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Developmental Biology

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

doi: 10.3791/61403 Published: October 7, 2020
Maxwell E Edmonds1, Micah D. Forshee1, Teresa K. Woodruff1

Abstract

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.

Introduction

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 “in vitro”, 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’s reliance upon pre-existing native testis tubules, these recent advances are more accurately described as examples of “ex vivo” testicular function and spermatogenesis, tissue function that was maintained or resumed upon removal from an organism’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’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’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.

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Protocol

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

  1. Use two different enzymatic solutions (Solution 1 and Solution 2), both made using a basal culture medium solution (BM).
  2. 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 μm 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 °C for a minimum of 1 h.
    NOTE: BM can be stored at 4 °C for up to 1 week, after which fresh BM should be made.
  3. 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 μL aliquots at -20 °C for later use. Aliquots should be thawed only once.
  4. 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 μL aliquots at -20 °C for later use. Aliquots should be thawed only once.
  5. 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 μL aliquots at -20 °C for later use. Aliquots can be thawed and re-frozen several times without loss of enzymatic activity.
  6. To prepare dissociation Solution 1, add 10 μL collagenase I and 10 μL DNase I into 1 mL of sterile, pre-equilibrated BM (Final concentrations: 1 mg/mL collagenase I and 5 μg/mL DNase I). Triturate gently with a pipette to mix the solution, and pre-warm to 37 °C before use with the tissue.
    NOTE: Solution 2 is prepared by adding 33 μL of hyaluronidase (prewarmed to 37 °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.

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.

  1. 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.
  2. 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.
  3. 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.
    1. 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.
  4. Cut the testes into smaller pieces (≤ 2 mm3) and place them into 1 mL of pre-warmed (37 °C) dissociation Solution 1.
    1. Incubate at 37 °C for 10 min.
    2. 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.).
    3. 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).
  5. Add 33 μL of hyaluronidase stock solution (pre-warmed at 37 °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.
    1. Triturate (50x) using a P1000 and incubate at 37 °C for 5 min.
    2. Triturate (50x) using a P200 pipette.
    3. 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).
  6. 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 μm cell strainer to produce a single-cell suspension.
  7. Centrifuge cells at 100 x g for 7 min, discard the supernatant, and resuspend the cells in fresh BM.
  8. Count the total and viable cell concentrations using trypan blue exclusion on a hemocytometer. Add 10 μL of 1:1 diluted, cell suspension: trypan blue solution, into the hemocytometer cell counting chamber (see Table of Materials).
    1. 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 ≥ 80% viability for generating organoids.
    2. 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 – 0.56 x 106 cells/mL, 2D ECM – 0.56 x 106 cells/mL , 3D ECM-Free – 4.66 x 106 cells/mL, 3D ECM – 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.

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 °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.

  1. For 2D ECM-free culture, no special preparation is necessary, plate single cell suspensions (500 μL of 0.56 x 106 cells/mL in BM) directly onto 4- well chamber slides, and place into a 35 °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.
  2. For 2D ECM culture, dispense 100 μL of cold 1:1 diluted extracellular basement matrix medium into a 4 well chamber slide, ensuring the gel covers the entirety of the dish bottom.
    1. Place the chamber slide in a 35 °C incubator for a minimum of 30 min to allow the ECM to polymerize into a gel.
    2. Add the cell suspension (500 μL 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.
  3. For 3D ECM-free culture, prepare agarose 3D Petri dish inserts before starting the cell culture.
    1. 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.
    2. Within a sterile workspace, dispense molten agarose into the 3D Petri dish mold until the meniscus is level with the sides of the mold.
    3. 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 °C for upwards of one month.
    4. 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 °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).
    5. To prepare for cell seeding, remove all BM from the well and dispense 200 μL 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.
    6. Dispense the single cell suspension (4.66 cells/mL in 60 μL 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.
    7. Place into in a humidified 35 °C incubator for culture. The following day, remove the 200 μL 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.
    8. 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.
  4. 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).
    1. Immediately dispense the cell-ECM mixture into a 4 well chamber slide, ensuring the mixture covers the entire bottom of the plate.
    2. Place the chamber slides at 35 °C in an incubator and allow its contents to polymerize. This should take at least 30 min. After the polymerization, add 500 μL 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.

4. Organoid maintenance

  1. Culture all organoid model types at 35 °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 °C for use with immunoassays or other analyses later (i.e., quantification of secreted reproductive hormones or cytokines).
  2. 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.
  3. 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.

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 °C.

