Establishment and Maintenance of Patient-derived Prostate Cancer Organoids: A Detailed Experimental Protocol

Summary

Prostate cancer heterogeneity and treatment resistance remain challenges. We describe a protocol for generating patient-derived organoids (PDOs) that preserve key parental tumor characteristics. This standardized model supports drug screening and precision medicine advancements.

Abstract

Prostate cancer (PCa) is a highly heterogeneous malignancy and a leading cause of cancer-related mortality in men, with significant clinical challenges in addressing treatment resistance and disease progression. Existing preclinical models, such as two-dimensional (2D) cell lines and patient-derived xenografts (PDXs), have limitations in faithfully recapitulating the complexity of PCa, including tumor heterogeneity, androgen receptor (AR)-dependent signaling, and microenvironmental interactions. To address this gap, this study presents a robust and reproducible protocol for establishing Patient-derived organoids (PDOs) that preserve the genetic, phenotypic, and histological features of the parental tumors. This method enables the culture of organoids from both localized and advanced PCa, providing a biologically relevant model for studying disease mechanisms, drug response, and biomarker discovery. The results demonstrate the protocol's ability to generate organoids from a variety of clinical specimens, with applications in high-throughput drug screening and precision medicine. By offering a standardized workflow that addresses key technical challenges, this study highlights the importance of PDOs as versatile tools for advancing prostate cancer research and developing more effective therapeutic strategies.

Introduction

Prostate cancer (PCa) remains one of the most prevalent malignancies in men worldwide, second only to lung cancer in cancer-related mortality. According to recent epidemiological studies, the global incidence of PCa has been steadily rising, and it is projected that the number of new cases of prostate cancer annually will rise from 1.4 million in 2020 to 2.9 million by 2040¹. Despite advancements in early detection and the development of novel therapies, including androgen deprivation therapy (ADT) and targeted agents, prostate cancer remains a significant clinical challenge due to its heterogeneous nature and the inevitable development of treatment resistance, particularly in advanced stages². Castration-resistant prostate cancer (CRPC), which emerges after ADT failure, represents a lethal form of the disease, characterized by poor survival outcomes and limited therapeutic options³. Understanding the molecular mechanisms driving disease progression and resistance is therefore crucial for identifying novel therapeutic targets and improving patient outcomes.

The development of preclinical models that accurately mimic the complexity of human prostate cancer is critical for advancing research in this field. Traditional 2D cell cultures, though widely used, fail to recapitulate the cellular heterogeneity, tumor architecture, and microenvironment observed in vivo⁴. As a result, the translational utility of these models is limited, particularly for studying tumor progression, drug resistance, and the dynamic interactions between cancer cells and the tumor microenvironment⁵. In contrast, patient-derived xenografts (PDXs) and organoid cultures have emerged as more sophisticated tools that bridge the gap between conventional cell lines and clinical tumors^4,5,6.

Organoid culture systems have revolutionized cancer modeling. These three-dimensional (3D) structures are derived from patient tissues and can retain the histological, genomic, and phenotypic characteristics of the original tumor^7,8,9,10. Importantly, prostate cancer organoids preserve critical features of disease heterogeneity, including AR signaling, resistance to treatment, and genomic instability, making them ideal tools for translational research¹¹. Compared to PDX models, organoids are more cost-effective, scalable, and amenable to genetic manipulation, which facilitates high-throughput drug screening, biomarker discovery, and functional studies. Organoid platforms have already shown promising results in other malignancies, including colorectal, breast, and pancreatic cancers, underscoring their potential for personalized oncology^4,12.

However, establishing and maintaining prostate cancer organoids (PCOs) presents unique technical challenges. The prostate epithelium is inherently dependent on androgen signaling and extracellular matrix (ECM) support, necessitating specialized culture conditions. Moreover, prostate cancer exhibits a wide range of clinical behaviors, from localized hormone-sensitive tumors to highly aggressive, treatment-refractory CRPC, which further complicates the development of reproducible organoid systems^11,12. Successful protocols for PCO generation must address these biological complexities by optimizing key steps, including tissue processing, enzymatic digestion, ECM embedding, and supplementation with critical growth factors, hormones, and small molecules. Such protocols are essential for ensuring the fidelity of organoids to their parental tumors and facilitating their application in downstream analyses.

