This work was funded by the National Institutes of Health, National Institute of Child Health and Human Development (NICHD) F31 HD089693, the National Institute for Environmental Health Sciences / National Center for Advancing Translational Sciences (NIEHS/NCATS) UH3TR001207 and 4UH3ES029073-03, and the Thomas J. Watkin&#8217;s Memorial Professorship.
The authors would like to thank Eric W. Roth for their assistance with transmission electron microscopy. This work made use of the BioCryo facility of Northwestern University&#8217;s NUANCE Center, which has received support from the Soft and Hybrid Nanotechnology Experimental (SHyNE) Resource (NSF ECCS-1542205); the MRSEC program (NSF DMR-1720139) at the Materials Research Center; the International Institute for Nanotechnology (IIN); and the State of Illinois, through the IIN. It also made use of the CryoCluster equipment, which has received support from the MRI program (NSF DMR-1229693). Graphics in Figure 1 were designed using BioRender.com.

All mouse experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Northwestern University, and all procedures were performed under IACUC-approved protocols.
1. Preparation of enzymatic tissue-dissociation solutions
<ol>
	<li>Use two different enzymatic solutions (Solution 1 and Solution 2), both made using a basal culture medium solution (BM).</li>
	<li>To prepare BM, add serum and penicillin-streptomycin to minimum essential medium to final concentrations of 10% and 1% respectively (see Table of Materials for specific reagents). Then sterile filter the BM through a 0.22 &#956;m filter. Before use with cells, pre-equilibrate sterile BM to a neutral pH by dispensing into a culture dish and placing within a humidified, 5% CO2 incubator at 37 &#176;C for a minimum of 1 h. 
	NOTE: BM can be stored at 4 &#176;C for up to 1 week, after which fresh BM should be made.</li>
	<li>To prepare collagenase I stock solutions, first dissolve 100 mg of collagenase I into 1 mL of sterile embryo grade H2O (final concentration 10% m/v), invert or swirl to dissolve, and store 20 &#956;L aliquots at -20 &#176;C for later use. Aliquots should be thawed only once.</li>
	<li>To prepare deoxyribonuclease I (DNase I) stock solutions, add 20 mg of DNase I into 1 mL of sterile embryo grade H2O (final concentration 2% m/v), invert or swirl to dissolve (do not vortex), and store 20 &#956;L aliquots at -20 &#176;C for later use. Aliquots should be thawed only once.</li>
	<li>For hyaluronidase stock solutions, add 30 mg into 1 mL of sterile phosphate buffered saline (PBS; final concentration 3% m/v hyaluronidase in PBS containing Ca++/Mg++), invert or swirl to dissolve, and store 100 &#956;L aliquots at -20 &#176;C for later use. Aliquots can be thawed and re-frozen several times without loss of enzymatic activity.</li>
	<li>To prepare dissociation Solution 1, add 10 &#956;L collagenase I and 10 &#956;L DNase I into 1 mL of sterile, pre-equilibrated BM (Final concentrations: 1 mg/mL collagenase I and 5 &#956;g/mL DNase I). Triturate gently with a pipette to mix the solution, and pre-warm to 37 &#176;C before use with the tissue. 
	NOTE: Solution 2 is prepared by adding 33 &#956;L of hyaluronidase (prewarmed to 37 &#176;C) per 1 mL of Solution 1, (to a final concentration of 1 mg/mL). This occurs mid-way through enzymatic dissociation of testis tissue at step 2.5 below.</li>
</ol>
2. Testis tissue dissociation
NOTE: All mice were housed within polypropylene cages and provided with food and water ad libitum. Animals were fed irradiated chow which does not contain phytoestrogens. Juvenile CD-1 mice, 5 days post-partum (dpp), were used for all experiments and anesthetized prior to euthanasia and tissue collection, within an anesthesia chamber attached to an isoflurane vaporizer (2.5 L/min in O2). Mice were confirmed for full anesthesia via the absence of a response to toe-prick, after which mice were euthanized via decapitation.
<ol>
	<li>Anesthetize mice in an isoflurane chamber, ensure anesthesia via a toe-prick, and then decapitate the mouse using a sharp scissor. Place the euthanized mouse supine on a dissection mat and sterilize the abdomen with 70% ethanol. Tent the skin of the lower abdomen with forceps and open the abdomen with scissors.</li>
	<li>Locate the testes in the lower left and right inguinal regions of the abdomen. Cut their connections to the vas deferens and any anchoring connective tissue, then lift the entire testis (with epididymis still attached) from the animal. Place testes in a Petri dish of pre-equilibrated BM.</li>
	<li>Under a dissection microscope and within a sterile field, make a small incision in the tunica albuginea on one end of each testis with either a small microdissection scissor or by tearing gently using two fine forceps.
	<ol>