  1. For 2D ECM-free cultures, first rinse the sample with fresh PBS, and then add fixative directly on top of the adhered constructs.
  2. 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.
  3. 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.
  4. Before processing into paraffin, embed many organoids (≥ 20) within a small volume (~ 30 μL) 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.

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Representative Results

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 manipulation (i.e., pipetting). Successfully generated tissues were initially observed as 3D cell “clusters” (yellow arrows in 6 h column of Figure 1). Within ECM-free environments (2D and 3D), these constructs visibly appeared to “compact” across the first 24 h of culture, especially when in 3D agarose Petri dishes (Figure 1A,C). In ECM environments (2D and 3D) cell clusters possessed clear margins between the cluster and their surrounding environment (Figure 1B,D). Cell clusters were also observed to migrate across the ECM and fuse together forming larger clusters (red arrows in Figure 1B). The time required to appreciate separate self-assembled structures was measured, and there was no significant difference in time required between both 2D and 3D ECM-free conditions and 2D ECM, however, 3D ECM culture required significantly more time to assemble de novo structures than all other culture methods (Figure 1E). 2D ECM-free and 3D ECM culture generated cell clusters with significantly smaller sizes than 2D ECM and 3D ECM-free culture methods; 3D ECM-free culture produced the largest clusters with a single large and compact cluster within each well of the 3D agarose Petri dish (Figure 1C,F). In summary, these data demonstrate the ease with which to produce testicular biological constructs from juvenile mouse primary cells in four archetypal culture environments and highlight different cell-directed assembly-phenotypes within these different culture environments.

A goal of all organoid models is to recapitulate an inner morphology mimetic of native tissue. To assess for this outcome, biological constructs assembled within each culture condition were cultured for 72 h and then probed for cell-specific markers and visualized with immunofluorescence (Figure 2). Variability in tissue morphology was observed between different culture methods. 2D ECM-free organoids presented as clusters of Sertoli cells (SOX9 and βCatenin) with some germ cells (DDX4, a pan germ cell marker) adhered on top of a 2D basal confluent layer containing many somatic cells, including Sertoli, peritubular (αSMA), and Leydig cells (3βHSD) (Figure 2A-D). Note that Figure 2B-D are epifluorescent images of the entire 2D ECM-free sample, not a 5 μm section; this enables visualization of both the basal somatic cell layer and the superiorly oriented aggregates of Sertoli and germ cells. In contrast, 2D ECM culture presented with a largely different phenotype, as clear biological structures with distinct borders were easily discerned on top of the basal ECM gel (Figure 2E). These structures possessed a complex inner morphology with de novo compartmentalization of tubule vs. interstitial cell types of the testis, and so were deemed successful organoids. Tubular regions contained Sertoli, peritubular, and germ cells, and interstitial regions contained Leydig, peritubular cells and non-labeled cells (Figure 2F-H). Similarly, 3D ECM-free organoids also possessed a compartmentalized inner morphology and so were deemed successful organoids. In particular, 3D ECM-free organoids were distinguished by tubular regions containing peritubular cells that specifically oriented around groups of Sertoli and germ cells with high fidelity (Figure 2I-L). 3D ECM assembled structures did not possess a sophisticated morphology, but instead were observed to be clusters of Sertoli cells, with occasional germ and Leydig cells. In these samples, many Sertoli and un-labeled cells remained suspended in between organoids in a “stroma-like” orientation (Figure 2M-P). Together, these data highlight the variability in morphological phenotypes that different organoid generation methods produce and supports the use of two specific organoid assembly environments, 2D ECM and 3D ECM-free, for the generation of testicular organoids with an inner morphology highly mimetic of testicular compartmentalization.