In this study, we present a detailed and reproducible protocol for the establishment and long-term maintenance of PDOs. Our workflow spans the entire process, from tissue acquisition and processing of biopsy and surgical specimens to organoid passaging, cryopreservation, and resuscitation (Figure 1). This protocol has been optimized to preserve the phenotypic and genomic integrity of prostate cancer tissues, enabling robust 3D culture systems suitable for a wide range of applications, including genomic profiling, drug response testing, and functional assays. Notably, our method addresses common technical hurdles, such as low success rates in organoid culture, suboptimal growth, and difficulties in subsequent passaging, and provides standardized guidelines, thereby facilitating reproducibility across laboratories.

The development of reliable prostate cancer organoid models holds immense significance for both basic and translational research¹³. These systems offer a powerful platform for deciphering disease mechanisms, identifying novel therapeutic targets, and advancing precision medicine approaches tailored to individual patients. By addressing existing technical gaps and providing a comprehensive protocol, our work aims to accelerate the adoption of organoid technology in prostate cancer research and support the global efforts to combat this challenging disease¹⁴.

Protocol

This study was reviewed and approved by the Medical Ethics Committee of Wuxi No. 2 People's Hospital (Approval Number: Y-38 (2023)). All participants provided written informed consent forms. Patient data underwent de-identification to ensure privacy protection in compliance with ethical standards.

1. Sample collection

1. Tissue acquisition
   1. Preparation before sampling
      1. Inclusion Criteria: Ensure all participants have a confirmed diagnosis of prostate cancer, as determined by pathological or imaging examination. Derive tumor samples from surgical resection, biopsy, or other clinical specimens. Ensure all participants are 18 years or older and do not have any other malignancies, to exclude potential interference from other cancers in the study of prostate cancer. From all participants, collect the written informed consent to participate in the study and allow their samples to be used for scientific research.
      2. Exclusion Criteria: Exclude patients whose samples do not contain enough tumor tissue for culture or analysis. Exclude patients who withdraw consent or are unable to provide the required samples during the study.
   2. Prostate core needle biopsy samples:Use a special needle to obtain samples from prostate tissue. Immediately place the tissue in preservation solution (Advanced DMEM/F-12 (adDMEM/F-12) medium + rock inhibitor solution (1:2000 dilution)) and transport it to the laboratory on ice, ensuring delivery within 24 h.
      NOTE: Precisely target prostate cancer tissue and perform multisite sampling across different anatomical regions.Perform 1-2 punctures per site, each yielding ≥ 1 cm core samples.
   3. Radical prostatectomy samples: After collection, quickly place the samples in preservation solution (adDMEM/F-12 medium + rock inhibitor solution (1:2000 dilution)) and transport on ice, ensuring delivery to the laboratory within 24 h.
      NOTE: More tissue fragments can be obtained through radical prostatectomy¹⁵.
      Collect samples ≥ 1g (≈ 5 mm diameter), minimizing contamination by normal tissue. Preoperatively coordinate with clinicians to specify: 1) tumor core; 2) peri-tumoral tissue; 3) normal control. Prioritize firm fragments with gray-yellow discoloration. If absent, implement systematic random sampling. Refer to Table 1 for sample collection information.
2. Sample processing
   1. Labeling and categorization: Clearly label the collected tissue samples (including patient name, hospitalization number, tissue name, tissue site, and tissue type).
   2. Cryopreservation: Process tissue samples immediately (≤ 12h post-collection). Place the samples into pre-cooled (4 °C) preservation solution (adDMEM/F-12 medium + rock inhibitor solution (1:2000 dilution)) and store them at 4 °C. Proceed with further experimental steps within 4-6 h for optimal results.
3. Transportation and storage
   1. Cold chain transport: Transport the tissue samples to the laboratory under low-temperature conditions.
   2. Sample registration: Upon arrival at the laboratory, verify and register the sample information in detail, including the patient's basic personal information, tissue type, name, and sampling site.
4. Laboratory receipt and inspection:
   1. Sample inspection: Open the samples in a biosafety cabinet and inspect for any loss, contamination, or visible abnormalities.
   2. Sample dissection: Extract the sample and place it in a culture dish. Use tissue scissors or a scalpel to divide the sample into small pieces (approximately 1-3 mm³) for use in subsequent experiments.

