A Defined Hydrogel-Based Method For Generating Three-Dimensional Human Breast Organoids That Recapitulate Mammary Morphogenesis

Summary

This protocol describes a defined hydrogel-based method to generate human breast organoids that recapitulate key features of mammary morphogenesis in a controlled three-dimensional culture system.

Abstract

The development of physiologically relevant human model systems that recapitulate tissue architecture and cell-state dynamics remains a major challenge in studying breast development and early events in carcinogenesis. Conventional two-dimensional cultures and many three-dimensional systems fail to capture the structural organization and microenvironmental cues that define the human mammary gland. Here, we describe a reproducible method for generating three-dimensional human breast organoids from primary epithelial cells embedded within a defined hydrogel matrix composed of type I collagen, laminin, fibronectin, and hyaluronic acid. This system supports the progression of single cells through key stages of mammary morphogenesis, including progenitor expansion, epithelial patterning, and the formation of terminal ductal lobular unit-like structures, as well as the emergence of a mesenchyme-like compartment, over a 21-day culture period. We provide a step-by-step protocol for hydrogel preparation, cell seeding and culture conditions. The method is compatible with high-content imaging and quantitative analysis of organoid number, size distribution, and architectural complexity. This platform enables mechanistic studies of epithelial plasticity and environmental perturbations, providing a scalable and biologically relevant system for investigating early tissue-level changes associated with breast cancer risk.

Introduction

Understanding human mammary gland development and the early events that predispose tissue to malignant transformation requires experimental systems that faithfully recapitulate tissue architecture, cellular hierarchy, and microenvironmental signaling. While two-dimensional epithelial cultures have provided important mechanistic insights, they lack the structural context necessary to model epithelial organization and morphogenesis¹. Existing three-dimensional culture systems, including those based on basement membrane extracts, have advanced the field but remain limited by variable composition, incomplete control over extracellular matrix components, and inconsistent support of higher-order tissue architecture¹. In addition, many organoid systems based on basement membrane extracts are constrained by their inability to consistently support the emergence of tissue-like organization¹. In particular, many systems rely on exogenous stromal components rather than enabling the endogenous development of supportive microenvironments, thereby limiting their ability to model epithelial–mesenchymal interactions that are central to tissue development and disease. These limitations restrict the reproducible study of developmental processes, epithelial plasticity, and the effects of environmental or molecular perturbations on tissue organization.

To address these limitations, we developed a defined hydrogel-based three-dimensional human breast organoid model that enables primary epithelial cells to generate organized structures within a defined extracellular matrix^2,3,4,5,8. The hydrogel consists of type I collagen, laminin, fibronectin, and hyaluronic acid, components selected based on their established roles in mammary gland development and epithelial morphogenesis. These extracellular matrix molecules engage distinct cellular receptors, including integrins, discoidin domain receptors, CD44, and RHAMM, and are known to regulate epithelial polarity, branching morphogenesis, stem cell maintenance, mechanotransduction, and tissue organization in the mammary gland^6,7. Previous studies describing this hydrogel system demonstrated that incorporation of these extracellular matrix components markedly improved ductal-lobular morphogenesis and epithelial maturation compared to collagen-only or Matrigel-based conditions^3,8. In addition, the physical properties of this hydrogel formulation were previously characterized by atomic force microscopy, demonstrating that the composite extracellular matrix hydrogel exhibited lower stiffness and increased swelling relative to collagen-only gels (Young’s modulus: 256.7 ± 20.0 Pa versus 559.2 ± 204.0 Pa, respectively), consistent with a softer and more hydrated matrix environment⁸.