		<li>Then, while holding the testis from the opposite end of the incision, gently squeeze the testis with fine forceps and push in a gentle sweeping motion towards the hole in the tunica; this will release the testicular tissue as one cohesive piece.</li>
	</ol>
	</li>
	<li>Cut the testes into smaller pieces (&#8804; 2 mm3) and place them into 1 mL of pre-warmed (37 &#176;C) dissociation Solution 1.
	<ol>
		<li>Incubate at 37 &#176;C for 10 min.</li>
		<li>For more than 10 testes, increase the total dissociation solution volume by 1 mL, ensuring a minimum of 1 mL of dissociation solution per 10 testes (e.g., 2 mL for 20 testes, 3 mL for 30 testes, etc.).</li>
		<li>Gently triturate the testis pieces 50 times (50x) in solution 1 using a P1000 pipette. Ensure that the tubules separate from one another and from interstitial tissue at this point. If clumps remain, incubate for an additional 5 min and triturate once more (50x).</li>
	</ol>
	</li>
	<li>Add 33 &#956;L of hyaluronidase stock solution (pre-warmed at 37 &#176;C, from step 1.5) per 1 mL of solution 1 dissociation mixture (containing the partially dissociated testicular tissue and tubules). After adding hyaluronidase, this is called solution 2.
	<ol>
		<li>Triturate (50x) using a P1000 and incubate at 37 &#176;C for 5 min.</li>
		<li>Triturate (50x) using a P200 pipette.</li>
		<li>Ensure that at this point no visible tubules or clumps of cells are present. If clumps persist, incubate for up to 5 more min, with further trituration using a P200 pipette (50x).</li>
	</ol>
	</li>
	<li>Quench the dissociation enzymes by adding fetal bovine serum (FBS) to 10% of the total volume of solution 2. Triturate several times using a P200 pipette to ensure no clumps remain, and filter through a 40 &#956;m cell strainer to produce a single-cell suspension.</li>
	<li>Centrifuge cells at 100 x g for 7 min, discard the supernatant, and resuspend the cells in fresh BM.</li>
	<li>Count the total and viable cell concentrations using trypan blue exclusion on a hemocytometer. Add 10 &#956;L of 1:1 diluted, cell suspension: trypan blue solution, into the hemocytometer cell counting chamber (see Table of Materials).
	<ol>
		<li>Re-centrifuge cells at 100 x g for 7 min and resuspend in fresh BM. 
		NOTE: Only use viable cells for calculating cell concentration and number. Only use cell suspensions of &#8805; 80% viability for generating organoids.</li>
		<li>Prepare the single cell suspension into cell concentrations as described in order to aliquot 280,000 cells given the volumes used in the protocol specific steps below in section 3: 2D ECM-Free &#8211; 0.56 x 106 cells/mL, 2D ECM &#8211; 0.56 x 106 cells/mL , 3D ECM-Free &#8211; 4.66 x 106 cells/mL, 3D ECM &#8211; 2.8 x 106 cells/mL. 
		NOTE: All culture experiments presented here start with 280,000 cells seeded per culture well. These numbers are matched to the representative data in Figure 1, Figure 2, Figure 3 and Figure 4.</li>
	</ol>
	</li>
</ol>
3. Preparation of organoid culture dishes and seeding of cells
NOTE: To ensure a homogenous ECM, pre-thaw frozen aliquots of ECM overnight before experimentation. ECM aliquots should be submerged within a bucket of ice within a 4 &#176;C refrigerator or cold room to guarantee a slow, gradual increase in temperature. All ECM is used at a 1:1 final dilution in BM for culture. Keep thawed ECM and 1:1 diluted ECM on ice until immediately before use, otherwise the ECM might polymerize prematurely.
<ol>
	<li>For 2D ECM-free culture, no special preparation is necessary, plate single cell suspensions (500 &#956;L of 0.56 x 106 cells/mL in BM) directly onto 4- well chamber slides, and place into a 35 &#176;C incubator for culture. 
	NOTE: Cells should adhere to the bottom of the culture dish within the first 24 h of culture and may exhibit some small 3D cell clusters within this same time.</li>
	<li>For 2D ECM culture, dispense 100 &#956;L of cold 1:1 diluted extracellular basement matrix medium&#160;into a 4 well chamber slide, ensuring the gel covers the entirety of the dish bottom.
	<ol>
		<li>Place the chamber slide in a 35 &#176;C incubator for a minimum of 30 min to allow the ECM to polymerize into a gel.</li>
		<li>Add the cell suspension (500 &#956;L of 0.56 x 106 cells/mL in BM) directly on the top of the 2D gel once it has polymerized. 
		NOTE: Cells should cluster together to form small 3D clusters within the first 24 h of culture.</li>
	</ol>
	</li>
	<li>For 3D ECM-free culture, prepare agarose 3D Petri dish inserts before starting the cell culture.
	<ol>
		<li>First, autoclave 1.5 g agarose powder in a 100 mL beaker, then add 75 mL sterile, distilled water and microwave to produce molten 2% agarose for 3D Petri dish casting.</li>
		<li>Within a sterile workspace, dispense molten agarose into the 3D Petri dish mold until the meniscus is level with the sides of the mold.</li>