2. Experimental procedures

1. Preparation of primary tissue cell suspensions
   1. Preservation: Place the collected tissue blocks into 50 mL sterile conical centrifuge tubes, ensuring that the tissues are categorized by type. Label each tube appropriately. Add preservation solution (adDMEM/F-12 medium + rock inhibitor solution (1:2000 dilution)) to each tube until the tissue is completely submerged.Store the samples at 4 °C until further processing.
      NOTE: Tissue types should include both tumor and non-tumor tissues. Sample labels should typically include the medical record number.
   2. Tissue separation: Using sterilized tweezers, transfer the prostate cancer tissue samples into a medium-sized culture dish. Wash the tissue 2-3 times with 2 mL of Dulbecco's Phosphate-Buffered Saline (D-PBS) to remove blood and debris. Carefully remove adipose tissue, muscle, and other non-epithelial components using fine-tipped tweezers, retaining only the epithelial regions. Subsequently, mince the cleaned tissue into small fragments approximately 1-3 mm³ in size with scissors.
      NOTE: Complete removal of non-epithelial components is critical for downstream applications, as contaminating stromal or adipose tissue may reduce the efficiency and specificity of cell isolation. Maintain sterile conditions throughout the procedure to minimize the risk of microbial contamination.
   3. Tissue digestion: Transfer the prepared tissue pieces into tissue processing tubes. Add the tumor tissue digestion solution (Table 2) to each tube. Activate the digestion device (Single-cell suspension preparation apparatus) using the Medium-Hard Tissue Dissociation Program for enzymatic-mechanical dissociation.
      NOTE: The digestion protocol is as follows: forward rotation at 200 rpm for 3 s, reverse rotation at 200 rpm for 3 s, forward rotation at 300 rpm for 4 s, reverse rotation at 300 rpm for 4 s, forward rotation at 600 rpm for 5 s, reverse rotation at 600 rpm for 5 s, forward rotation at 400 rpm for 3 s, reverse rotation at 400 rpm for 3 s. This sequence constitutes one cycle, which should be repeated three times. After completing the first two cycles, continue with slow rotation at 20 rpm for approximately 25 min. The duration may be adjusted based on experimental conditions. Digest the tissues at 37 °C for 40 min to 1 h.
   4. Digestion termination: Add an equal or double volume of FBS-supplemented adDMEM/F-12 medium to the cell suspension to terminate the enzymatic activity.
      NOTE: If undigested residue remains, collect the supernatant, add fresh digestive enzymes to the residue, and continue digestion at 37 °C for an additional 15-20 min. Monitor the process periodically, and combine cell suspensions from both digestion steps.
   5. Filtration: Pre-wet a 100 µm cell strainer with 1 mL of adDMEM/F-12 medium, then filter the cell suspension. Rinse the tissue processing tube and cell strainer twice with 5 mL of adDMEM/F-12 medium supplemented with rock inhibitor solution (1:2000 dilution). Collect the filtrate into a 50 mL sterile conical centrifuge tube. Pass suspension through 70 µm strainer and collect all cell suspensions into a new 50 mL tube.
      NOTE: The number of filtration steps and the pore size of the strainers may be adjusted according to the sample characteristics.
   6. Red blood cell lysis:Centrifuge the collected cell suspension at 200 × g for 5 min at 25 °C. Carefully discard the supernatant, then resuspend the cell pellet in 2 mL of red blood cell lysis buffer. Incubate the suspension on ice for 2-3 min.
      NOTE: The incubation time for red blood cell lysis may be adjusted based on the efficiency of lysis. Prolonged incubation may lead to increased cell death. It is recommended to monitor the sample under a microscope to ensure complete lysis of red blood cells without excessive damage to other cell types.
   7. Terminate the lysis:Add 6 mL of adDMEM/F-12 medium supplemented with rock inhibitor solution (1:2000 dilution) to the cell suspension. Transfer the mixture to a 15 mL sterile conical centrifuge tube and centrifuge at 200 x g for 5 min at 25 °C. Discard the supernatant. Wash the cell pellet 1-2 times with the same supplemented medium and transfer the final cell suspension to a 1.5 mL Eppendorf tube.
   8. Viability assessment: Mix 10 µL of the cell suspension with an equal volume of trypan blue solution and observe cell viability under a microscope.
      NOTE: If significant amounts of dead cells or debris are observed, perform a low-speed centrifugation at 50 x g for 5 min at 25 °C to remove them before proceeding with counting.
   9. Cell counting:Resuspend the pellet in 1 mL of adDMEM/F-12 medium, dilute the suspension as needed (typically in a 1:20 dilution), and place 6 µL of the diluted suspension on a hemocytometer to count cells under a microscope.
   10. Cell pellet collection: Centrifuge at 200 x g, 25 °C, for 5 min. Collect the final cell pellet into a 1.5 mL Eppendorf tube.