Importantly, this model supports the emergence of a mesenchyme-like compartment that arises alongside epithelial structures, providing endogenous structural and signaling support that more closely reflects native tissue organization³. The physiological relevance of this hydrogel system has been evaluated through direct comparisons with conventional Matrigel-based organoid cultures using both morphological and transcriptomic analyses. Earlier studies demonstrated that hydrogel cultures supported more organized ductal-lobular tissue formation and multilineage differentiation than Matrigel-based cultures while also preserving hormone responsiveness⁸. More recently, integrated single-cell RNA sequencing analyses comparing hydrogel-derived organoids, Matrigel-grown organoids, and primary human breast tissue demonstrated that hydrogel-grown organoids more faithfully recapitulate the epithelial hierarchy, cellular diversity, and epithelial–mesenchymal interactions present in native human breast tissue³. In contrast, Matrigel-grown organoids were enriched for proliferative hybrid basal states and lacked stromal-like populations, consistent with epithelial self-assembly rather than directed organogenesis.

The protocol described here provides a reproducible and scalable method for generating organoids from primary human tissue, with optional steps for fibroblast depletion and dissociation to single cells. Because this platform is compatible with live imaging, high-content imaging, quantitative morphometric analysis, cell tracking, and genetic perturbation studies, it can be integrated with downstream approaches assessing organoid number, size, architecture, and growth dynamics. This platform therefore provides a biologically relevant system for studying human breast development, epithelial plasticity, epithelial–microenvironment interactions, and tissue-level responses to developmental, molecular, or environmental perturbations.

A schematic overview of the workflow, including tissue processing, hydrogel preparation, organoid culture, developmental progression, and downstream analyses, is provided in Figure 1, while representative organoid morphologies generated using this platform are shown in Figure 2.

Figure 1. Overview of the hydrogel-based organoid generation workflow. (A) Flowchart summarizing the protocol workflow, including tissue collection, tissue processing, cryopreservation, recovery and optional cell preparation, hydrogel preparation, organoid culture, organoid development, and downstream analytical applications. (B) Visual schematic depicting the major stages of the hydrogel-based organoid generation protocol, including tissue processing and preparation, recovery and optional cell preparation, and hydrogel preparation and organoid seeding. Please click here to view a larger version of this figure.

Figure 2. Representative organoid morphologies generated in the defined hydrogel system. Representative brightfield images of organoids derived from different human tissues cultured in the defined hydrogel matrix. Breast organoids generated from reduction mammoplasty-derived single epithelial cells were imaged at day 21 of culture. Patient-derived xenograft organoids generated from tumor fragments were imaged at day 16 of culture. Salivary gland organoids generated from epithelial fragments were imaged at day 3 of culture. Kidney organoids generated from epithelial fragments were imaged at day 17 of culture. Scale bars = 50 µm. Please click here to view a larger version of this figure.

Protocol

Primary tissues that would otherwise have been discarded as medical waste following surgery were obtained in compliance with all relevant laws using protocols approved by the institutional review boards at Maine Medical Center and Tufts Medical Center. All tissues were anonymized prior to transfer and could not be traced to specific patients. For this reason, this research was granted exemption status by the Committee on the Use of Humans as Experimental Subjects at the Massachusetts Institute of Technology and at Tufts University Health Sciences (IRB #13521). All patients enrolled in this study signed an informed consent form agreeing to participate in the study and to the publication of the results.

1. Tissue Processing and Preparation

NOTE: Perform all procedures involving human tissue in accordance with institutional biosafety and ethical guidelines. Refer to the workflow schematics shown in Figure 1A and 1B for a visual overview of the protocol steps and organoid preparation workflow prior to beginning the procedure.

