinsic, self-organization is an innate property of mammary epithelial cells15,16,17, recombining these cell fractions allows researchers to generate bilayered, mosaic organoids. We begin by enzymatically digesting the adipose tissue, and then incubating the mammary fragments on a tissue culture dish for 24 h (Figure 1). The tissue fragments settle on polystyrene dishes as bilayered fragments with their in vivo organization: outer myoepithelial layer surrounding inner luminal layers. This cellular organization allows for the isolation of the outer MyoECs by trypsin-EDTA (0.05%) treatment for 3-6 min followed by a second round of trypsin-EDTA (0.05%) treatment that detaches the remaining inner LECs (Figure 2). Thus, these cell types with different trypsin sensitivity are isolated and can subsequently be mixed and plated in ECM (Figure 3). The cells undergo self-organization to form bilayered spheres, comprising an outer layer of MyoECs surrounding inner LECs. Lumen formation occurs as the cells grow in a medium containing a cocktail of growth factors (see recipes for Growth Medium)13. After 5 days, organoids can be differentiated into milk-producing acini by switching to Alveologenesis Medium (see recipes and Figure 3F) and incubated for another 5 days. Alternatively, organoids will continue to expand and branch in Growth Medium for at least 10 days. Organoids can be analyzed using immunofluorescence (Figure 3D-F) or released from the ECM using a recovery solution (see Table of Materials) and analyzed via other methods (e.g., immunoblot, RT-qPCR).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 ml High-Clarity Polypropylene Conical Tube (BD Falcon)</td>
			<td>Fisher Scientific</td>
			<td>352096</td>
			<td></td>
		</tr>
		<tr>
			<td>24 well ultra-low attachment plate (Corning)</td>
			<td>Fisher Scientific</td>
			<td>CLS3473-24EA</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm TC-treated Easy-Grip Style Cell Culture Dish (BD Falcon)</td>
			<td>Fisher Scientific</td>
			<td>353001</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml High-Clarity polypropylene conical tube (BD Falcon)</td>
			<td>Fisher Scientific</td>
			<td>352098</td>
			<td></td>
		</tr>
		<tr>
			<td>60 mm TC-treated Easy-Grip Style Cell Culture Dish (BD Falcon)</td>
			<td>Fisher Scientific</td>
			<td>353004</td>
			<td></td>
		</tr>
		<tr>
			<td>70 &#181;M nylon cell strainer (Corning)</td>
			<td>Fisher Scientific</td>
			<td>08-771-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic (100X)</td>
			<td>Thermo Fisher Scientific</td>
			<td>15240062</td>
			<td>Pen/Strep also works</td>
		</tr>
		<tr>
			<td>B27 supplement without vitamin A (50x)</td>
			<td>Thermo Fisher Scientific</td>
			<td>12587010</td>
			<td></td>
		</tr>
		<tr>
			<td>B6 ACTb-EGFP mice</td>
			<td>The Jackson Laboratory</td>
			<td>003291</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Insulin syringe 0.5 mL</td>
			<td>Thermo Fisher Scientific</td>
			<td>14-826-79</td>
			<td></td>
		</tr>
		<tr>
			<td>Class 2 Dispase (Roche)</td>
			<td>Millipore Sigma</td>
			<td>4942078001</td>
			<td></td>
		</tr>
		<tr>
			<td>Class 3 Collagenase</td>
			<td>Worthington Biochemical</td>
			<td>LS004206</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Cell Recovery solution</td>
			<td>Fisher Scientific</td>
			<td>354253</td>
			<td>Follow the guidelines for use &#8211; Extraction of Three-Dimensional Structures 
			from Corning Matrigel Matrix</td>
		</tr>
		<tr>
			<td>Corning Costar Ultra-Low Attachment 6-well</td>
			<td>Fisher Scientific</td>
			<td>CLS3471</td>
			<td></td>
		</tr>
		<tr>
			<td>Dexamethasone</td>
			<td>Millipore Sigma</td>
			<td>D4902-25MG</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM/F12, no phenol red</td>
			<td>Thermo Fisher Scientific</td>
			<td>11039-021</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase (Deoxyribonuclease I)</td>
			<td>Worthington Biochemical</td>
			<td>LS002007</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-Goat 647</td>
			<td>Thermo Fisher Scientific</td>
			<td>A21447</td>
			<td>Use at 1:500, Lot: 1608641, stock 2 mg/mL, RRID:AB_2535864</td>