3. Organoid establishment from primary cell suspensions

1. Seeding: Determine the required volume of matrix gel based on cell count. Using pre-chilled pipette tips (stored at -20 °C), carefully transfer the appropriate volume of matrix gel to the cell pellet. Mix the cell-gel mixture on ice gently, avoiding the formation of air bubbles to ensure uniform seeding and optimal matrix gel integrity.
   NOTE: Thaw the matrix gel at 4 °C overnight prior to use. Ensure that all handling tools, including pipette tips and tubes, are pre-chilled to prevent premature gelation.
2. Plating: Dispense the cell-matrix gel mixture onto a low-adhesion 24-well culture plate, applying approximately 30 µL per drop. Ensure that each drop contains more than 10,000 cells to support effective three-dimensional (3D) structure formation.
   NOTE: Avoid excessive dilution of the matrix gel. To maintain structural integrity and support 3D culture formation, the matrix gel should constitute more than 70% of the total volume of the mixture. Organoids derived from human biopsy and prostatectomy specimens are equivalent in nature, primarily differing in viable cell yield, and identical seeding densities were applied in this protocol.
3. Incubation: Place the culture plate in a humidified incubator at 37 °C with 5% CO₂ for 5 min, allowing the matrix gel to solidify preliminary. Invert the plate and let it sit for 20 min to keep the matrix gel in 3D form. Add 800-1000 µL of culture medium (Table 3) to each well. Replace the culture medium every 2-3 days.
   NOTE: Ensure that the matrix gel is not disturbed during the inversion step to maintain the 3D structure.

4. Organoid passaging

1. Organoid collection and preparation: Using a pre-chilled pipette tip, mechanically disrupt the matrix gel to fragment the organoid clusters. Rinse each well 1-2 times with adDMEM/F-12 medium + rock inhibitor solution (1:2000 dilution). Transfer the fragmented clusters along with the culture medium into a 15 mL sterile conical centrifuge tube. Centrifuge at 200 × g for 5 min at 25 °C and discard the supernatant.
   NOTE: When organoids attain a diameter of 100 µm (typically at culture day 14). If excessive medium volume interferes with mechanical disruption, aspirate part of the medium before breaking up the clusters. For residual matrix gel that adheres to the plate surface, gently scrape using a pipette tip and rinse 2-3 times with adDMEM/F-12 medium + rock inhibitor solution (1:2000 dilution). All rinses should be pooled with the initial collection for maximum yield.
2. Organoid dissociation: Resuspend the resulting pellet in 2 mL of pre-warmed recombinant trypsin-like enzyme solution. Pipette up and down to initiate dissociation, then incubate at 37 °C for 15-30 min. Monitor dissociation under the microscope every 5 min until small cell clusters (approximately 5-6 cells per cluster) dominate the field, then proceed to the next step.
   NOTE: Timely termination of digestion is crucial to avoid over-dissociation and to preserve cell viability. Frequent microscopic observation ensures that digestion is stopped at the optimal time for cluster formation.
3. Terminate the digestion: Terminate the digestion by adding an equal or double volume of adDMEM/F-12 medium supplemented with fetal bovine serum (FBS) to quench enzymatic activity. Centrifuge at 200 x g for 5 min at 25 °C and discard the supernatant.
4. Matrix gel removal: Resuspend the pellet in 2 mL of cell recovery solution and incubate at 4 °C for 20-40 min to dissolve residual matrix gel. Centrifuge at 200 x g for 5 min at 4 °C, discard the supernatant, and repeat the wash twice with adDMEM/F-12 medium to eliminate residual matrix components. Transfer the final pellet toa 1.5 mL sterile Eppendorf tube for further processing.
   NOTE: Low-temperature incubation is critical for effective matrix gel depolymerization while preserving cell integrity. Minimize time outside the cold environment to prevent premature re-gelation.
5. Seeding and culture: Proceed with reseeding the dissociated organoid clusters following the procedures described in steps 3.1-3.3. Ensure proper cell density and matrix gel ratio for optimal 3D structure reformation.
   NOTE: Detailed protocols for organoid cryopreservation, thawing, re-embedding, and histological processing are available upon request.