1. Prepare MEGM using mammary epithelial basal medium supplemented with 52 µg/mL bovine pituitary extract, 10 ng/mL human epidermal growth factor, 5 µg/mL insulin, 500 ng/mL hydrocortisone, 1% (v/v) GlutaMAX, and 1% (v/v) Antibiotic-Antimycotic (100X).
2. Prepare the dissociation medium by adding 1.5 mg/mL collagenase A (stored at −20°C) and 100 U/mL hyaluronidase (stored at 4°C) to mammary epithelial growth medium (MEGM).
3. Receive human breast tissue obtained from prophylactic mastectomies and reduction mammoplasties within 24 h post-excision, unfixed in phosphate-buffered saline (PBS), and maintained at 4°C. Weigh the tissue on a precision balance to determine the total tissue weight.
4. Place the tissue in a sterile biosafety cabinet. Mechanically mince the tissue into 3–5 mm³ fragments using sterile scalpels. Transfer approximately 2–3 g of minced tissue into a 15 mL conical tube. Add 10 mL of dissociation medium to each tube. 
5. Incubate the tubes at 37°C on an orbital rotator at 8 rpm for 12–18 h to enzymatically digest the tissue. Consider digestion complete when no large tissue fragments remain and a homogeneous suspension of evenly fragmented material is visible throughout the medium.
6. Allow epithelial fragments to settle by gravity for 5 min. Confirm that no visible fragments remain suspended in the medium and that a distinct pellet is present. Carefully decant and discard the supernatant without disturbing the pellet.
   NOTE: The supernatant contains stromal cells and matrix components, and may be washed and used or preserved for other studies. 
7. Resuspend the pellet in 10 mL wash medium consisting of 5% fetal bovine serum (FBS) in PBS. Centrifuge at 250 × g for 5 min in a swing-bucket centrifuge at room temperature.
8. Repeat the wash step three additional times or until the supernatant is clear and free of visible debris.
9. Resuspend the final pellet in freezing medium consisting of 10% dimethyl sulfoxide (DMSO) in MEGM at a volume of 1 mL per gram of initial tissue weight.
   CAUTION: DMSO is harmful upon contact or inhalation. Handle using appropriate personal protective equipment and work in a chemical safety hood if required by institutional guidelines.
10. Aliquot 1 mL of the suspension into each cryovial. Place the cryovials at −80°C in a freezing container prior to long-term storage in liquid nitrogen.
    NOTE: This step represents a pause point. Store samples at −80°C before transferring to long-term storage conditions.

2. Recovery and Optional Cell Preparation

1. Thawing and Initial Recovery
   1. Remove cryopreserved tissue from −80°C storage after freezing for at least 24 h, or from the liquid nitrogen long term storage. Immediately submerge the cryovial up to the cap in a 37°C water bath and thaw rapidly for 1 min to minimize cell death.
      NOTE: Gently agitate the cryovial during thawing to ensure uniform warming.
   2. Once fully thawed, immediately transfer the contents of the cryovial to a 15 mL conical tube containing 10 mL of MEGM. Centrifuge at 250 × g for 5 min.
2. Optional Fibroblast Depletion
   NOTE: Fibroblast depletion by pre-plating is an optional enrichment step used to reduce rapidly adherent stromal or fibroblast populations when a more epithelial-enriched starting population is desired. This step is not required for organoid formation and may be adjusted depending on the experimental objective.
   1. Resuspend the pellet in 10 mL of MEGM.
   2. Transfer the resuspended tissue to a 10 cm cell culture dish. Incubate at 37°C in a humidified incubator with 5% CO₂ for 90 min to allow fibroblast attachment.
      NOTE: Do not disturb the culture dish during incubation to ensure efficient fibroblast adherence.
   3. Collect the supernatant containing non-adherent cells using a 10 mL serological pipette while gently tilting the culture dish to approximately 45°. Gently rinse the plate with 5 mL PBS and combine the rinse with the collected supernatant.
   4. Centrifuge the combined suspension at 250 × g for 5 min to recover epithelial-enriched material.
      NOTE:  Cells that adhered to the plate are enriched for mammary gland fibroblasts, and may be propagated separately by adding medium composed of DMEM + 10% FBS and antibiotics. 
3. Optional Dissociation to Single Cells
   1. Resuspend the pellet in 500 µL of prewarmed (37°C) dissociation enzyme solution consisting of 0.25% trypsin. Alternative dissociation reagents may require optimization. Transfer the suspension to a 1.5 mL microcentrifuge tube. Triturate the sample 20 times using a P1000 pipette to promote dissociation.
      CAUTION: Enzymatic dissociation reagents may be harmful. Handle using appropriate personal protective equipment and avoid skin or eye contact.
   2. Incubate the tube at 37°C for 3-5 min. Mechanically dissociate the sample immediately following incubation by triturating 20 times using a P1000 pipette.
   3. Add 1 mL of serum-containing wash medium to neutralize the enzymatic reaction. Centrifuge at 500 × g for 5 min.
   4. Resuspend the pellet in 300 µL dispase solution (5 U/mL) and 30 µL DNase solution (1 mg/mL). Incubate at 37°C for 3–5 min.
   5. Mechanically dissociate the sample by pipetting 15–20 times. Add 700 µL serum-containing wash medium to the suspension.
   6. Filter the cell suspension through a 40 µm cell strainer into a clean tube. Either a standard 40 µm cell strainer or a Flowmi pipette-tip strainer may be used. Flowmi strainers may reduce sample loss when working with limited cell numbers.
   7. Determine viable cell number using trypan blue exclusion. Mix 10 µL of the cell suspension with trypan blue at a 1:1 ratio and load 10 µL of the mixture into a cell counting chamber or counting slide. Determine cell viability using an automated cell counter or hemocytometer.
   8. Centrifuge the suspension at 500 × g for 5 min at room temperature.
   9. Resuspend the cell pellet in MEGM at the desired concentration for hydrogel seeding.
      NOTE: For single-cell seeding experiments, typical inputs range from approximately 1,000–5,000 cells per 200 µL hydrogel, depending on the experimental objective and donor sample characteristics. Lower seeding densities are generally used for live imaging and morphogenesis studies to facilitate tracking of individual organoid development, whereas higher densities are commonly used for endpoint molecular analyses, including RNA and protein collection.