		<li>Allow the agarose to cool and solidify. When solid, turn the mold upside down and gently flex repeatedly until the agarose 3D Petri dish falls free from the mold. 
		NOTE: At this point, one can prepare many agarose 3D Petri dishes and store them in sterile H2O or DPBS at 4 &#176;C for upwards of one month.</li>
		<li>Prior to culturing, place agarose 3D Petri dishes into a 24 well culture dish, and cover them with 1 mL of BM. Let the 3D Petri dishes equilibrate in BM for at least 30 min within a 37 &#176;C culture incubator. Discard the BM, and repeat the equilibration once more with 1 mL fresh BM. After equilibration of 3D Petri dishes in BM, they will appear the same color as the BM (i.e., pink).</li>
		<li>To prepare for cell seeding, remove all BM from the well and dispense 200 &#956;L of fresh BM around, but not inside the center recess of the 3D Petri dish. Also, collect any remaining BM from inside the center cell-seeding recess of the microwell insert.</li>
		<li>Dispense the single cell suspension (4.66 cells/mL in 60 &#956;L of BM) into the center recess of the agarose 3D Petri dish. Gently triturate up and down to mix cells and guarantee a single cell suspension at the start of culture.</li>
		<li>Place into in a humidified 35 &#176;C incubator for culture. The following day, remove the 200 &#956;L of BM from around the microwell insert, and replace with 1 mL of fresh BM. This will bring the liquid level above the plane of the insert, submerging the entire culture.</li>
		<li>Slowly and carefully remove/add media from outside of the agarose 3D Petri dish. The organoids should have compacted overnight, allowing them to rest at the bottom and enabling media changes to leave organoids undisturbed.</li>
	</ol>
	</li>
	<li>For 3D ECM culture, prepare a single cell suspension by combining, in equal parts, the cell suspension in BM with cold, pre-thawed ECM (final concentration = 2.8 x 106 cells/mL).
	<ol>
		<li>Immediately dispense the cell-ECM mixture into a 4 well chamber slide, ensuring the mixture covers the entire bottom of the plate.</li>
		<li>Place the chamber slides at 35 &#176;C in an incubator and allow its contents to polymerize. This should take at least 30 min. After the polymerization, add 500 &#956;L of BM on the top of the culture. 
		NOTE: Cells should have clustered together to form small 3D aggregates within the first 24 h of culture.</li>
	</ol>
	</li>
</ol>
4. Organoid maintenance
<ol>
	<li>Culture all organoid model types at 35 &#176;C. For All culture types exchange half of their media with fresh BM every 2 days. To ensure that organoids are not accidentally collected while exchanging medium, always collect media slowly from a corner of the chamber slide dish, and from an external point outside of agarose 3D Petri dishes. All media can be stored at -20 &#176;C for use with immunoassays or other analyses later (i.e., quantification of secreted reproductive hormones or cytokines).</li>
	<li>After 7 days in culture, use BM containing follicle stimulating hormone (final concentration 20 mIU/mL) and human chorionic gonadotropin (final concentration 4.5 IU/mL). This applies to all organoid culture types.</li>
	<li>Routinely image all organoid cultures (i.e., time-lapse imaging) for characterizing organoid formation and quantifying metrics of self-assembly, development, and growth over time.</li>
</ol>
5. Organoid Collection
NOTE: All organoids can be fixed with 4% paraformaldehyde in PBS for downstream immunolabeling and histological analyses. Fix for 2 h at room temperature with rotation, or overnight at 4 &#176;C.
<ol>
	<li>For 2D ECM-free cultures, first rinse the sample with fresh PBS, and then add fixative directly on top of the adhered constructs.</li>
	<li>For ECM (2D and 3D) culture methods, rinse once with PBS, and then either add fixative directly on the top of the ECM-organoid sample (to fix the ECM gel and organoids together), or alternatively, gently pipette the organoids up and down to free them from the surrounding ECM, and transfer to a separate tube for fixation.</li>
	<li>For 3D ECM-free culture, gently pipette the organoids up and down within the center recess of the agarose 3D Petri dish; this will flush the organoids out facilitating their easy collection with a pipette. Then transfer organoids to a separate tube for fixation.</li>
	<li>Before processing into paraffin, embed many organoids (&#8805; 20) within a small volume (~ 30 &#956;L) of tissue processing gel; this helps orient and concentrate organoids into a small area within paraffin blocks, facilitating easier observation when sectioning and easier visual identification within paraffin sections. 
	NOTE: Organoids can be challenging to identify after paraffin embedding and sectioning upon a microtome.</li>
</ol>