5. Organoid cryopreservation

1. Organoid collection: Remove the culture plate from the incubator and mechanically dissociate organoid hydrogel aggregates using a pre-chilled pipette tip. Transfer the dissociated aggregates along with the culture medium into a 15 mL conical centrifuge tube. Rinse the wells 2-3 times with adDMEM/F-12 medium, combining the washes with the primary suspension. Centrifuge at 200 × g for 5 min at 25 °C, and discard the supernatant.
   NOTE: To ensure maximal yield, gently disrupt the organoid aggregates without causing excessive damage. All washes should be combined with the primary collection to reduce the loss of organoids during the collection process.
2. Enzymatic dissociation: Add 2-4 mL of pre-warmed recombinant trypsin-like enzyme solution to the organoid pellet in a conical tube and gently mix the suspension. Incubate the suspension at 37 °C for 15-20 min. Monitor the dissociation process microscopically every 5 min.
   NOTE: If necessary, gently pipette to assist in breaking up larger clusters into smaller aggregates (approximately 5-6 cells per cluster). Shorter digestion times are required for cryopreservation to prevent over-dissociation and loss of cell viability. Frequent observation ensures that digestion is terminated at the appropriate stage to preserve organoid integrity.
3. Digestion termination: Add an equal or greater volume of FBS-supplemented adDMEM/F-12 medium to stop the enzymatic activity. Centrifuge at 200 × g for 5 min at 4 °C and discard the supernatant.
   NOTE:If the pellet is free of residual matrix gel, proceed to step 5.5 for cryopreservation. If matrix gel contamination persists, proceed with step 5.4.
4. Matrix gel removal (optional): Follow the procedure outlined in step 4.4 to remove the matrix gel. Resuspend the pellet in2 mL of cell recovery solution and incubate at 4 °C for 20-40 min. Centrifuge at 200 × g for 5 min at 4°C, discard the supernatant, and repeat the washing process 1-2 times to ensure complete matrix gel removal.
   NOTE: The removal of residual matrix gel is critical to ensure that only dissociated cells are cryopreserved. Incomplete gel removal may affect the viability and structural integrity of the organoids upon thawing.
5. Cryopreservation: Resuspend the cell pellet in 1.5 mL of organoid freezing medium by gentle pipetting to ensure even suspension. Transfer the suspension into a cryovial and store it at -80 °C for short-term preservation. After 24 h, transfer the vials to liquid nitrogen for long-term storage.
   NOTE: Ensure that the cryopreservation medium is thoroughly mixed with the cells to prevent uneven freezing, which can lead to cell damage. Vials should be transferred to liquid nitrogen promptly after the short-term storage period to ensure long-term preservation and prevent cell degradation.