3. Hydrogel Preparation and Organoid Seeding

1. Stock Preparation and Volume Calculations
   1. Place laminin solution (1.18 mg/mL), hyaluronic acid solution (1 mg/mL in sterile water), fibronectin solution (2 mg/mL in sterile water), and PBS 1× on ice prior to use. Store laminin and fibronectin aliquots at −80°C and store hyaluronic acid solution at 4°C. Thaw frozen components slowly on ice at 0°C–4°C prior to use.
   2. Prepare a 25× extracellular matrix supplement stock solution. To prepare 1 mL, combine 425 µL laminin, 250 µL hyaluronic acid, 250 µL fibronectin, and 75 µL PBS in a tube maintained on ice. Mix gently by inverting the tube 3–4 times. Store the solution at 4°C for up to 1 month. 
   3. Determine the desired final hydrogel volume and composition prior to preparation. Prepare at least 20% excess volume to account for material loss during pipetting and transfer.
      NOTE: The hydrogel formulation consists of collagen I, a 1× final concentration of extracellular matrix supplements (from a 25× stock), and 12.5% (v/v) 0.1 N sodium hydroxide relative to the collagen I volume for pH neutralization and polymerization.
      NOTE: The final hydrogel concentrations are 1.7 mg/mL collagen I, 20 µg/mL laminin, 20 µg/mL fibronectin, and 10 µg/mL hyaluronic acid.
   4. Calculate the required volume of collagen I (V_collagen) using the following formula:
      
      Here, C_final = 1.7 mg/mL, V_total is the desired final hydrogel volume, and C_stock is the concentration of the collagen stock solution. Use collagen stock concentrations between 2–12 mg/mL.
      NOTE: Collagen stock concentration may vary between batches. Higher concentrations increase viscosity and should be pipetted slowly.
   5. Calculate the volume of 0.1 N sodium hydroxide (V_NaOH) using the following formula:
      V_NaOH = 0.125 x V_collagen
      Prepare a 0.1 N sodium hydroxide working solution by diluting 1 N sodium hydroxide in sterile water.
   6. Calculate the volume of extracellular matrix supplement (V_ES) using the following formula:
      