To further assess testicular organoid function, 3D ECM-free assembled organoids were selected for a deeper long-term analysis. For this study, these organoids were cultured for 14 days and then probed for cell and structure specific markers. Upon immunofluorescent analysis at 14 days, 3D ECM-free organoids were observed to contain tubule-like-structures (TLS) and a tissue architecture remarkably similar to in vivo testes (Figure 3A-D). Interstitial cells were appropriately located in separate regions from TLS. Tissue sections were then probed for the pan germ cell marker, DDX4, spermatogonial stem cell marker, SALL4, and meiosis marker, SCP3 (Figure 3E-G). Rare DDX4-positive and SALL4-positive cells were observed, however, no SCP3 signal was identified. Upon deeper characterization of TLS, they were observed to contain a lumen-appearing space surrounded by polarized Sertoli cells and an external monolayer of peritubular cells (Figure 3I-L). Sertoli cells also exhibited tight junctions between one another, as visualized with transmission electron microscopy and with labeling against ZO-1, a junctional protein and component of the blood-testis barrier (Figure 3H,L). Next, 3D ECM-free organoids were studied for endocrine function in 12-week, long-term culture with gonadotropin stimulation (Figure 4). Testosterone and inhibin B, reproductive hormones from Leydig and Sertoli cells respectively, were both identified and quantified from organoid conditioned medium (Figure 4A). Over 12-weeks of culture, both hormones were measured to significantly respond to the supplementation of gonadotropins FSH and hCG (red arrow denotes the beginning of supplementation, during weeks 2 – 12). At completion, a test was performed to determine if endocrine responsiveness was preserved. Gonadotropins were removed for 48 h, during which both testosterone and inhibin B concentrations significantly decreased in concentration. After 48 h, gonadotropins were returned to culture and both hormone concentrations significantly increased again over a final 24 h, demonstrating proper endocrine-responsiveness (Figure 4B). Collectively, these histological and endocrine assay results demonstrate that testicular organoids are a useful model for studying testicular development (i.e., de novo compartmentalization and tubulogenesis) and somatic cell testicular function in vitro (e.g., tight junction formation and endocrine function).

Figure 1
Figure 1: Organoids self-assemble in 2D and 3D, ECM and ECM-free culture conditions.
5 dpp murine testicular cells were cultured in four different conditions: 2D ECM-free (row A), 2D ECM (row B), 3D ECM-free (row C), and 3D ECM (row D). Graphics depicting the culture method are provided in the left-hand column. Representative image montages were assembled from time-lapse images captured during the live culture. Time points for each image are labeled at the top margin. Time 0 h is before any organoid assembly has occurred, time 3 h is during ongoing cell-driven organoid assembly, and times 6 h and 9 h demonstrate representative, successfully formed organoids. Yellow arrows mark cell clusters and red arrows mark locations where separate cell clusters migrated and merged together. All scale bars = 1 mm. (E) Time required before separate cell clusters could be visibly appreciated was recorded for each condition. 2DF = 2D ECM-free; 2DE = 2D ECM; 3DF = 3D ECM-free; 3DE = 3D ECM. (F) Area per cluster was measured for each culture condition. Images were selected from n= 3 – 5 separate experiments. One-way ANOVA with Tukey’s multiple comparisons test was used to determine significance in 1E and 1F; graphs were assembled from the means of n=4 separate experiments. This figure has been modified from Edmonds and Woodruff37. © IOP Publishing. Reproduced with permission. All rights reserved. Please click here to view a larger version of this figure.

Figure 2
Figure 2: 2D ECM and 3D ECM-free cultured organoids exhibit compartmentalization of tubular and interstitial cell types.
Representative brightfield and immunofluorescent images of self-assembled organoids after 72 h of culture. 2D ECM-free samples were imaged whole mount, all other samples were imaged in 5 μm tissue sections. (A – D) 2D ECM-free culture. (E – H) 2D ECM culture. (I – L) 3D ECM-free culture. (M – P) 3D ECM culture. Cell-specific markers used for analysis are labeled on the top margin: Sertoli cell nuclei (SOX9) and cell bodies (βCatenin), germ cells (DDX4, a pan germ cell marker), Leydig cells (3βHSD), and peritubular cells (αSMA). All fluorescent samples were co-stained for DNA with DAPI. Yellow arrows point to DDX4-marked germ cells in the second column from the left, red arrows point to Sertoli cell clusters in the right two columns. Bright field scale bars = 400 μm, fluorescent scale bars = 100 μm. This figure has been modified from Edmonds and Woodruff37. © IOP Publishing. Reproduced with permission. All rights reserved. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Organoids develop tubule-like structures populated by rare germ cells.
3D-ECM-free assembled organoids were cultured for 14 days. (A–D) Representative H&E and immunofluorescent images detailing cell types and tissue features around tubule-like structures (TLS); the same organoid is depicted in adjacent tissue sections enabling side-by-side comparison of morphological features: Leydig cells (3βHSD), peritubular cells (αSMA), Sertoli cell bodies (βCatenin), collagen membrane (COL IV), and Sertoli cell nuclei (SOX9); scale bars = 100 μm. (E – G) Immunofluorescent labeling was performed against germ cells, including a pan germ cell marker (DDX4), spermatogonial stem cell marker (SALL4) and meiotically active spermatocytes (SCP3). Highly magnified insets are outlined by yellow panels in 3E and 3F. Green triangles point to DDX4-labeled cells; Red arrows point to SALL4-labeled cells. (H) Representative transmission electron micrograph (TEM) of a tight junction between Sertoli cells within an organoid; TEM scale bar = 100 nm. (I – L) High magnification representative images of a TLS labeled for key features of the seminiferous epithelium including tight junctions (ZO1) and laminin. The same TLS is depicted in adjacent tissue sections in panels I – L. All fluorescent samples were co-stained for DNA with DAPI. Images were selected from n=7 separate biological experiments. This figure has been modified from Edmonds and Woodruff37. © IOP Publishing. Reproduced with permission. All rights reserved. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Organoids secrete testosterone and inhibin B over 12-weeks of culture in response to gonadotropins FSH and hCG.
Conditioned media collected from 3D-ECM-free organoids was measured for testosterone and inhibin B via enzyme-linked immunosorbent assay (ELISA). (A) Organoids were cultured for twelve weeks, with FSH and hCG supplementation during weeks 2 – 12 (beginning of supplementation is marked with a red arrow on the x-axis); all values were compared to day 7 of culture for statistical tests. (B) Magnified graph of a “re-stimulation test” to determine if endocrine responsiveness was preserved after 12 weeks. The period of the test is outlined by the gray box in 4A. At the completion of 12-weeks in culture, gonadotropins were removed for 48 h and then re-supplemented for a final 24 h of culture. Hormones were measured at 0, 2, 6, 12, and 24 h after gonadotropin re-stimulation; red-shaded areas designate time periods during culture with FSH and hCG, non-shaded areas designate time periods without FSH and hCG; noted p-values are relative to 2 h after re-stimulation. Two-way ANOVA with Tukey’s multiple comparison’s test was used to determine significance for all endocrine data, n=5 separate biological experiments. This figure has been modified from Edmonds and Woodruff37. © IOP Publishing. Reproduced with permission. All rights reserved. Please click here to view a larger version of this figure.