		</tr>
		<tr>
			<td>Donkey anti-Mouse 647</td>
			<td>Jackson ImmunoResearch</td>
			<td>715-606-150</td>
			<td>Use at 1:1000, Lot: 140554, stock 1.4 mg/mL</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F12)</td>
			<td>Thermo Fisher Scientific</td>
			<td>11330-057</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s phosphate-buffered saline (DPBS)</td>
			<td>Thermo Fisher Scientific</td>
			<td>14190-250</td>
			<td>Without Mg2+/Ca2+</td>
		</tr>
		<tr>
			<td>EGF</td>
			<td>Fisher Scientific</td>
			<td>AF-100-15-100ug</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>VWR</td>
			<td>97068&#8211;085</td>
			<td>100% US Origin, premium grade, Lot: 059B18</td>
		</tr>
		<tr>
			<td>Fluoromount-G (Southern Biotech)</td>
			<td>Fisher Scientific</td>
			<td>0100-01</td>
			<td>Referred to as mounting media in text</td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15710064</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>Fisher Scientific</td>
			<td>BP381-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-WAP</td>
			<td>Santa Cruz Biotech</td>
			<td>SC-14832</td>
			<td>Use at 1:250, Lot: J1011, stock 200 &#181;g/mL, RRID:AB_677601</td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>AnaSpec</td>
			<td>AS-83218</td>
			<td>Use 1:2000, stock is 20mM</td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>Millipore Sigma</td>
			<td>I6634-100mg</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher Scientific</td>
			<td>P217-500</td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Fisher Scientific</td>
			<td>P285-500</td>
			<td></td>
		</tr>
		<tr>
			<td>KRT14&#8211;CreERtam</td>
			<td>The Jackson Laboratory</td>
			<td>5107</td>
			<td></td>
		</tr>
		<tr>
			<td>Matrigel Growth Factor Reduced (GFR); Phenol Red-Free; 10 mL</td>
			<td>Fisher Scientific</td>
			<td>CB-40230C</td>
			<td>Lot: 8204010, stock concentration 8.9 mg/mL</td>
		</tr>
		<tr>
			<td>MillexGV Filter Unit 0.22 &#181;m</td>
			<td>Millipore Sigma</td>
			<td>SLGV033RS</td>
			<td></td>
		</tr>
		<tr>
			<td>Millicell EZ SLIDE 8-well glass, sterile</td>
			<td>Millipore Sigma</td>
			<td>PEZGS0816</td>
			<td>These chamber slides are great for gasket removal but other brands can work well (e.g. Lab Tek II).</td>
		</tr>
		<tr>
			<td>Mouse anti-SMA</td>
			<td>Millipore Sigma</td>
			<td>A2547</td>
			<td>Use at 1:500, Lot: 128M4881V, stock 5.2 mg/mL, RRID:AB_476701</td>
		</tr>
		<tr>
			<td>N-2 Supplement (100x)</td>
			<td>Thermo Fisher Scientific</td>
			<td>17502048</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Fisher Scientific</td>
			<td>S671-3</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Fisher Scientific</td>
			<td>S468-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Nrg1</td>
			<td>R&#38;D</td>
			<td>5898-NR-050</td>
			<td></td>
		</tr>
		<tr>
			<td>Ovine Pituitary Prolactin</td>
			<td>National Hormone and Peptide Program</td>
			<td></td>
			<td>Purchased from Dr. Parlow at Harbor-UCLA Research and Education Institute</td>
		</tr>
		<tr>
			<td>Paraformaldahyde</td>
			<td>Millipore Sigma</td>
			<td>PX0055-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Pentobarbital</td>
			<td>Millipore Sigma</td>
			<td>P3761</td>
			<td></td>
		</tr>
		<tr>
			<td>R26R-EYFP</td>
			<td>The Jackson Laboratory</td>
			<td>6148</td>
			<td></td>
		</tr>
		<tr>
			<td>Rho inhibitor Y-27632</td>
			<td>Tocris</td>
			<td>1254</td>
			<td></td>
		</tr>
		<tr>
			<td>R-spondin</td>
			<td>Peprotech</td>
			<td>120-38</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Hydroxide</td>
			<td>Fisher Scientific</td>
			<td>S318-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile Filtered Donkey Serum</td>
			<td>Equitech-Bio Inc.</td>
			<td>SD30-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile Filtered Donkey Serum</td>
			<td>Equitech-Bio Inc.</td>
			<td>SD30-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Millipore Sigma</td>
			<td>x100-500ML</td>
			<td>Laboratory grade</td>
		</tr>
		<tr>
			<td>Trypsin EDTA 0.05%</td>
			<td>Thermo Fisher Scientific</td>
			<td>25300-062</td>
			<td></td>
		</tr>
	</tbody>
</table>