   7. Calculate the remaining volume to be filled with growth medium using the following formula:
      V_{growth medium} = V_total - (V_collagen + V_NaOH + V_ES + V_cells/tissue)
      NOTE: Determine the volume of cells or tissue fragments for each experiment individually. Use a typical range of 10–100 µL within the allowable volume. Precise fragment counting is not performed because fragment size and composition vary substantially between preparations. Standardize seeding by visual assessment of fragment abundance and distribution.
2. Preparation of Hydrogel Master Mix
   1. Place collagen I, extracellular matrix supplement, sodium hydroxide (0.1 N), and growth medium on ice at 0°C–4°C. Maintain all components at low temperature to prevent premature polymerization.
   2. Add the calculated volume of collagen I to a chilled microcentrifuge tube.
   3. Add the calculated volume of pre-chilled growth medium to the collagen solution.
   4. Add the calculated volume of 0.1 N sodium hydroxide to neutralize the solution.
      NOTE: Previous optimization established that these conditions produce the appropriate gel pH; therefore, routine pH testing of the hydrogel mixture is not required.
      NOTE: Perform all subsequent steps rapidly to prevent premature collagen polymerization.
   5. Mix the solution by shaking the tube vigorously at a rate of approximately one shake per second for at least 6 shakes.
   6. Add the calculated volume of extracellular matrix supplement to the mixture.
   7. Add the calculated volume of single cells or tissue fragments using a P1000 pipette or wide-bore pipette tip. Mix immediately by shaking as described in Step 3.2.5 and proceed immediately to the next step.
3. Hydrogel Deposition
   1. Pipette the hydrogel mixture into culture vessels at the desired volume per well. Use 200 µL hydrogel per well for 4-well chamber slides, 100 µL per well for 8-well chamber slides, and 20 µL per well for 96-well plates.
   2. Spread the hydrogel into a thin pad across the surface when using chamber slides. Position the pipette tip at the center upper edge of the chamber well and drag the tip across the surface while dispensing the gel to create an even pad. For 96-well plates, position the pipette tip at the center of the well while maintaining contact with the surface and dispense the gel centrally to maintain a dome structure. Avoid pipetting air into the gel as this will create bubbles.
   3. Incubate the culture vessels at 37°C in a humidified incubator with 5% CO₂ for 60 min to allow complete polymerization.
      NOTE: Polymerization conditions described here were optimized for the culture formats used in this study. Minor adjustments in polymerization time may be required depending on culture vessel geometry, hydrogel volume, and incubation conditions.
4. Culture and Maintenance
   1. Add prewarmed MEGM to each well. Adjust the volume according to the culture format used.
   2. Gently detach the hydrogel from the culture surface using a pipette tip to allow the gel to float. Slide the pipette tip around the perimeter of the gel and gently lift beneath the polymerized hydrogel to release it from the surface.
   3. Replace the MEGM twice weekly with fresh prewarmed medium. Maintain cultures in a humidified incubator with 5% CO₂ for approximately 3–4 weeks and terminate cultures before complete hydrogel collapse occurs. Identify collapsed hydrogels by the appearance of dense, opaque, plug-like structures resulting from extensive cellular outgrowth within the gel (see representative examples in Supplementary Figure 1).

Results

Successful execution of this protocol results in the formation of three-dimensional organoid structures that exhibit organized epithelial morphology and tissue-specific architectural features. Organoids begin to form within 3–7 days following seeding and continue to develop throughout the culture period. Previous characterization of this hydrogel organoid system demonstrated reproducible organoid formation across multiple independent primary human donors³. In that study, primary epithelial cells isolated from 12 disease-free reduction mammoplasty donors were evaluated, and 11 of 12 donor samples successfully generated organoids under the described culture conditions. Across donors, the median number of organoid structures formed per 100 seeded cells was 1.775 (95% CI: 0.45–4.10). Although substantial inter-donor variability in organoid formation efficiency and growth dynamics was observed, complex ductal-lobular and acinar morphologies were reproducibly generated across donors.