The authors have nothing to disclose.

With the completion of this organoid generation protocol, the user will have four different culture techniques available to them for assembling testicular constructs and organoids in either ECM or ECM-free environments. Importantly, all four methods allow the researcher to non-invasively observe organoid self-assembly over time through time-lapse imaging or video recording, and to noninvasively collect conditioned media for analysis of secreted hormones and cytokines, without disturbing tissues during culture. In all met...

Testicular organoids are a pioneering technique for studying testicular development, spermatogenesis, and physiology in vitro1,2,3,4. Several methods have been explored for organoid generation; these include a variety of extracellular matrix (ECM) and ECM-free culture systems, in both two-dimensional (2D) and three-dimensional (3D) orientations. Different generation methods can promote distinct cellular assembly strategies; this results in a high level of morphological and functional variability between published organoid models. The purpose of this article is to discuss the current state of in vitro testicular models, and to serve as a template for future investigators, when designing testicular organoid experiments. Within the present study, four different culture system archetypes are defined and characterized in experimental process and biological outcome. These include: 2D ECM-Free, 2D ECM, 3D ECM-Free, and 3D ECM culture methods. The strategies presented herein are intended to be simple, accessible, and highly reproducible between different laboratories and research groups.
Historically for the testis, the designation &#8220;in vitro&#8221;, has been used for several different culture methods of testicular tissues and cells. These include organotypic tissue/organ culture methods (i.e., explant culture)5, isolated seminiferous tubule culture6, testicular cell culture7, and methods of de novo tissue morphogenesis (i.e., biological constructs and organoids)1. The first investigations into in vitro spermatogenesis were performed approximately 100 years ago, with the culture of rabbit testis explants in 19208, and later in 1937 with mouse explants9. Within these initial experiments spermatogonia were observed to largely degenerate across the first week of culture, though some meiotically differentiating cells were identified. Reminiscent of these historical reports, testis explant culture was revived and optimized in 2011 to become a feasible technique for studying the testis10. Since 2011, explant culture has produced fertility competent sperm in multiple reports11,12,13. Yet, due to explant culture&#8217;s reliance upon pre-existing native testis tubules, these recent advances are more accurately described as examples of &#8220;ex vivo&#8221; testicular function and spermatogenesis, tissue function that was maintained or resumed upon removal from an organism&#8217;s body. Despite its prevalence in the literature, long-term germ cell maintenance and differentiation within testicular explants is challenging to replicate14,15,16,17,18, especially over timeframes long enough to fully observe in vitro spermatogenesis (~35 days in mice19 and 74 in humans20). It is intriguing to appreciate that many of the same challenges experienced 100 years ago, are still experienced within ex vivo spermatogenesis today.
Different than ex vivo approaches, testicular organoids are de novo assembled microtissues generated entirely in vitro from cellular sources (i.e., primary testicular cells). Testicular organoids provide a creative strategy to circumvent the field&#8217;s historical reliance upon pre-existing native tissue, and to recapitulate testicular biology completely in vitro. There are multiple requirements shared by most organoid tissue models; these include (1) in vivo-mimetic tissue morphology or architecture, (2) multiple major cell types of the represented tissue, (3) self-assembly or self-organization in their generation, and (4) the ability to simulate some level of the represented tissue&#8217;s function and physiology21,22,23,24. For the testis, this can be captured in four major hallmarks: (1) the inclusion of major testicular cell types, germ, Sertoli, Leydig, peritubular, and other interstitial cells, (2) cell-directed tissue assembly, (3) appropriately-compartmentalized cell types into separate tubular compartments (germ and Sertoli) and interstitial regions (all other cell types), and (4) some degree of tissue function (e.g., reproductive hormone secretion or tissue responses, and germ cell maintenance and differentiation). Considering the historic challenges in maintaining germ cell differentiation ex vivo and in vitro, the recapitulation of in vivo-mimetic testicular architectures (i.e., structures resembling seminiferous tubules) with additional markers suggesting simulation of testicular physiology (e.g., endocrine function), are priority milestones towards generating organoids which might one day sustain in vitro spermatogenesis.
The majority of published testicular organoid methods take advantage of commercially available ECM (e.g., collagen or proprietary ECM formulations)25,26,27 or custom-sourced ECMs (i.e., decellularized testis ECM-derived hydrogels)28,29,30. Exogenous ECM promotes de novo tissue formation through providing an assembly-supportive scaffold for tissue generation. ECM methods have afforded an impressive level of tissue formation, including some germ cell presence and tissue-mimetic morphology25,28. However, the ECMs they utilize are not always universally available (i.e., decellularized ECM-derived hydrogels), and some methods require sophisticated gel and cell seeding orientations (e.g., 3-layer gradients of ECM and 3D printing)25,31,32. Scaffold-free methods (e.g., hanging drop and nonadherent culture plates)33,34,35 have also generated robust and highly reproducible organoids without the need of ECM gels or scaffolds. However, the tissue morphology of these scaffold-free organoids is often dissimilar to in vivo testes, and most of these reports incorporate a biochemical ECM additive to promote tissue formation33,34,36, or alternatively, rely upon centrifugation for forced cell aggregation and compaction34, making them less ideal for studying cell-directed migration and self-organization.
The four organoid generation methods presented in this manuscript include both ECM-dependent and independent strategies, each using simple cell seeding that enables the observation of cell-driven organoid self-assembly. All four techniques can be performed from the same cell suspensions or can make use of custom and cell-type enriched populations. A strength of these methods is the ability to observe organoids self-assemble in real-time, and to directly compare how testicular structures self-assemble between different culture microenvironments. The phenotypic differences between these four culture methods should be considered for their impact on the research question or subject of the investigator. Each method produces biological constructs or organoids within 24 h or less. In conclusion, the methods presented here provide a toolkit of organoid assembly techniques for studying testicular organoid assembly, tissue development, and testicular physiology in vitro.