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 protocol presented here describes a method for investigating specific lineage contributions of mammary epithelial cells by making use of mosaic organoids. To obtain primary murine cells for organoids, the mammary gland epithelium must first be isolated from the surrounding adipocyte rich stroma (Figure 1). This process is described briefly here and is also described in a previously published study18. To obtain enough cells, it is r...

Watch this Scientific Journal Video about Generation of Mosaic Mammary Organoids by Differential Trypsinization at JoVE.com

Generation of Mosaic Mammary Organoids by Differential Trypsinization

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 ...

generation-of-mosaic-mammary-organoids-by-differential-trypsinization

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

6.2K Views.  University of California, Santa Cruz. 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.

Video: Generation of Mosaic Mammary Organoids by Differential Trypsinization

Mosaic Zebrafish Transgenesis for Functional Genomic Analysis of Candidate Cooperative Genes in Tumor Pathogenesis

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers. To robustly and efficiently identify oncogenic &#8220;drivers&#8221; among these tumors and define their complex relationships with concurrent genetic alterations during tumor pathogenesis remains a daunting task. Recently, zebrafish have emerged as an important animal model for studying human diseases, largely because of their ease of maintenance, high fecundity, obvious advantages for in vivo imaging, high conservation of oncogenes and their molecular pathways, susceptibility to tumorigenesis and, most importantly, the availability of transgenic techniques suitable for use in the fish. Transgenic zebrafish models of cancer have been widely used to dissect oncogenic pathways in diverse tumor types. However, developing a stable transgenic fish model is both tedious and time-consuming, and it is even more difficult and more time-consuming to dissect the cooperation of multiple genes in disease pathogenesis using this approach, which requires the generation of multiple transgenic lines with overexpression of the individual genes of interest followed by complicated breeding of these stable transgenic lines. Hence, use of a mosaic transient transgenic approach in zebrafish offers unique advantages for functional genomic analysis in vivo. Briefly, candidate transgenes can be coinjected into one-cell-stage wild-type or transgenic zebrafish embryos and allowed to integrate together into each somatic cell in a mosaic pattern that leads to mixed genotypes in the same primarily injected animal. This permits one to investigate in a faster and less expensive manner whether and how the candidate genes can collaborate with each other to drive tumorigenesis. By transient overexpression of activated ALK in the transgenic fish overexpressing MYCN, we demonstrate here the cooperation of these two oncogenes in the pathogenesis of a pediatric cancer, neuroblastoma that has resisted most forms of contemporary treatment.

The goal of this study is to demonstrate how the mosaic transgenesis strategy can be used in zebrafish to rapidly and efficiently assess the relative contributions of multiple oncogenes in tumor initiation and progression in vivo.

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers.  To robustly and efficiently ...