Breast organoids derived from single epithelial cells display progressive morphogenesis, forming branching structures by day 21 of culture (Figure 2). These structures are characterized by elongated projections and multicellular organization. Organoids exhibited projected areas ranging from 10,000–90,000 µm² by day 18, with circularity values below 0.3, calculated as 4π × (Area/Perimeter²)^3,9. Organoids derived from tissue fragments typically formed structures exceeding 90,000 µm² by day 18, whereas organoids derived from tumor fragments formed denser and more irregular structures with reduced organization and increased radial projections, consistent with altered growth behavior.

Organoids derived from salivary gland and kidney epithelial fragments also exhibit distinct morphologies under the same hydrogel conditions (Figure 2). Salivary gland organoids form compact structures at early time points (day 3), whereas kidney organoids develop elongated and asymmetric morphologies by day 17. These observations are included to demonstrate the broader adaptability of the hydrogel-based organoid system beyond mammary tissue and illustrate its ability to support organoid formation from multiple epithelial tissue sources.

Immunostaining and molecular analyses previously performed^3,5,8 demonstrated the presence of epithelial lineage markers, including the luminal markers KRT8, KRT18, KRT19, E-cadherin, GATA3, JAG1, Notch1, and MUC1, as well as the basal or myoepithelial markers KRT5, KRT14, Slug, SOX9, and TP63, indicating preservation of epithelial heterogeneity within the organoids. Structural features consistent with terminal ductal lobular unit-like organization were observed by confocal microscopy and three-dimensional reconstruction and were defined as structures containing elongated ductal regions connected to terminal lobule- or alveolar-like buds resembling the organization of native human terminal ductal lobular units. These structures also exhibited layered epithelial organization with luminal and basal cell patterning consistent with previously published analyses of this platform³.

A mesenchyme-like compartment was observed in association with and between epithelial structures and was characterized by migratory behavior and expression of markers including VIM, SNAI1, ZEB1, S100A, CD90, and FAPα. Together with time-lapse microscopy analyses, these observations support the emergence of a supportive microenvironment within the culture system³.

Importantly, this protocol is intended to provide a broadly adaptable methodological framework rather than establish a single fixed biological benchmark. Quantitative outcomes, including organoid formation efficiency, size, branching complexity, and cellular composition, may vary depending on donor source, menopausal status, starting material (e.g., tissue fragments versus single cells), and experimental conditions.

Suboptimal outcomes include reduced organoid formation efficiency (<1 organoid per 500 seeded cells), excessive cellular debris, failure of collagen polymerization or failure to establish organized structures. These outcomes are commonly associated with low cell viability, incomplete tissue dissociation, incorrect hydrogel composition, or improper pH neutralization.

Quantitative analysis of organoid development can be performed using live imaging and high-content imaging approaches to measure organoid number, size distribution, branching behavior, structural complexity, and cell dynamics. Previous studies using this platform performed longitudinal live imaging, cell tracking, morphometric analysis, and genetic perturbation studies to quantify organoid growth dynamics and lineage behavior over time^3,5. Imaging was performed using confocal microscopy, and image analysis was conducted using NIS-Elements software.

Supplementary Figure 1. Representative example of a collapsed hydrogel during long-term organoid culture at day 18. Collapsed hydrogels appear as dense, opaque, plug-like structures resulting from extensive cellular outgrowth and matrix contraction. These morphological features were used as criteria for terminating cultures prior to complete hydrogel collapse. Scale bar = 500 µm.Please click here to download this file.

