Culture of Bladder Cancer Organoids as Precision Medicine Tools

Summary

Patient-derived organoids (PDOs) are a powerful tool in translational cancer research, reflecting both the genetic and phenotypic heterogeneity of the disease and response to personalized anti-cancer therapies. Here, a consolidated protocol to generate human primary bladder cancer PDOs in preparation for the evaluation of phenotypic analyses and drug responses is detailed.

Abstract

Current in vitro therapeutic testing platforms lack relevance to tumor pathophysiology, typically employing cancer cell lines established as two-dimensional (2D) cultures on tissue culture plastic. There is a critical need for more representative models of tumor complexity that can accurately predict therapeutic response and sensitivity. The development of three-dimensional (3D) ex vivo culture of patient-derived organoids (PDOs), derived from fresh tumor tissues, aims to address these shortcomings. Organoid cultures can be used as tumor surrogates in parallel to routine clinical management to inform therapeutic decisions by identifying potential effective interventions and indicating therapies that may be futile. Here, this procedure aims to describe strategies and a detailed step-by-step protocol to establish bladder cancer PDOs from fresh, viable clinical tissue. Our well-established, optimized protocols are practical to set up 3D cultures for experiments using limited and diverse starting material directly from patients or patient-derived xenograft (PDX) tumor material. This procedure can also be employed by most laboratories equipped with standard tissue culture equipment. The organoids generated using this protocol can be used as ex vivo surrogates to understand both the molecular mechanisms underpinning urological cancer pathology and to evaluate treatments to inform clinical management.

Introduction

Bladder cancer is the most prevalent urinary tract cancer and the tenth most common human malignancy worldwide¹. It encompasses a genetically diverse and phenotypically complex spectrum of disease². Urothelial non-muscle-invasive forms of bladder cancer (NMIBC) are the most common bladder cancer diagnoses (70%-80%), and these cancers display considerable biological heterogeneity and variable clinical outcomes^2,3,4. Patients with NMIBC typically experience a high risk of disease recurrence (50-70%) and one-third of cancers will progress and develop into significantly more aggressive muscle-invasive bladder cancer (MIBC)². Although 5-year survival rates for NMIBC are high (>90%), these patients must undergo long-term clinical management⁵. On the other hand, locally advanced (unresectable) or metastatic MIBC is generally considered incurable⁶. Consequently, bladder cancer has one of the highest lifetime treatment costs within cancer care and is a significant burden for both the individual and the healthcare system^3,7. The underlying genetic aberrations in advanced disease renders therapeutic management of bladder cancer a clinical challenge, and therapeutic options for invasive urothelial tumors have only recently improved since the approval of immunotherapies for both advanced and high-risk NMIBC^8,9. Currently, clinical decision-making has been guided by conventional clinical and histopathological features, despite individual bladder cancer tumors showing large differences in disease aggressiveness and response to therapy¹⁰. There is an urgent need to accelerate research into clinically useful models to improve the prediction of individual patient prognosis and identification of effective treatments.

Three-dimensional (3D) organoids show great potential as tumor models due to their ability to self-organize and recapitulate the original tumor's intrinsic in vivo architecture and pharmacogenomic profile, and their capability to mirror the native cellular functionality of the original tissue from which they were derived^11,12,13. Although established bladder cancer cell lines are readily available, relatively cost-effective, scalable, and simple to manipulate, the in vitro cell lines largely fail to mimic the spectrum of diverse genetic and epigenetic alterations observed in clinical bladder cancers^12,14 and were all established and maintained under 2D, adherent culture conditions. Additionally, cell lines derived from primary and metastatic bladder tumors harbor significant genetic divergence from the original tumor material. ^8,15.