<table>
	<tbody>
		<tr>
			<td>0.22 um Media Sterile Filters</td>
			<td>Millipore Sigma</td>
			<td>scgpu05re</td>
			<td>For sterile filtering media</td>
		</tr>
		<tr>
			<td>3&#946;HSD primary antibody</td>
			<td>Cosmo Bio Co</td>
			<td>K0607</td>
			<td>Leydig cell marker, 1:500 dilution</td>
		</tr>
		<tr>
			<td>AlexaFluor 568 &#945;-Mouse</td>
			<td>Thermo Fisher Scientific</td>
			<td>A-21202</td>
			<td>Fluorescence-tagged secondary antibody</td>
		</tr>
		<tr>
			<td>AlexaFluor 568 &#945;-Rabbit</td>
			<td>Thermo Fisher Scientific</td>
			<td>A10042</td>
			<td>Fluorescence-tagged secondary antibody</td>
		</tr>
		<tr>
			<td>Alpha Minimum Essential Medium</td>
			<td>Thermo Fisher Scientific</td>
			<td>11-095-080</td>
			<td>Base of culture media</td>
		</tr>
		<tr>
			<td>Collagenase I</td>
			<td>Worthington Bio</td>
			<td>LS004197</td>
			<td>For dissociation solution 1</td>
		</tr>
		<tr>
			<td>Corning Matrigel Membrane Matrix, LDEV-free</td>
			<td>Corning</td>
			<td>354234</td>
			<td>Extracellular matrix used for casting 2D and 3D ECM culture gels</td>
		</tr>
		<tr>
			<td>Countess Cell counter</td>
			<td>Thermo Fisher Scientific</td>
			<td>C10227</td>
			<td>Autmated cell counter (hemacytometer machine)</td>
		</tr>
		<tr>
			<td>Countess Cell Counting Chamber Slides</td>
			<td>Thermo Fisher Scientific</td>
			<td>C10228</td>
			<td>Hemacytometer slide for use with Countess automated counter</td>
		</tr>
		<tr>
			<td>DDX4 primary antibody</td>
			<td>Abcam</td>
			<td>138540</td>
			<td>Spermatogonia marker, 1:500 dilution</td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I (2,280 u/mgDW)</td>
			<td>Worthington Bio</td>
			<td>LS002140</td>
			<td>For dissociation solution 1</td>
		</tr>
		<tr>
			<td>DPBS 1X, + CaCl + MgCl</td>
			<td>Thermo Fisher Scientific</td>
			<td>14040182</td>
			<td>For reconstituting Hyaluronidase</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Phosphate Buffered Saline +Ca/+Mg</td>
			<td>Thermo Fisher Scientific</td>
			<td>14040117</td>
			<td>PBS</td>
		</tr>
		<tr>
			<td>Embryo Grade H2O</td>
			<td>MIllipore Sigma</td>
			<td>W1503</td>
			<td>For reconstituting Collagenase I and Dnase I</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Thermo Fisher Scientific</td>
			<td>16000044</td>
			<td>For quencing enzyme dissocation solutions</td>
		</tr>
		<tr>
			<td>Follicle stimulating hormone</td>
			<td>Abcam</td>
			<td>ab51888</td>
			<td>For long-term organoid culture</td>
		</tr>
		<tr>
			<td>Human chorionic gonadotropin</td>
			<td>Millipore Sigma</td>
			<td>C1063</td>
			<td>For long-term organoid culture</td>
		</tr>
		<tr>
			<td>Hyaluronidase, from bovine testes</td>
			<td>Millipore Sigma</td>
			<td>H4272</td>
			<td>For dissociation solution 2</td>
		</tr>
		<tr>
			<td>Inhibin B Enzyme-linked Immunosorbent Assay</td>
			<td>Ansh Labs</td>
			<td>AL-107</td>
			<td>Inhibin B ELISA Kit</td>
		</tr>
		<tr>
			<td>KnockOut Serum Replacement</td>
			<td>Thermo Fisher Scientific</td>
			<td>10828-028</td>
			<td>Serum source for Basal media</td>
		</tr>
		<tr>
			<td>MicroTissues 3D Petri Dish micro-mold spheroids (24-35, 5x7 array)</td>
			<td>Millipore Sigma</td>
			<td>Z764051</td>
			<td>For 3D ECM-Free organoid fabrication</td>
		</tr>
		<tr>
			<td>Nunc, Lab Tek II Chamber Slide System, 4-well</td>
			<td>Thermo Fisher Scientific</td>
			<td>12-565-7</td>
			<td>For 2D ECM-free, and 2D, 3D ECM culture</td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15-140-122</td>
			<td>Antibiotic for media</td>
		</tr>
		<tr>
			<td>Richard-Allan Scientific; Histogel, Specimen processing gel</td>
			<td>Thermo Fisher Scientific</td>
			<td>HG-4000-012</td>
			<td>For aiding paraffin embedding</td>
		</tr>
		<tr>
			<td>SOX9 primary antibody</td>
			<td>Millipore Sigma</td>
			<td>AB5535</td>
			<td>Sertoli Marker, 1:500 dilution</td>
		</tr>
		<tr>
			<td>Tedklad Global Mouse Chow (Breeder)</td>
			<td>Teklad Global</td>
			<td>2920</td>
			<td>Mouse food without phytoestrogens</td>
		</tr>
		<tr>
			<td>Tedklad Global Mouse Chow (Maintenance)</td>
			<td>Teklad Global</td>
			<td>2916</td>
			<td>Mouse food without phytoestrogens</td>
		</tr>
		<tr>
			<td>Testosterone Enzyme-linked Immunosorbent Assay</td>
			<td>Calbiotech</td>
			<td>TE373S</td>
			<td>Testosterone ELISA Kit</td>
		</tr>
		<tr>
			<td>Trypan Blue Solution, 0.4%</td>
			<td>Thermo Fisher Scientific</td>
			<td>15250061</td>
			<td>For cell counting</td>
		</tr>
		<tr>
			<td>&#945;SMA primary antibody</td>
			<td>Millipore Sigma</td>
			<td>A2547</td>
			<td>Peritubular marker, 1:500 dilution</td>
		</tr>
		<tr>
			<td>&#946;Catenin primary antibody</td>
			<td>BD Biosciences</td>
			<td>610154</td>
			<td>Sertoli Cytoplasm marker, 1:100 dilution</td>
		</tr>
	</tbody>
</table>