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

Generation of iPSC-derived Human Brain Organoids to Model Early Neurodevelopmental Disorders

The restricted availability of suitable in vitro models that can reliably represent complex human brain development is a significant bottleneck that limits the translation of basic brain research into clinical application. While induced pluripotent stem cells (iPSCs) have replaced the ethically questionable human embryonic stem cells, iPSC-based neuronal differentiation studies remain descriptive at the cellular level but fail to adequately provide the details that could be derived from a complex, 3D human brain tissue. 
This gap is now filled through the application of iPSC-derived, 3D brain organoids, "Brains in a dish," that model many features of complex human brain development. Here, a method for generating iPSC-derived, 3D brain organoids is described. The organoids can help with modeling autosomal recessive primary microcephaly (MCPH), a rare human neurodevelopmental disorder. A widely accepted explanation for the brain malformation in MCPH is a depletion of the neural stem cell pool during the early stages of human brain development, a developmental defect that is difficult to recreate or prove in vitro. 
To study MCPH, we generated iPSCs from patient-derived fibroblasts carrying a mutation in the centrosomal protein CPAP. By analyzing the ventricular zone of microcephaly 3D brain organoids, we showed the premature differentiation of neural progenitors. These 3D brain organoids are a powerful in vitro system that will be instrumental in modeling congenital brain disorders induced by neurotoxic chemicals, neurotrophic viral infections, or inherited genetic mutations.

Modeling human brain development has been hindered due to the unprecedented complexity of neural epithelial tissue. Here, a method for the robust generation of brain organoids to delineate early events of human brain development and to model microcephaly in vitro is described.

The restricted availability of suitable in vitro models that can reliably represent complex human brain development is a significant bottleneck that ...

Transfer of Mammary Gland-forming Ability Between Mammary Basal Epithelial Cells and Mammary Luminal Cells via Extracellular Vesicles/Exosomes

Cells can communicate via exosomes, ~100-nm extracellular vesicles (EVs) that contain proteins, lipids, and nucleic acids. Non-adherent/mesenchymal mammary epithelial cell (NAMEC)-derived extracellular vesicles can be isolated from NAMEC medium via differential ultracentrifugation. Based on their density, EVs can be purified via ultracentrifugation at 110,000 x g. The EV preparation from ultracentrifugation can be further separated using a continuous density gradient to prevent contamination with soluble proteins. The purified EVs can then be further evaluated using nanoparticle-tracking analysis, which measures the size and number of vesicles in the preparation. The extracellular vesicles with a size ranging from 50 to 150 nm are exosomes. The NAMEC-derived EVs/exosomes can be ingested by mammary epithelial cells, which can be measured by flow cytometry and confocal microscopy. Some mammary stem cell properties (e.g., mammary gland-forming ability) can be transferred from the stem-like NAMECs to mammary epithelial cells via the NAMEC-derived EVs/exosomes. Isolated primary EpCAMhi/CD49flo luminal mammary epithelial cells cannot form mammary glands after being transplanted into mouse fat pads, while EpCAMlo/CD49fhi basal mammary epithelial cells form mammary glands after transplantation. Uptake of NAMEC-derived EVs/exosomes by EpCAMhi/CD49flo luminal mammary epithelial cells allows them to generate mammary glands after being transplanted into fat pads. The EVs/exosomes derived from stem-like mammary epithelial cells transfer mammary gland-forming ability to EpCAMhi/CD49flo luminal mammary epithelial cells.

This protocol describes methods for purifying, quantitating, and characterizing extracellular vesicles (EVs)/exosomes from non-adherent/mesenchymal mammary epithelial cells and for using them to transfer mammary gland-forming ability to luminal mammary epithelial cells. EVs/exosomes derived from stem-like mammary epithelial cells can transfer this cell property to cells that ingest the EVs/exosomes.

Cells can communicate via exosomes, ~100-nm extracellular vesicles (EVs) that contain proteins, lipids, and nucleic acids. Non-adherent/mesenchymal ...