An alternative approach is to use genetically engineered and carcinogen-induced mouse models. However, while these models recapitulate some of the natural oncogenic cascades involved in human neoplasia (reviewed in refs^16,17,18), they lack tumor heterogeneity, are expensive, poorly represent invasive and metastatic bladder cancer, and are not viable for rapid term drug testing as tumors can take many months to develop^14,19. Patient-derived models of cancer (including organoids, conditionally reprogrammed primary cell culture, and xenografts) provide invaluable opportunities to understand the effects of drug treatment before clinical treatment²⁰. Despite this, few groups routinely use these patient-proximal models due to limited access to fresh primary patient tissue and the extensive optimization required to reproducibly generate patient-derived organoid (PDO) culture conditions. In an in vivo setting, oncogenic cells can interact and communicate with various compositions of the surrounding constituents, including stromal cells, tissue infiltrating immune cells, and matrix¹². Similarly, for PDOs grown in a 3D format, cellular/matrix complexity can be customized to include other relevant components. PDOs can be rapidly generated and are often able to be passaged extensively or cryopreserved for later use, despite having a finite lifespan^21,22,23. Pharmacodynamics (i.e., response to a drug) can be evaluated using multiple read-outs, including organoid viability and morphology, and characterization of immunohistochemistry targets or transcriptional changes.

Here, the procedures for the establishment of bladder cancer organoids from patient material collected from transurethral resection of bladder tumor (TURBT) or surgical removal of the bladder (radical cystectomy) are described. The method to generate PDOs is illustrated, using readily available wet laboratory materials and tools. Endpoints include changes in cell morphological characteristics and viability. These were measured using fluorescence microscopy, in vitro viability (metabolic and cell membrane integrity) assays, and histopathological analysis. Figure 1 shows the workflow for establishing human bladder cancer PDOs from clinical material obtained during elective surgery.

Protocol

Patients have consented to this study following their admission under the Urology team at the Princess Alexandra Hospital, Brisbane, Australia. This study was performed in accordance with the principles of the Declaration of Helsinki and within ethical and institutional guidelines (ethics number HREC/05/QPAH/95, QUT 1000001165).

NOTE: As eligibility criteria, patients were aged ≥ 18 with cancer, and able to understand and provide consent. Those who were not able to give informed consent were excluded. Those having a primary language other than English were excluded as the provision of interpreters was not possible due to logistical and budgetary considerations. Also excluded were patients whose tumors were not accessible to biopsy or unlikely to be available in adequate amount after routine pathology.

1. Organoid Medium Preparation

NOTE: Human bladder cancer organoid medium requires growth factors that aid in the survival, growth, and continuous expansion of organoids derived from dissociated clinical material (Table 1). For complete details of each supplement used in this procedure, please refer to the Table of Materials.

1. Thaw frozen ingredients on ice or in a refrigerator at 2-8 °C. Avoid repeated freeze/thaw cycles and work from frozen aliquots (stored at -20 °C).
2. Basal medium: Prepare basal medium by supplementing advanced Dulbecco's Modified Eagle Medium/Ham's F-12 (adDMEM/F12) with HEPES (10 mM) and glutamine (2 mM). This is also used as a transport medium, and during organoid washing steps.
   NOTE: Advanced DMEM/F-12 is used as the basal medium to aid in the expansion of organoids in the presence of limited serum; however, it requires supplementation with HEPES and L-glutamine.
3. Briefly centrifuge fibroblast growth factor 10 (FGF-10), fibroblast growth factor 2 (FGF-2), epithelial growth factor (EGF), SB202190, prostaglandin E2 (PGE2), and A 83-01 before opening to ensure components are at the bottom of the vial.
4. Complete organoid medium: Supplement basal medium with human noggin- and R-spondin 1-conditioned media at a final concentration of 5% v/v, human EGF (50 ng/mL), human FGF-2 (5 ng/mL), human FGF-10 (20 ng/mL), A 83-01 (500 nM), SB202190 (10 µM), B27 (1x), nicotinamide (10 mM), N-acetylcysteine (1.25 mM), Y-27632 (10 µM), PGE2 (1 µM) and a broad-spectrum antibiotic formation at the concentration indicated by the manufacturer.
5. Store organoid media at 4 °C in the dark and use it within 2 weeks (1 month at most). Do not freeze. Avoid extended exposure to light sources.
   NOTE: The organoid medium is prepared serum-free; however, serum and penicillin/streptomycin could be supplemented on a user-to-user basis as required.

2. A day before procedure outlined in 3

1. Thaw growth factor reduced basement membrane (BME; see Table of Materials) overnight for at least 12 h before use in a 4 °C refrigerator or cold room. If required, dispense BME into 1 mL aliquots in a 1.5 mL polypropylene single-use tube to avoid freeze-thaw cycles.
2. Place filtered pipette tips into a 4 °C refrigerator or cold room.
   NOTE: This section refers to pipette tips that will be used when handling BME to prevent premature polymerization and reduce the coating of BME on the surface of the tips.
3. Sterilize all surgical equipment required for the procedure.