6. Organoid recovery

1. Thawing of cryopreserved organoids: Remove cryovials containing organoids from liquid nitrogen or -80 °C storage. Immediately immerse the vial in a 37 °C water bath and gently agitate until the contents are fully thawed. As soon as thawing is complete, transfer the organoid suspension to a 15 mL conical centrifuge tube using a pre-cooled (-20 °C) pipette tip.
   NOTE: Rapid and uniform thawing is critical to minimize cryo-induced cell damage. Use pre-cooled pipette tips to prevent premature warming and matrix gel reformation during transfer.
2. Cryoprotectant removal: Carefully add 1-2 mL of adDMEM/F-12 medium along the wall of the conical tube to dilute the cryoprotectant gradually. Gently mix by inversion. Centrifuge at 200 × g for 5 min at 4 °C and discard the supernatant.
   NOTE: Avoid vigorous pipetting or vortexing at this stage, as sudden osmotic shifts and mechanical stress can compromise organoid viability. A slow dilution approach minimizes cellular shock.
3. Washing and Resuspend: Add 1 mL of fresh adDMEM/F-12 medium to the pellet. Resuspend the cells gently using a pre-cooled (-20 °C) pipette tip to prevent premature matrix gel adhesion. Transfer the suspension to a 1.5 mL microcentrifuge tube.
   NOTE: Maintaining a low temperature during washing is essential to prevent thermal-induced reassembly of matrix gel components, which could hinder subsequent organoid reformation.
4. Collection: Centrifuge at 200 × g for 5 min at 4 °C and discard the supernatant. Repeat the wash step 2-3 times to ensure complete removal of residual cryoprotectant and contaminants.
5. Seeding and culture: Resuspend the final cell pellet in organoid culture medium prepared according to experimental needs. Proceed with seeding following the protocol described in steps 3.1-3.3, ensuring proper matrix gel proportion and cell density to support 3D structure re-establishment.
   NOTE: Post-thaw cell viability and recovery are highly dependent on gentle handling, proper dilution of cryoprotectants, and immediate re-establishment of a supportive 3D matrix gel environment.

7. Pre-analysis processing for organoid characterization

1. Organoid harvesting: Mechanically dissociate aggregates when organoid diameters reach 70-100 µm and transfer them to a 15 mL conical tube containing medium for subsequent culture. Rinse the wells 2-3 times with adDMEM/F-12 medium and combine the washes with the organoid suspension. Centrifuge at 200 × g for 5 min at 25 °C, and discard the supernatant.
   NOTE: For small-scale collections (e.g., 1-2 wells), use a 1.5 mL microcentrifuge tubeto ensure efficient handling and reduce sample loss. Be cautious to avoid disrupting the organoid structures during dissociation.
2. Resuspension: Resuspend the pellet in 1 mL of adDMEM/F-12 medium and transfer to a 1.5 mL microcentrifuge tube. Centrifuge at 200 × g for 5 min at 25°C and discard the supernatant.
   NOTE: Ensure the pellet is thoroughly resuspended before centrifugation to prevent clumping and to facilitate uniform processing in subsequent steps.
3. Washing and fixation: Add 1 mL of D-PBS to the pellet and mix gently by pipetting or inversion. Centrifuge at 200 × g for 5 min at 25 °C, discard the supernatant, and repeat the wash step 1-2 times. Fix the pellet with 1-2 mL of 4% paraformaldehyde (PFA) and incubate overnight at 4 °C.
   NOTE: Carefully perform the washing steps to ensure thorough removal of residual medium and other contaminants. The fixation in paraformaldehyde is crucial to preserve organoid morphology for subsequent analysis.
4. Post-Fixation Washing: Centrifuge the samples at 200 × g for 5 min at 4 °C and discard the supernatant. Wash the fixed samples 2-3 times with D-PBS, mixing gently by finger-flicking or gentle inversion to ensure proper distribution of the wash buffer and prevent loss of organoids.
   NOTE: Proper post-fixation washing is essential to remove excess paraformaldehyde and reduce background staining in downstream applications, such as immunohistochemistry.
5. Ethanol Re-Fixation: Resuspend the pellet in 1 mL of 70% ethanol mix gently by finger-flicking, and incubate at room temperature for 20 min. Centrifuge at 200 × g for 5 min at 4 °C and discard supernatant.
   NOTE: Ethanol fixation at this stage helps to preserve organoid integrity and prepare the samples for embedding or further processing. Ensure uniform suspension in ethanol to avoid uneven fixation.
6. Matrix gel clearance (Optional): If residual matrix gel remains in the pellet, follow the protocol described in step 4.4 to remove it. This step involves resuspending the pellet in cell recovery solution and incubating at 4 °C for 20-40 min, followed by centrifugation and multiple washes to ensure complete gel removal.
   NOTE: Removing matrix gel is particularly important for downstream histological analysis or imaging, as residual gel may interfere with tissue sectioning and analysis.
7. Agarose embedding: Prepare about 6 mL of 2% agarose by melting at 96 °C during organoid ethanol fixation. Mix 100 µL of the melted agarose with the organoid pellet, ensuring minimal air bubble formation. Transfer the mixture to a mold, solidify at 4°C, then process for paraffin embedding.
   NOTE: Pre-warm pipette tips (with trimmed ends) to prevent clogging. Agarose embedding facilitates the preservation of organoid structure while allowing for easy handling and sectioning. Avoid bubble formation during mixing to ensure even embedding and optimal tissue sectioning.