A Novel Mammary Fat Pad Transplantation Technique to Visualize the Vessel Generation of Vascular Endothelial Stem Cells

Endothelial cells (ECs) are the fundamental building blocks of the vascular architecture and mediate vascular growth and remodeling to ensure proper vessel development and homeostasis. However, studies on endothelial lineage hierarchy remain elusive due to the lack of tools to gain access as well as to directly evaluate their behavior in vivo. To address this shortcoming, a new tissue model to study angiogenesis using the mammary fat pad has been developed. The mammary gland develops mostly in the postnatal stages, including puberty and pregnancy, during which robust epithelium proliferation is accompanied by extensive vascular remodeling. Mammary fat pads provide space, matrix, and rich angiogenic stimuli from the growing mammary epithelium. Furthermore, mammary fat pads are located outside the peritoneal cavity, making them an easily accessible grafting site for assessing the angiogenic potential of exogenous cells. This work also describes an efficient tracing approach using fluorescent reporter mice to specifically label the targeted population of vascular endothelial stem cells (VESCs) in vivo. This lineage tracing method, coupled with subsequent tissue whole-mount microscopy, enable the direct visualization of targeted cells and their descendants, through which the proliferation capability can be quantified and the differentiation commitment can be fate-mapped. Using these methods, a population of bipotent protein C receptor (Procr) expressing VESCs has recently been identified in multiple vascular systems. Procr+ VESCs, giving rise to both new ECs and pericytes, actively contribute to angiogenesis during development, homeostasis, and injury repair. Overall, this manuscript describes a new mammary fat pad transplantation and in vivo lineage tracing techniques that can be used to evaluate the stem cell properties of VESCs.

This work demonstrates a novel approach to assess the proliferation, differentiation, and vessel-forming potential of vascular endothelial stem cells (VESCs) through mammary fat pad transplantation followed by whole-mount tissue preparation for microscopic observation. A lineage tracing strategy to investigate the behavior of VESCs in vivo is also presented.

Endothelial cells (ECs) are the fundamental building blocks of the vascular architecture and mediate vascular growth and remodeling to ensure proper ...

Generation of Standardized and Reproducible Forebrain-type Cerebral Organoids from Human Induced Pluripotent Stem Cells

The human cortex is highly expanded and exhibits a complex structure with specific functional areas, providing higher brain function, such as cognition. Efforts to study human cerebral cortex development have been limited by the availability of model systems. Translating results from rodent studies to the human system is restricted by species differences and studies on human primary tissues are hampered by a lack of tissue availability as well as ethical concerns. Recent development in human pluripotent stem cell (PSC) technology include the generation of three-dimensional (3D) self-organizing organotypic culture systems, which mimic to a certain extent human-specific brain development in vitro. Currently, various protocols are available for the generation of either whole brain or brain-region specific organoids. The method for the generation of homogeneous and reproducible forebrain-type organoids from induced PSC (iPSC), which we previously established and describe here, combines the intrinsic ability of PSC to self-organize with guided differentiation towards the anterior neuroectodermal lineage and matrix embedding to support the formation of a continuous neuroepithelium. More specifically, this protocol involves: (1) the generation of iPSC aggregates, including the conversion of iPSC colonies to a confluent monolayer culture; (2) the induction of anterior neuroectoderm; (3) the embedding of neuroectodermal aggregates in a matrix scaffold; (4) the generation of forebrain-type organoids from neuroectodermal aggregates; and (5) the fixation and validation of forebrain-type organoids. As such, this protocol provides an easily applicable system for the generation of standardized and reproducible iPSC-derived cortical tissue structures in vitro.

Cerebral organoids represent a new model system to investigate early human brain development in vitro. This article provides the detailed methodology to efficiently generate homogeneous dorsal forebrain-type organoids from human induced pluripotent stem cells including critical characterization and validation steps.

The human cortex is highly expanded and exhibits a complex structure with specific functional areas, providing higher brain function, such as cognition. ...

Real-time Measurement of Epithelial Barrier Permeability in Human Intestinal Organoids

Advances in 3D culture of intestinal tissues obtained through biopsy or generated from pluripotent stem cells via directed differentiation, have resulted in sophisticated in vitro models of the intestinal mucosa. Leveraging these emerging model systems will require adaptation of tools and techniques developed for 2D culture systems and animals. Here, we describe a technique for measuring epithelial barrier permeability in human intestinal organoids in real-time. This is accomplished by microinjection of fluorescently-labeled dextran and imaging on an inverted microscope fitted with epifluorescent filters. Real-time measurement of the barrier permeability in intestinal organoids facilitates the generation of high-resolution temporal data in human intestinal epithelial tissue, although this technique can also be applied to fixed timepoint imaging approaches. This protocol is readily adaptable for the measurement of epithelial barrier permeability following exposure to pharmacologic agents, bacterial products or toxins, or live microorganisms. With minor modifications, this protocol can also serve as a general primer on microinjection of intestinal organoids and users may choose to supplement this protocol with additional or alternative downstream applications following microinjection.