3. Generation of bladder tumor organoids

NOTE: This is an initial step for PDO establishment from primary patient tumors. This procedure is adapted for bladder cancer tissues from methods established by Gao et al.²⁴.

1. Use a class II biohazard hood to prepare the specimen. Retrieve dry and wet ice and print Patient Specimen Processing Sheet (Supplemental File).
2. Call research personnel to the operating room when surgical resection is near complete.
   NOTE: Confirm that the patient meets the eligibility criteria and has signed the participant consent form. Tissue is provided for research only if surplus to requirements for clinical histopathologic assessment.
3. Collect a fresh macroscopically viable tumor specimen from surgery. Ensure that the sample is submerged in transport medium (either 1x adDMEM/F12 or 1x Dulbecco's phosphate-buffered saline (DPBS)) in a sterile 50 mL conical tube or urine specimen jar during transit.
   NOTE: At some clinical centers, tissue may need to be transported to a pathology laboratory for to be allocated for research. Under these circumstances, it is recommended that antibiotics and antimycotics be added to the transport medium. Tissue can be stored at 4 °C in basal medium for up to 24 h post-surgery and still generate viable organoid cultures.
4. Record specimen details, including tissue weight (g or mg), sample description, and details regarding any blood and urine samples on the Patient Specimen Processing Sheet (Supplemental File).
5. Carefully remove the transport medium and replace it with 10 mL of basal media. Allow the tumor tissue to settle by gravity.
   NOTE: Transport medium is considered as clinical waste and must be collected in an appropriately labeled waste container in an appropriate amount of decontamination solution. Once the liquid has been chemically decontaminated, it can be disposed of according to institutional guidelines for hazardous waste.
6. Remove the tumor tissue with forceps and place it in a sterile 90 mm Petri dish (Figure 2A). Record the weight of the tissue in mg or g on the clinical specimen processing sheet and dissection grid (Figure 2B).
7. Remove non-cancerous tissue (including adipose tissue) and macroscopically visible necrotic regions using sterile forceps and disposable scalpel blade mounted to scalpel handle (Figure 2C). Wash tumor pieces 1-2 times with cold 1x DPBS. Collect the tumor pieces and transfer them to a new sterile 90 mm Petri Dish.
   NOTE: As scalpel blades are sharp, take caution when manually slicing. Tumor-adjacent adipose tissue can be identified by distinctly soft, gelatinous, pale areas immediately adjacent to the visual tumor perimeter. Macroscopic adipose tissue, and focally dark regions representing areas of necrosis will require personal judgment when dissecting from an excisional bladder tumor biopsy.
8. Take a photograph, draw a diagram of tissue, and plan tissue dissection on a clinical processing grid (Figure 2B and Figure 2C).
   NOTE: It is important to keep a visual record and rough sketch of the individual macroscopic tumor characteristics and dissection for each specific case.
9. Dissect tumor tissue pieces and allocate for histopathological (Figure 2D) and molecular analyses (Figure 2E).
   1. For histological analyses: Place approximately 50 mg of tumor tissue into a labeled disposable plastic histology cassette (Figure 2D). Submerge histology cassette into a container with 5x to 10x volume of 10% neutral buffered formalin (NBF). Incubate overnight at RT.
   2. Remove 10% NBF and replace with 70% (w/w) ethanol the next day for storage at 4 °C until tissue can be processed using routine tissue processing protocol
   3. For molecular analyses: Snap freeze at least one 1-3 mm³ tumor piece in an RNase/DNase-free 1.5 mL cryovial using liquid nitrogen and store at -80 °C (Figure 2E).
10. Dispense 5 mL of organoid medium (Table 1) in the 90 mm Petri dish containing the remaining tumor piece(s).
11. Mechanically mince tissue as finely as possible (0.5-1 mm³ pieces or smaller) with a sterile #10 scalpel blade.
    NOTE: Larger fragments or whole chunks of tissue (>3 mm³) will take considerably longer to digest and will decrease the viability of the specimen. Omit the above step if the tissue is disaggregated and fragments are small enough to pipette with a 5 mL serological pipette tip.
12. Transfer finely minced tissue to a 50 mL conical tube and add 4 mL of organoid medium, 1 mL of 10x collagenase/hyaluronidase, and 0.1 mg/mL deoxyribonuclease 1 (DNase 1) to avoid cell clumping.
    NOTE: Enzyme solution should be freshly prepared each time. Increase the volume of the medium to be approximately 10x the visible amount of the tumor fragments for large samples.
13. Incubate the minced tumor tissue and enzyme solution for 1-2 h on an orbital shaker or rotator (150 rpm) in an incubator (37 °C, 5% CO₂) to dissociate fragments into a cell suspension and break down collagens. This can be checked with histological analysis (Figure 3A).
    NOTE: If the amount of tissue is large or no obvious dissociation is observed after 1-1.5 h, increase the incubation time checking the level of dissociation every 30 min. Timing for this step depends on the sample and must be determined empirically each time. A noticeably clearer solution with tissue fragments indiscernible to the eye (or very few fragments) indicates successful digestion.
14. Terminate the digestion with the addition of 2x volume (20 mL) of basal medium to the sample.