Results

Morphological and dynamic growth features of patient-derived prostate cancer organoids
Patient-derived prostate cancer organoids were successfully established from fresh tumor tissues collected during clinical procedures. Initial organoid formation was typically observed within 5-7 days of culture (Figure 2). Under brightfield microscopy, organoids exhibited distinct morphological features, forming compact, well-defined spheroids or glandular-like structures. Organoids d...

Discussion

The establishment of PDOs for PCa represents a critical advancement in preclinical modeling, offering significant advantages over conventional 2D cell cultures and PDXs¹⁶. The method presented in this study enables the generation of organoids that faithfully recapitulate the histological, genetic, and phenotypic characteristics of the original tumor. By preserving tumor heterogeneity, including AR signaling and cellular architecture, the organoids provide a more biologically relevant platform for ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
A 83-01	TOCRIS	2939
adMEM/F12	Gibco	12634010
biosafety cabinet	Jinan Xinbeixi Biotechnology Co., Ltd
Cell Recovery Solution	Corning	354253
centrifuge	Shenzhen Ruiwode Life Technology Co., Ltd	M1416R
centrifuge tubes	Nanjing iResearch Biotechnology Co., Ltd
Cultrex PathClear BM	R&D	3533-010-02
Cultrex Reduced Growth Factor BME, Type 2	R&D	3533-005-02
culture containers (24-well low-adhesion plates)	thermo	Nunc-174930-ZX
DHT	ISOREAG	521-18-6
electronic balance	Nanjing iResearch Biotechnology Co., Ltd
Fetal Bovine Serum	Gibco	10099141
GlutaminePlus-200mM	R&D	B90210
HEPES	R&D	1254/10
humidified cell culture incubator (37°C, 5% CO2)	Shanghai Hetian Scientific Instrument Co., Ltd	HY-160SY
ice box	Nanjing iResearch Biotechnology Co., Ltd
medium-sized culture dishes	absin	abs7005
N21-MAX Media Supplement (50X)	R&D	AR008
N-Acetylcysteine	Sigma	A9165-5G
Nicotinamide	R&D	3533-010-02
Normocin	Invivogen	ant-nr-2
Pipet-aid pipetting device	Nanjing Hotspot Scientific Instrument Co., Ltd
pipettes	Nanjing Hotspot Scientific Instrument Co., Ltd
Primocin	Invivogen	ant-pm-2
rhEGF	R&D	236-EG-01M
rhFGF	R&D	233-FB-025
rhFGF-10	R&D	345-FG-025
rhNoggin	R&D	6057-NG-100
rhR-Spondin 1	R&D	4656-RS-100
SB 202190	TOCRIS	1264
Single-cell suspension preparation apparatus	RWD	DSC-400
Tissue scissors/scalpel	Nanjing iResearch Biotechnology Co., Ltd
TryplE Express	Gibco	126A04013
Tumor Dissociation Kit, Human	Miltenyi Biotec	130-095-929
tweezers	Nanjing iResearch Biotechnology Co., Ltd
water bath	Changzhou Zhongjie Experimental Instrument Manufacturing Co., Ltd	HH-600
Y-27632 dihydrochloride	R&D	1254/10