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.

Organoid generation was considered unsuccessful if testicular cells did not self-assemble within 72 h of culture, however, all methods presented here assemble within 24 h when using juvenile (5 dpp) murine cells. Failure of biological construct generation presented as a continuation of freely suspended cells (0 h column in Figure 1) even after extended culture (72 h). In the absence of tissue self-assembly, any apparent cell clusters easily dispersed into individual cells upon even gentle ma...

Watch this Scientific Journal Video about Extra Cellular Matrix-Based and Extra Cellular Matrix-Free Generation of Murine Testicular Organoids at JoVE.com

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

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 ...

extra-cellular-matrix-based-extra-cellular-matrix-free-generation

Research

JoVE Journal

Developmental Biology

6.7K Views.  Northwestern University. 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.

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

Ex Utero Electroporation and Organotypic Slice Culture of Mouse Hippocampal Tissue

Mouse genetics offers a powerful tool determining the role of specific genes during development. Analyzing the resulting phenotypes by immunohistochemical and molecular methods provides information of potential target genes and signaling pathways. To further elucidate specific regulatory mechanisms requires a system allowing the manipulation of only a small number of cells of a specific tissue by either overexpression, ablation or re-introduction of specific genes and follow their fate during development. To achieve this ex utero electroporation of hippocampal structures, especially the dentate gyrus, followed by organotypic slice culture provides such a tool. Using this system to generate mosaic deletions allows determining whether the gene of interest regulates cell-autonomously developmental processes like progenitor cell proliferation or neuronal differentiation. Furthermore it facilitates the rescue of phenotypes by re-introducing the deleted gene or its target genes. In contrast to in utero electroporation the ex utero approach improves the rate of successfully targeting deeper layers of the brain like the dentate gyrus. Overall ex utero electroporation and organotypic slice culture provide a potent tool to study regulatory mechanisms in a semi-native environment mirroring endogenous conditions.

Ex Utero Electroporation and Organotypic Slice Culture of Mouse Hippocampal Tissue

Here we present a protocol providing a tool to examine regulatory mechanisms of specific genes during hippocampal development. Employing ex utero electroporation and organotypic slice culture allows the up- and down-regulation of the expression of genes of interest in single cells and follow their fate during development.

Mouse genetics offers a powerful tool determining the role of specific genes during development. Analyzing the resulting phenotypes by immunohistochemical ...