15. Centrifuge the sample at 261 x g for 5 min at room temperature (RT), aspirate, and discard the supernatant.
16. To lyse contaminating red blood cells (RBCs), resuspend the pellet obtained from the above step in 5 mL of ammonium-chloride-potassium (ACK) buffer. Incubate the tube at RT for 3 min or until the complete lysis of the RBCs is seen (suspension becomes clear).
    NOTE: If RBCs are not observed as a small red clump in the pellet, this step may be omitted.
17. Add 20 mL of the basal medium into the tube. Centrifuge the tube at 261 x g for 5 min at RT and aspirate the supernatant.
18. At this step, place a 10 mL aliquot of 2x and 1x organoid medium in a 37 °C water bath to warm.
19. Filter the sample through a pre-wet reversible 100 µm strainer into a new 50 mL tube to remove large insoluble material.
    NOTE: Large undigested material (>100 µm) collected by the filter may contain cells of interest and can be cultured in 90 mm Petri dish or 6-well cell culture plate in organoid medium to derive 2D cultures (Figure 1 (step 5) and Figure 3B).
20. Filter the eluate through a pre-wet reversible 37 µm strainer to collect single cells and small clusters for single-cell and immune cell isolation (Tube 37-1; Figure 1 (step 6) and Figure 3B).
    NOTE: The yield of tumor cells during this step may be increased by passing the filtered cell suspension through the strainer again.
21. Reverse the 37 µm strainer and use 10 mL of basal media to collect small and moderate-sized clusters (37-100 µm) (tube 70-1; Figure 3B).
22. Top up each of the new 50 mL tubes (tubes 70-1 and 37-1) with DPBS to 40 mL. Centrifuge the suspension at 261 x g for 5 min at RT. Aspirate and discard the supernatant.
23. Add 10 mL of basal media to tube 37-1 and count the cells using trypan blue exclusion dye and an automated cell counter (as per the manufacturer's specifications). Determine the cell number and viability of cells.
24. Centrifuge the remainder of the sample at 261 x g for 5 min at RT. Aspirate the medium and replace it with cell freezing solution or basal media containing 10% fetal bovine serum (FBS), 1% penicillin/streptomycin, and 10% dimethyl sulfoxide (DMSO).
25. Place samples in 1.5 mL cryovials and store them in a cell-freezing container. Immediately transfer the container to a -80 °C freezer for optimal rate of cooling. Following overnight freezer storage at -80 °C, transfer the cryovials into cryogenic liquid or air phase storage (-196 °C) for long-term storage.
26. Resuspend cells from tube 70-1 with 500 µL of pre-warmed 2x organoid medium.
    NOTE: Seeding density must be high for successful organoid propagation. The volume of the organoid medium and, subsequently, BME should be altered empirically on the number of cells isolated from the filtration step. This step provides a relevant starting point based on studies in our laboratory.
27. Add BME to cells (at a 1:1 ratio with 2x organoid medium) with ice-cold P1000 sterile filter pipette tips and mix gently to suspend cells. Quickly and carefully pipette 100 µL of reconstituted cells/ BME mixture to wells of an ultra-low attachment flat-bottom 96-well plate. Place the 96-well plate in an incubator (37 °C, 5% CO₂) for 20-30 min to solidify.
28. Add 1:2 ratio of 1x organoid medium on top of BME cell suspension depending on the empirically assessed volume. In the relevant starting point, add 50 µL of 1x organoid medium media on top of 100 µL of reconstituted cells/ BME mixture.
29. Place the 96-well plate in an incubator (37 °C, 5% CO₂) for 20 min to equilibrate.
30. Take the cell culture plate out of the incubator and assemble it on a specimen holder on the stage of a microscope. Assess organoids visually under phase contrast or brightfield settings.
    NOTE: Images are best acquired by an inverted phase-contrast microscope equipped with differential interference (DIC) optics, digital camera, and associated software to observe the formation of spherical PDOs in preparation for endpoint analysis.
    1. If distinct clusters are visible, place the cell culture plate into an onstage electronically heated microscope chamber (37 °C, 5% CO₂) for live-cell imaging over the first 24-72 h.
    2. Ensure the flexible heated collar is attached to the lens to reduce thermal drift and the humidifier is filled with dH₂O.
    3. Perform the initial imaging using a 4x or 10x objective lens (N.A. 0.30, W.D. 15.2 mm). Time-lapse every 5-10 min on the phase-contrast setting.
    4. Check for the successful isolation as characterized by the appearance of >10 self-organizing organoids per well after a 24-72 h period.
    5. Allow cultures to continue for up to two weeks if organoid numbers are low (Figure 3C).
31. Top up the medium every 2-3 days using 50 µL of pre-warmed organoid medium to replenish depleted growth factors and overall volume.
32. Acquire images (as described in step 3.30) on days 1, 2, and 3 (time-lapse series), and on days 5, 7, and 10 before passaging (if applicable).
    NOTE: Organoids (observed as generally round structures where you cannot see the edges of individual cells) are typically passaged 7-10 days following setup, depending on the success of the isolation. Before or following passaging, organoids can be treated with cytotoxics or therapeutic agents for up to 6 days and drug response measured using cell viability and cytotoxicity assays. Alternatively, organoids can be retrieved from BME (using 1 mg/mL dispase) and cryopreserved (bio banked), prepared for molecular testing, or embedded and assessed histologically (Figure 4).