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Discussion

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 methods, over the course of 24 h, an experimenter can generate as many as several hundred organoids / testicular constructs, as cell numbers allow. These methods promote tissue self-assembly into constructs with different sizes and morphologies; organoid size depends on the cell number and concentration used in culture, as seen in other organoid reports34. Reducing the organoid size or diameter might help reduce the development of inner regions of necrosis which are sometimes presents in larger organoids. A particular strength of 2D ECM and 3D ECM-free protocol methods are their ability to generate morphologically mimetic testicular organoids, containing de novo compartmentalization of tubular versus interstitial cell types. Furthermore, 3D ECM-free assembled organoids provide a model for de novo tubulogenesis of seminiferous TLS, with appropriately compartmentalized and oriented Sertoli and peritubular cells. This is an important phenotype for studying testicular organoids, and is still a variable outcome amongst different testicular organoid reports; multiple other reports lack tubule versus interstitium compartmentalization and some even develop an “inside-out tubule” phenotype32,33,34,38. While none of the organoid generation methods presented in the present manuscript were characterized to maintain large germ cell populations over extended days of culture, as germ cells were rarely observed as early as 72 h, both 2D ECM and 3D ECM-free methods might provide a useful tool to study in vitro tubulogenesis and the somatic cell component of a spermatogonial niche environment. With this goal in mind, testicular organoids provide a potential platform for optimizing future protocols to improve in vitro germline stem cell maintenance, meiotic progression, and differentiation.

Another advantage of these protocols is the ability to customize and scale organoid generation. Customized, drug-treated, or engineered cell populations can replace, or be used in addition to, primary mouse testicular cells37. If cell suspensions are of a low viability (<80 %), methods should be taken to improve cell viability of the cellular suspension. These can include reducing the time spent in dissociation media and minimizing trituration during tissue digestion (steps 2.4.1 – 2.5.2), increasing the number of washes after dissociation, or removing dead cells after dissociation with cell-sorting or dead-cell labeling kits. However, none of these steps were necessary to produce the representative data shared in this report. For 2D and 3D ECM culture methods, these protocols can be used with other structural protein-based biomaterial matrices in addition to those used for the studies presented here. These other biomaterials include collagen, gelatin, commercially available ECM extracts, and custom-made decellularized ECM-derived hydrogels25,26,28. There are a few pointers for trouble-shooting ECM gel dispensing and creating high-quality agarose 3D Petri dish inserts. When casting ECM gels onto chamber plates, be sure to work quickly and use cold pipette tips to prevent premature polymerization of the ECM within a tube or pipette tip prior to dispensing in the culture dish. When casting molten agarose into the 3D Petri dish molds (for 3D ECM-free culture), use only hot and not warm agarose to ensure high quality casting of the inserts with minimal variation, and check that the agarose has fully cooled to room temperature and solidified before attempting to remove from the mold. Agarose 3D Petri dishes are best handled gently with fine forceps. When collecting organoids for fixation, be sure to work under a dissection microscope to visualize the collection of all organoids. Organoids can stick to the plastic side of culture dishes and the inside of pipette tips; glass pipettes exhibit less organoid adherence than plastic. Fixing organoids while still encapsulated within or on top of ECM is a more challenging and delicate process than removing them from ECM prior to fixation. Tissue processing gel can be cast above the ECM-organoid construct prior to removal from the culture chamber to help reinforce the gel before fixation39. Fixation should be performed at room temperature to retrieve ECM hydrogels as they are likely to de-polymerize if lowered to 4 °C. Additionally, 0.1 % - 1.0 % glutaraldehyde can be added to the 4 % PFA solution to help further cross-link the ECM; however, this method increases background autofluorescence of the sample.