Discussion

The protocol described here enables the reproducible generation of three-dimensional human breast organoids within a defined hydrogel microenvironment that supports key features of mammary morphogenesis³. Several steps are critical for the success of this method. First, tissue processing and enzymatic dissociation must be carefully controlled to preserve epithelial viability while minimizing overdigestion, which can reduce cell yield and impair subsequent morphogenesis¹⁰. In particular, brief and sequential enzymatic treatments, combined with gentle mechanical dissociation are essential for maintaining functional epithelial populations. Second, hydrogel preparation requires precise control of collagen concentration, pH, and timing¹¹. Neutralization of collagen initiates polymerization; therefore, all steps following sodium hydroxide addition must be performed rapidly and on ice to ensure consistent gel formation. Incomplete or delayed polymerization can result in poorly structured or collapsed hydrogels that do not support organoid development. Finally, seeding density must be empirically optimized for each donor sample, as excessive tissue or cell loading can lead to rapid gel contraction and loss of structural integrity¹².

Several troubleshooting considerations can improve reproducibility. Poor organoid formation may result from low cell viability, suboptimal hydrogel composition, or improper gel handling during deposition¹³. Ensuring that collagen stocks are maintained at 4°C and have not undergone premature polymerization is essential for consistent gel quality¹¹. Additionally, successful detachment of hydrogels from the culture surface following polymerization serves as an indicator of proper gel formation; failure to detach typically reflects incomplete polymerization or inappropriate surface conditions¹⁴. Variability in organoid size and morphology is expected across donor samples and reflects biological heterogeneity rather than technical failure. Previous studies using this platform demonstrated reproducible organoid formation across a broad set of independent primary human donors despite substantial inter-donor variability in organoid formation efficiency and morphogenesis³.

This method has several limitations. While the model supports the emergence of a mesenchyme-like compartment alongside epithelial structures, it does not fully recapitulate the complexity of the in vivo stromal, immune, and vascular microenvironment. The composition of the hydrogel, although defined, represents a simplified extracellular matrix and may not capture all biomechanical or biochemical cues present in native tissue^15,16. Additionally, donor-to-donor variability can influence organoid growth dynamics and morphology, necessitating empirical optimization for specific applications. Despite these limitations, the ability of this system to support self-organization and endogenous epithelial–mesenchymal interactions represents a significant advance over many existing culture models³.

Compared to commonly used basement membrane extract-based systems, this approach provides greater control over extracellular matrix composition and reduces variability associated with undefined materials¹⁷. Previous studies directly comparing this hydrogel platform with Matrigel-based organoid systems demonstrated substantial differences in tissue organization and cellular composition^3,8. Hydrogel-grown organoids more closely resembled native human breast tissue at both the morphological and transcriptomic levels, preserving multilineage epithelial populations, including luminal, basal, progenitor, and mesenchymal-like compartments, whereas Matrigel-grown organoids were dominated by proliferative hybrid basal-like states and lacked stromal populations. In addition, unlike co-culture systems that rely on the addition of exogenous stromal cells, this model enables the intrinsic emergence of a supportive mesenchyme-like compartment, allowing the study of epithelial–microenvironment interactions in a more physiologically relevant and less artificially engineered context. Emerging mesenchymal-like populations within the hydrogels were previously validated through complementary live imaging, immunostaining, and single-cell transcriptomic analyses³. These studies demonstrated the emergence of highly motile stromal-like cells expressing mesenchymal and epithelial–mesenchymal transition-associated markers including Vimentin, THY1/CD90, FAP, S100A4, ZEB1, and Snail, together with reciprocal epithelial–mesenchymal signaling interactions identified through ligand–receptor analyses. These features make the system particularly well suited for investigating processes that depend on tissue architecture and cell–cell communication, including morphogenesis, epithelial plasticity, and early tissue remodeling^4,5.