Representative Results

3D organoids were successfully established from human bladder cancer patient TURBT and cystectomy tissues. Briefly, this technique highlights a rapid formation of 3D multicellular structures that are both viable and suitable for other endpoint analyses such as histological evaluation, molecular characterization (by immunohistochemistry or quantitative real-time PCR), and drug screening. During the procedure (Figure 1), the various eluates during our filtration phases (Figure 1, steps 1-6) could be taken advantage of to cryopreserve single cells for isolation of tumor-infiltrating immune cells (TILs) and generation of a single cell biobank. The procedure allows for multiple aspects of a heterogeneous primary tumor to be biobanked appropriately and in a way that can be tracked across patients (Figure 2A,B), including histology (Figure 2C,D) and fresh frozen samples (Figure 2E).

A 2 h digestion with commercially available collagenase/hyaluronidase and DNase 1 allows for sufficient digestion of 0.5-1 mm³ pieces of the more complex cystectomy biopsy tissue, including the neoplastic cells within the lamina propria and muscular layers of the bladder (Figure 3A). Larger post-digestion fragments (>100 microns) have proven suitable for culture in standard tissue culture plates of any size for the isolation of novel 2D cultures (Figure 3B). However, extensive molecular characterization is required to validate the cancer origin of such cell lines. Organoids which are successfully generated with this protocol undergo processes of dynamic self-assembly, including phases of aggregation, compaction, and final formation, whereby they form into tight cellular structures (Figure 3C) that can be assessed for response to standard-of-care therapies. Figure 3C highlights the efficient self-assembly properties of the organoids. Histologically, these structures recapitulate the original patient tumor (Figure 4). Organoids generated in this procedure are suitable for a multitude of purposes, including further characterization, biobanking, and drug efficacy testing (Figure 5).