Generation of Genetically Modified Organotypic Skin Cultures Using Devitalized Human Dermis

Organotypic cultures allow the reconstitution of a 3D environment critical for cell-cell contact and cell-matrix interactions which mimics the function and physiology of their in vivo tissue counterparts. This is exemplified by organotypic skin cultures which faithfully recapitulates the epidermal differentiation and stratification program. Primary human epidermal keratinocytes are genetically manipulable through retroviruses where genes can be easily overexpressed or knocked down. These genetically modified keratinocytes can then be used to regenerate human epidermis in organotypic skin cultures providing a powerful model to study genetic pathways impacting epidermal growth, differentiation, and disease progression. The protocols presented here describe methods to prepare devitalized human dermis as well as to genetically manipulate primary human keratinocytes in order to generate organotypic skin cultures. Regenerated human skin can be used in downstream applications such as gene expression profiling, immunostaining, and chromatin immunoprecipitations followed by high throughput sequencing. Thus, generation of these genetically modified organotypic skin cultures will allow the determination of genes that are critical for maintaining skin homeostasis.

The goal of this paper is to provide a comprehensive and detailed protocol on how to generate genetically modified human organotypic skin from epidermal keratinocytes and devitalized human dermis.

Organotypic cultures allow the reconstitution of a 3D environment critical for cell-cell contact and cell-matrix interactions which mimics the function ...

Isolation of Murine Embryonic Hemogenic Endothelial Cells

The specification of hemogenic endothelial cells from embryonic vascular endothelium occurs during brief developmental periods within distinct tissues, and is necessary for the emergence of definitive HSPC from the murine extra embryonic yolk sac, placenta, umbilical vessels, and the embryonic aorta-gonad-mesonephros (AGM) region. The transient nature and small size of this cell population renders its reproducible isolation for careful quantification and experimental applications technically difficult. We have established a fluorescence-activated cell sorting (FACS)-based protocol for simultaneous isolation of hemogenic endothelial cells and HSPC during their peak generation times in the yolk sac and AGM. We demonstrate methods for dissection of yolk sac and AGM tissues from mouse embryos, and we present optimized tissue digestion and antibody conjugation conditions for maximal cell survival prior to identification and retrieval via FACS. Representative FACS analysis plots are shown that identify the hemogenic endothelial cell and HSPC phenotypes, and describe a methylcellulose-based assay for evaluating their blood forming potential on a clonal level.

Hematopoietic stem and progenitor cells (HSPC) derive from specialized (hemogenic) endothelial cells during development, yet little is known about the process by which some endothelial cells specify to become blood forming. We demonstrate a flow-cytometry based method allowing simultaneous isolation of hemogenic endothelial cells and HSPC from murine embryonic tissues.

The specification of hemogenic endothelial cells from embryonic vascular endothelium occurs during brief developmental periods within distinct tissues, ...

Evaluation of Injury-induced Senescence and In Vivo Reprogramming in the Skeletal Muscle

Cellular senescence is a stress response that is characterized by a stable cellular growth arrest, which is important for many physiological and pathological processes, such as cancer and ageing. Recently, senescence has also been implicated in tissue repair and regeneration. Therefore, it has become increasingly critical to identify senescent cells in vivo. Senescence-associated &#946;-galactosidase (SA-&#946;-Gal) assay is the most widely used assay to detect senescent cells both in culture and in vivo. This assay is based on the increased lysosomal contents in the senescent cells, which allows the histochemical detection of lysosomal &#946;-galactosidase activity at suboptimum pH (6 or 5.5). In comparison with other assays, such as flow cytometry, this allows the identification of senescent cells in their resident environment, which offers valuable information such as the location relating to the tissue architecture, the morphology, and the possibility of coupling with other markers via immunohistochemistry&#160;(IHC). The major limitation of the SA-&#946;-Gal assay is the requirement of fresh or frozen samples.
Here, we present a detailed protocol to understand how cellular senescence promotes cellular plasticity and tissue regeneration in vivo. We use SA-&#946;-Gal to detect senescent cells in the skeletal muscle upon injury, which is a well-established system to study tissue regeneration. Moreover, we use&#160;IHC to detect Nanog, a marker of pluripotent stem cells, in a transgenic mouse model. This protocol enables us to examine and quantify cellular senescence in the context of induced cellular plasticity and in vivo reprogramming.

Evaluation of Injury-induced Senescence and In Vivo Reprogramming in the Skeletal Muscle

Here we present a detailed protocol to detect both senescent and pluripotent stem cells in the skeletal muscle upon injury while inducing in vivo reprogramming. This method is suitable for evaluating the role of cellular senescence during tissue regeneration and reprogramming in vivo.

Cellular senescence is a stress response that is characterized by a stable cellular growth arrest, which is important for many physiological and ...