The described platform has broad applications in basic and translational research. It can be used to study human breast development, model early events in disease initiation, and evaluate the effects of molecular or environmental perturbations on tissue organization. Previous studies using this platform demonstrated that functional perturbation of developmental regulators such as DDR1 and RUNX1 alters lineage differentiation, epithelial organization, and ductal-lobular morphogenesis in three-dimensional culture^5,18. Compatibility with quantitative imaging and high-content analysis further enables systematic interrogation of phenotypic outcomes, including changes in organoid size, structure, and complexity. As such, this method provides a scalable and biologically relevant platform for studying human tissue organization and its disruption in disease-relevant contexts.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
15 mL conical tubes	VWR	89039-664	Sterile tubes for tissue processing and centrifugation
40 μm cell strainer	VWR	732-2757	Filtration device for single-cell suspension preparation
40 μm cell strainer for low volumes	Bel-Art	136800040	Filtration device for single-cell suspension preparation
Automated cell counter	Bio-Rad	1450102	Device used to count cells and determine viability
Bovine pituitary extract	Thermo Scientific	13028014	Supplement for epithelial cell media
Cell counting slides	Bio-Rad	145-0011	Dual-chamber slides used for cell counting
Centrifuge	N/A	N/A	Benchtop centrifuge/s need to have at least 500 x g speed capability and accommodation of 15 mL and 1.5 mL tubes.
Collagen I	Millipore Sigma	08-115	Extracellular matrix protein used for hydrogel formation
Collagenase A	Sigma-Aldrich	11088793001	Enzyme used for tissue dissociation
Cryovials	Corning	976171	Sterile vials for cryogenic storage of samples
Culture vessels (e.g., chamber slides, multiwell plates)	Corning	354104 354108 3603	Platforms for hydrogel deposition and organoid culture.
Dimethyl sulfoxide	Millipore Sigma	317275	Cryoprotectant used in freezing medium
Dispase II	Roche	4942078001	Enzyme used for secondary tissue dissociation
DNase I	Roche	10104159001	Enzyme used during cell dissociation.
Fetal bovine serum	Gibco	10437	Serum supplement used in wash and neutralization media
Fibronectin	Sigma-Aldrich	F2006	Extracellular matrix protein component
GlutaMAX	Thermo Scientific	35050061	Supplement for epithelial cell media
Human epidermal growth factor	Sigma-Aldrich	E9644	Supplement for epithelial cell media
Hydrocortisone	Sigma-Aldrich	H0888	Supplement for epithelial cell media
Hyaluronic acid	Millipore Sigma	385908	Extracellular matrix component for hydrogel formulation
Hyaluronidase	Sigma-Aldrich	H3506	Enzyme used for tissue dissociation
Incubator	Thermo Scientific	3598	Device used for tissue culture incubation
Insulin	Sigma-Aldrich	I9278	Supplement for epithelial cell media
Laminin	Gibco	23017-015	Extracellular matrix protein component
Mammary epithelial basal medium	Thermo Scientific	M171500	Growth medium for epithelial cells
Microcentrifuge tubes (1.5 mL)	Thermo Scientific	3451	Tubes used for small-volume reactions
Orbital rotator	Thermo Scientific	400110	Rotator used for tissue dispersion during enzymatic dissociation
P1000 pipette	Gilson	P1000	Device used to mix and transfer volumes up to 1,000 μL
Penicillin-streptomycin	Thermo Scientific	15140122	Antibiotic supplement for epithelial cell media
Phosphate-buffered saline	Gibco	20012-027	Buffer solution used for washing and rinsing
Precision balance	Mettler Toledo	ML303E	Balance used for tissue weighing
Serological pipette	Nunc	170356N	Pipette used for harvesting non-adherent epithelial cells
Sodium hydroxide (1 N)	Fisher Chemical	SS261	Reagent used for collagen neutralization
Sterile scalpels	Bard-Parker	372615	Tools used for mechanical tissue mincing
Trypan blue solution	Gibco	15250061	Dye used for cell viability assessment
Trypsin (0.25%)	Gibco	25200056	Enzymatic reagent used for cell dissociation
Water bath (37 °C)	VWR	10LA	Device used for controlled thawing of samples