Figure 1: Schematic representation of organoid tissue culture from the collection of tumor samples from surgery to endpoint analyses. URN, unit record number; TIL, Tumor-infiltrating lymphocyte. (1) Tissue is mechanically minced in a 90 mm petri dish as finely as possible with a sterile scalpel blade (0.5-1 mm³ pieces). (2) Tumor pieces are resuspended in an enzymatic dissociation medium consisting of the organoid medium, collagenase/hyaluronidase, and DNase 1. (3) Minced tumor tissue and enzyme solution are incubated for 1-2 h on a rotator incubator (150 rpm, 37 °C, 5% CO₂) and subsequently pelleted (4) to dissociate fragments into a finer cell suspension. The sample undergoes filtration through pre-wet reversible (5) 100 µm and (6) 37 µm strainers into new 50 mL tubes to remove large insoluble material and isolate 37-100 µm clusters. Following (7) centrifugation of the fraction isolated from the reverse of the 37 µm strainer, the cell clusters are suspended in (8) BME and organoid media before (9) live-cell imaging. Created with BioRender.com. Please click here to view a larger version of this figure.

Figure 2: Sterile tumor tissue handling for organoid establishment. (A) Upon excision and transport to the laboratory, the specimen is placed in a sterile petri dish, weighed, and subsequently dissected. (B) A specimen dissection grid is used to mark the specimen for various processes. An example of dissection is shown in (C) where samples are taken from the tumor periphery as much as possible to avoid areas of central necrosis and annotated (inset) before (D) grossing the tissue prior to tissue fixation. Scale bar: 6 mm. Inset scale bar: 4 mm. (E) Small fresh fragments are taken for subsequent freezing and biobanking. Scale bar: 5 mm. Elements of this image were created with BioRender.com. Please click here to view a larger version of this figure.

Figure 3: Enzymatic derivation and dynamic assembly of bladder tumor organoids. (A) Representative pictures of hematoxylin and eosin (H&E) stained parental bladder cancer tissue (pathology reviewed as T3bN0) on immediate fixation (left) and 2 h post-digestion with 1x collagenase/ hyaluronidase (right). Scale bar: 500 µm. Inset scale bar: 50 µm. (B) Representative images of the >100 µm, 37-1 and 70-1 (day 2) isolations. Scale bars are indicated within the image. (C) Representative images are shown for bladder cancer organoid isolation over 48 h and day 7 organoid H&E staining. Scale bar: 50 µm. Scale bar for the image in the inset: 20 µm. DIC: differential interference contrast. Elements of this image were created with BioRender.com Please click here to view a larger version of this figure.

Figure 4: Bladder cancer-derived organoid cultures maintain the histological architecture of parental tumors. Representative H&E image of low-grade bladder cancer (Ta) and corresponding bright-field image of organoid at day 7 of culture. The far-right panel shows associated H&E staining of the day 7 organoid. Scale bars are indicated in the figure. LG: low grade, BC: bladder cancer, H&E: haematoxylin and eosin, and DIC: differential interference contrast. Please click here to view a larger version of this figure.

Figure 5: Downstream applications with bladder cancer patient-derived organoids. Created with BioRender.com. Please click here to view a larger version of this figure.

Additive		Final Conc.	Stock Conc.	Solvent
Basal Media	Glutamax	2 mM	1 M	NaCl
	HEPES	10 mM	1 M	NaCl/NaHPO4 buffered solution
Supplements for complete media	R-spondin 1 conditioned media	5% v/v	Conditioned Media	Advanced DMEM/F-12
	Noggin conditioned media	5% v/v	Conditioned Media	Advanced DMEM/F-12
	EGF	50 ng/mL	0.5 mg/mL	PBS/0.1% BSA
	FGF-10	20 ng/mL	100 µg/mL	PBS/0.1% BSA
	FGF-2	5 ng/mL	50 µg/mL	PBS/0.1% BSA
	Nicotinamide	10 mM	1 M	1 g in 8.2 mL ddH20
	N-acetyl-L-cysteine	1.25 mM	500 mM	40 mL ddH20
	A 83-01	0.5 µM	50 mM	DMSO
	SB202190	10 µM	50 mM	DMSO
	Y27632	10 µM	100 mM	ddH20
	B-27 additive	1X	50x
	Prostaglandin E2	1 µM	10 mM	DMSO
	Primocin	100 µg/mL	50mg/mL

Table 1: Supplements used for basal and complete organoid media

Supplemental File: Please click here to download this File.