There are considerable germ cell-specific limitations to the results achieved by the organoid generation methods presented above, which represent priority areas for future innovation. Germ cells are poorly supported over extended culture, are only easily observed during the first few days of culture and are rarely observed by the end of the first week of culture and at later time points. While undifferentiated spermatogonia can be maintained within in vitro cell culture while maintaining their ability to restore spermatogenesis upon transplantation into in vivo tubules40, the tubule somatic microenvironment (i.e., direct Sertoli cell interactions) is hypothesized to be a prerequisite for differentiating pre-meiotic germ cells into and through meiosis and spermiogenesis in vitro1,5,40,41. Testicular organoids containing spermatogonia at early time points within a structurally mimetic TLS might enable the field to non-invasively study somatic-somatic, and somatic-germ cell interactions entirely in vitro. Optimization of media additives and cell preparation prior to culture (e.g., incorporation of agents used for in vitro spermatogonial stem cell culture)42,43 might increase the yield of germ cells in future studies, especially over culture periods longer than several days. Inversely, methods to re-introduce spermatogonia after TLS have formed, such as through microinjection, pose an interesting opportunity to restore germ cells within organoids, and test the capability of in vitro organized somatic environments to maintain germ cells. Other biological factors have been observed to affect ex vivo explant culture, including age/maturity45 and genetic background differences17. These same variables have yet to be directly investigated within the field of testicular organoid biology. However, it is plausible that different assembly, morphology, and functional phenotypes might result when organoids are generated from cells isolated from differently aged animals (i.e., neonatal, juvenile, adult), or animals of varying genetic background (e.g., different mouse strains).

In summary, the techniques shared in this manuscript provide four useful models for studying cell-driven testicular construct self-assembly, the generation of two separate structurally mimetic testicular organoid models, and the important achievement of TLS development and hormone-responsive endocrine function in vitro. These models encompass 2D and 3D spatial orientations in ECM and ECM-environments with minimally complex, completely defined culture medias. Each method is highly reproducible and uses only commonly available culture resources. These methods may prove advantageous for studying testicular morphogenesis in vitro and optimizing future culture conditions for in vitro spermatogenesis. More so, 2D ECM and 3D ECM-free methods provide a novel tool for studying the process of the de novo tissue compartmentalization unique to the testis, in vitro tubulogenesis, and somatic-somatic and somatic-germ cell interactions. Testicular organoids provide a flexible and scalable opportunity to investigate the development and regulation of somatic testicular physiologic hallmarks; including the blood-testis barrier and endocrine production and response; and, also, a useful tool for developing next-generation translational research tissue models. These include incorporating reproductive hormones into larger systems-level models, such as tissue-on-a-CHIP and micro-physiologic platforms45,46. Other translational goals towards which testicular organoids might one day be applied include reproductive toxicology and immunologic barrier testing, male contraceptive development, and assisted reproductive technology innovation.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

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’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’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.

Materials

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

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Cite this Article

Edmonds, M. E., Forshee, M. D., Woodruff, T. K. Extra Cellular Matrix-Based and Extra Cellular Matrix-Free Generation of Murine Testicular Organoids. J. Vis. Exp. (164), e61403, doi:10.3791/61403 (2020).More

Edmonds, M. E., Forshee, M. D., Woodruff, T. K. Extra Cellular Matrix-Based and Extra Cellular Matrix-Free Generation of Murine Testicular Organoids. J. Vis. Exp. (164), e61403, doi:10.3791/61403 (2020).

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