This protocol describes the measurement of epithelial barrier permeability in real-time following pharmacologic treatment in human intestinal organoids using fluorescent microscopy and live cell microscopy.

Advances in 3D culture of intestinal tissues obtained through biopsy or generated from pluripotent stem cells via directed differentiation, have resulted ...

Generation of Organoids from Mouse Extrahepatic Bile Ducts

Cholangiopathies, which affect extrahepatic bile ducts (EHBDs), include biliary atresia, primary sclerosing cholangitis, and cholangiocarcinoma. They have no effective therapeutic options. Tools to study EHBD are very limited. Our purpose was to develop an organ-specific, versatile, adult stem cell-derived, preclinical cholangiocyte model that can be easily generated from wild type and genetically engineered mice. Thus, we report on the novel technique of developing an EHBD organoid (EHBDO) culture system from adult mouse EHBDs. The model is cost-efficient, able to be readily analyzed, and has multiple downstream applications. Specifically, we describe the methodology of mouse EHBD isolation and single cell dissociation, organoid culture initiation, propagation, and long-term maintenance and storage. This manuscript also describes EHBDO processing for immunohistochemistry, fluorescent microscopy, and mRNA abundance quantitation by real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR). This protocol has significant advantages in addition to producing EHBD-specific organoids. The use of a conditioned medium from L-WRN cells significantly reduces the cost of this model. The use of mouse EHBDs provides almost unlimited tissue for culture generation, unlike human tissue. Generated mouse EHBDOs contain a pure population of epithelial cells with markers of endodermal progenitor and differentiated biliary cells. Cultured organoids maintain homogenous morphology through multiple passages and can be recovered after a long-term storage period in liquid nitrogen. The model allows for the study of biliary progenitor cell proliferation, can be manipulated pharmacologically, and may be generated from genetically engineered mice. Future studies are needed to optimize culture conditions in order to increase plating efficiency, evaluate functional cell maturity, and direct cell differentiation. Development of co-culture models and a more biologically neutral extracellular matrix are also desirable.

This protocol describes the production of a mouse extrahepatic bile duct 3-dimensional organoid system. These biliary organoids can be maintained in culture to study cholangiocyte biology. Biliary organoids express markers of both progenitor and biliary cells and are composed of polarized epithelial cells.

Cholangiopathies, which affect extrahepatic bile ducts (EHBDs), include biliary atresia, primary sclerosing cholangitis, and cholangiocarcinoma. They have ...

Generation of Multicellular Human Primary Endometrial Organoids

The human endometrium is one of the most hormonally responsive tissues in the body and is essential for the establishment of pregnancy. This tissue can also become diseased and cause morbidity and even death. Model systems to study human endometrial biology have been limited to in vitro culture systems of single cell types. In addition, the epithelial cells, one of the major cell types of the endometrium, do not propagate well or retain their physiological traits in culture, and thus our understanding of endometrial biology remains limited. We have generated, for the first time, endometrial organoids that consist of both epithelial and stromal cells of the human endometrium. These organoids do not require any exogenous scaffold materials and specifically organize so that epithelial cells encompass the spheroid-like structure and become polarized with stromal cells in the center that produce and secrete collagen. Estrogen, progesterone and androgen receptors are expressed in the epithelial and stromal cells and treatment with physiological levels of estrogen and testosterone promote the organization of the organoids. This new model system can be used to study normal endometrial biology and disease in ways that were not possible before.

A protocol to generate human primary endometrial organoids that consist of epithelial and stromal cells and retain characteristics of the native endometrial tissue is presented. This protocol describes methods from uterine tissue acquisition to the histologic processing of endometrial organoids.

The human endometrium is one of the most hormonally responsive tissues in the body and is essential for the establishment of pregnancy. This tissue can ...

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 ...