Discussion

While 3D organoid protocols derived from bladder cancer tissue are still in their infancy, they are an area of active research and clinical investigation. Here, an optimized protocol to successfully establish bladder cancer PDOs that are suitable for both NMIBC and MIBC specimens is detailed. This workflow integrates parallelly into hospital-based clinical trials and considers biobank sample accrual, including histological sample processing and fresh frozen tissue banking, which is an important consideration for clinical organoid pipelines. This protocol builds on existing methods and provides a consolidated methodology to build an organoid precision medicine pipeline.

The current success rate of this protocol - like others - is approximately 70%²⁵. It is anticipated that this will improve as isolation techniques and characterization of the bladder cancer tumor microenvironment evolves. As expected, the quality of the initial specimen influences the success rate and is an important limiting step. Specimens obtained from chemotherapy naïve patients provide the highest success rate, while those from patients who have received neoadjuvant chemotherapy may have a reduced number of viable cells^26,27. Typically, bladder cancer specimens (from both TURBT and cystectomies) provide an ample amount of samples with high tumor cellularity. This allows robust generation of organoids ranging between 30-100 µm within a 24-72 h growth period (given some <40 µm clusters are retained in the filtration steps). Importantly, the heterogeneity observed in organoid samples (i.e., cystic formations and more dense structures) indicates that this protocol provides conditions suitable to derive heterogeneous tumor cell populations.

Our optimized protocol can be performed on limited starting material (as is the case from some elective surgeries, including TURBT procedures or specimens acquired from patients on a clinical trial), enabling the incorporation of multiple analyses to maximize specimen value. This protocol includes methods to describe steps to grow 2D culture from extraneous tumor cells of interest that may reside in large insoluble tissues processed during the initial filtration step. Like other research groups which suggest processing specimens within 24 h post-surgery^28,29, it is observed that tissue hypoxia that occurs within this timeframe does not preclude generating viable organoids. Optimization is required to determine the density of cell (native clusters) seeding per well. In our experience, excessive numbers of clusters can result in poor growth of the cultures, while low visual density cultures can lead to cell death. This can be determined by the user empirically in the context of their volumes which can be diluted within a range as necessary. Additionally, TURBT specimens often contain burnt edges due to the resection process, which are apparent as prominent necrotic regions. A critical step is the careful isolation of macroscopic tumor tissue and subsequent dissection. This is paramount to deriving viable cultures and needs to be communicated with the surgical team. In some instances, this may be an initial limitation to the procedure.

An additional limitation to the current technique is the absence of the tumor microenvironment parallel to the organoids. A necessary development will be future co-culture systems to increase cellular complexity and provide other components of the tumor microenvironment (i.e., stroma and immune cells). As organoids are defined as self-organizing structures with the ability to self-perpetuate their growth, further work will be required to elucidate the discrete cancer stem-cells (such as CD44 and CD49) which may allow for improved in vitro growth characteristics^31,32,33. Establishing with certainty that the PDOs originate from tumor tissue is critical, as the contribution from normal urothelial cells in functional and molecular analyses will confound investigations into drug resistance. Sometimes the tumor origin of the urothelial cells in the PDOs is clear due to the use of metastatic tissue or papillary tumors (as is the case here; see Figure 4), but when the source tissue is excised from the bladder mucosa and thus contains margins of the benign urothelium, it is critical to confirm that the resultant PDOs are comprised of tumor cells using molecular techniques. Thus, immunohistochemical and genomic comparisons are required between the established organoids and the primary tumor from which they were derived³⁰.

Regarding the number and size distributions of the organoids per well, it is difficult to maintain uniform plating density of 3D organoids for cytotoxicity analysis. This is slightly mitigated by isolating defined ranges using the filters, but it is still a limitation if small clusters and larger organoids are isolated in the same well. Large particle sorters may mitigate this issue but are not widely available in medical research laboratories. Nonetheless, techniques that measure viability signals before and after treatment have been used successfully in bladder cancer organoids³⁴. As this technology advances, this will enable researchers to develop a library of bladder cancer PDOs with varying genomic profiles, and pharmacodynamic responses that can be explored to investigate novel therapies. Additionally, the specimen collection procedure and dissociation method described herein allow concurrent preparations for tumor spatial profiling, multiparametric flow cytometry, and single-cell RNA sequencing. Taken together, this allows for a diverse range of powerful endpoints which can be brought together to explore a patient's tumor heterogeneity and applicability of drugs such as immunotherapies.