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

In order to support the clinical application of human adipose-derived mesenchymal stem cell (haMSC) therapy for knee osteoarthritis (KOA), we examined the efficacy of cell persistence and biodistribution of haMSCs in animal models. We demonstrated a method to label the cell membrane of haMSCs with lipophilic fluorescent dye. Subsequently, intra-articular injection of the labeled cells in rats with surgically induced KOA was monitored dynamically by an in vivo imaging system. We employed the lipophilic carbocyanines DiD (DilC18 (5)), a far-red fluorescent Dil (dialkylcarbocyanines) analog, which utilized a red laser to avoid excitation of the natural green autofluorescence from surrounding tissues. Furthermore, the red-shifted emission spectra of DiD allowed deep-tissue imaging in live animals and the labeling procedure caused no cytotoxic effects or functional damage to haMSCs. This approach has been shown to be an efficient tracking method for haMSCs in a rat KOA model. The application of this method could also be used to determine the optimal administration route and dosage of MSCs from other sources in pre-clinical studies.

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

This protocol describes an efficient way to monitor the cell persistence and biodistribution of human adipose-derived mesenchymal stem cells (haMSCs) by far-red fluorescence labeling in a rat knee osteoarthritis (KOA) model via intra-articular (IA) injection.

In order to support the clinical application of human adipose-derived mesenchymal stem cell (haMSC) therapy for knee osteoarthritis (KOA), we examined the ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Generation of Scaffold-free, Three-dimensional Insulin Expressing Pancreatoids from Mouse Pancreatic Progenitors In Vitro

The pancreas is a complex organ composed of many different cell types that work together to regulate blood glucose homeostasis and digestion. These cell types include enzyme-secreting acinar cells, an arborized ductal system responsible for the transportation of enzymes to the gut, and hormone-producing endocrine cells. 
Endocrine beta-cells are the sole cell type in the body that produce insulin to lower blood glucose levels. Diabetes, a disease characterized by a loss or the dysfunction of beta-cells, is reaching epidemic proportions. Thus, it is essential to establish protocols to investigate beta-cell development that can be used for screening purposes to derive the drug and cell-based therapeutics. While the experimental investigation of mouse development is essential, in vivo studies are laborious and time-consuming. Cultured cells provide a more convenient platform for screening; however, they are unable to maintain the cellular diversity, architectural organization, and cellular interactions found in vivo. Thus, it is essential to develop new tools to investigate pancreatic organogenesis and physiology.
Pancreatic epithelial cells develop in the close association with mesenchyme from the onset of organogenesis as cells organize and differentiate into the complex, physiologically competent adult organ. The pancreatic mesenchyme provides important signals for the endocrine development, many of which are not well understood yet, thus difficult to recapitulate during the in vitro culture. Here, we describe a protocol to culture three-dimensional, cellular complex mouse organoids that retain mesenchyme, termed pancreatoids. The e10.5 murine pancreatic bud is dissected, dissociated, and cultured in a scaffold-free environment. These floating cells self-assemble with mesenchyme enveloping the developing pancreatoid and a robust number of endocrine beta-cells developing along with the acinar and the duct cells. This system can be used to study the cell fate determination, structural organization, and morphogenesis, cell-cell interactions during organogenesis, or for the drug, small molecule, or genetic screening.

Generation of Scaffold-free, Three-dimensional Insulin Expressing Pancreatoids from Mouse Pancreatic Progenitors In Vitro

Here, we present a protocol to generate insulin expressing 3D murine pancreatoids from free-floating e10.5 dissociated pancreatic progenitors and the associated mesenchyme.

The pancreas is a complex organ composed of many different cell types that work together to regulate blood glucose homeostasis and digestion. These cell ...

Induction and Validation of Cellular Senescence in Primary Human Cells

Cellular senescence is a state of permanent cell cycle arrest activated in response to different damaging stimuli. Activation of cellular senescence is a hallmark of various pathophysiological conditions including tumor suppression, tissue remodeling and aging. The inducers of cellular senescence in vivo are still poorly characterized. However, a number of stimuli can be used to promote cellular senescence ex vivo. Among them, most common senescence-inducers are replicative exhaustion, ionizing and non-ionizing radiation, genotoxic drugs, oxidative stress, and demethylating and acetylating agents. Here, we will provide detailed instructions on how to use these stimuli to induce fibroblasts into senescence. This protocol can easily be adapted for different types of primary cells and cell lines, including cancer cells. We also describe different methods for the validation of senescence induction. In particular, we focus on measuring the activity of the lysosomal enzyme Senescence-Associated &#946;-galactosidase (SA-&#946;-gal), the rate of DNA synthesis using 5-ethynyl-2&#39;-deoxyuridine (EdU) incorporation assay, the levels of expression of the cell cycle inhibitors p16 and p21, and the expression and secretion of members of the Senescence-Associated Secretory Phenotype (SASP). Finally, we provide example results and discuss further applications of these protocols.

Here, we discuss a series of protocols for induction and validation of cellular senescence in cultured cells. We focus on different senescence-inducing stimuli and describe the quantification of common senescence-associated markers. We provide technical details using fibroblasts as a model, but the protocols can be adapted to various cellular models.

Cellular senescence is a state of permanent cell cycle arrest activated in response to different damaging stimuli. Activation of cellular senescence is a ...

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 ...