In summary, we describe a protocol to establish and evaluate cultures of 3D patient-derived bladder cancer organoids in the advent of precision medicine endpoints. It is important to note that the steps and notes described here are routine approaches used to isolate bladder cancer PDOs in our laboratory. The culture medium used in our procedure was originally described for prostate cancer cells and has now been proven to be suitable for the culture of bladder cancer. Importantly, this protocol is centralized, suitable for routine use in laboratories, and broadly applicable across urological cancer types. In combination with high fidelity molecular phenotyping and drug response endpoints, this technique will provide a useful tool to rapidly explore the precise therapeutic management of bladder cancer patients.

Materials

Name	Company	Catalog Number	Comments
1.2 mL cryogenic vial	Corning	430487
1.5 mL Eppendorf tubes	Sigma-Aldrich	T9661-500EA	polypropylene single-use tube
100 µM reversible strainer	STEMCELL Technologies	#27270
100 mm Petri dish	Corning	430167
10x Collagenase/ Hyaluronidase	STEMCELL Technologies	#07912
37 µM reversible strainer	STEMCELL Technologies	#27250
37°C incubator
37°C water bath
50 mL falcon tube	Corning	CLS430829-500EA
6-well plate	Corning	CLS3516
70% (w/w) ethanol
-80°C freezer
96 well ultra low attachment plate (Black)	Sigma-Aldrich	CLS3474-24EA
A 83-01	BioScientific	2939	Prevents the growth-inhibitory effects of TGF-β
ACK lysis buffer	STEMCELL Technologies	#07850
adDMEM/F-12	Thermo Fisher Scientific	12634028	Base medium
Animal-free recombinant EGF	Sigma-Aldrich	518179	Growth factor
Automated cell counter (TC20)	Bio-rad	1450102
B27 additive	Gibco	17504044	Increases sphere-forming efficiency
Cell Counting Slides	Bio-rad	1450015
Centrifuge	Eppendorf	EP022628188
Computer system
CryoStor CS10	STEMCELL Technologies	#07930	Cell freezing solution
Dispase II, powder	Thermofisher	17105041	To enzymatically disrupt Matrigel
DNAse 1	STEMCELL Technologies	#07900
DPBS	Thermofisher	14190144
Dry and wet ice
Esky
Farmdyne (Iodine 16g/L)	Ecolab
Formalin solution, neutral buffered, 10%	Sigma	HT501128	Histological tissue fixative
Glutamax (L-alanine-L-glutamine)	Invitrogen	35050061	Source of nitrogen for the synthesis of proteins, nucleic acids
HEPES	Gibco	15630-080	All-purpose buffer
Histology cassette	ProSciTech	RCH44-W
Human FGF-10	Peprotech	100-26-25	Growth factor
Human FGF-2	Peprotech	100-18B-50	Growth factor
Liquid nitrogen
Matrigel (Growth Factor Reduced (GFR), phenol red-free, LDEV free)	In Vitro Technologies	356231	Basement membrane extract (BME)
Mr. Frosty freezing container	ThermoFisher	5100-0001	cell freezing container
N-acetyl-L-cysteine (NAC)	Sigma	A7250	Anti-oxidant required to protect against ROS-induced cytotoxicity
Nicotinamide	Sigma	N0636-100G	SIRT-1 inhibitor
NikonTs2U inverted microscope	Nikon	MFA510BB
NIS-Elements Advanced Research	Nikon	MQS31000
Noggin conditioned media	In-house		BMP inhibitor
Pipetboy acu 2	Integra	155000
Pipettes (p20, p100, p1000) with tips
Primocin	Jomar Bioscience	ant-pm2	Combination of antibacterial and antifungal compounds to protect cell cultures from contaminations
Prostaglandin E2 (PGE2)	Tocris	2296	support proliferation of cells
Rotary tube mixer	Ratek	RSM7DC
R-spondin 1 conditioned media	In-house		WNT signalling regulator
SB202190	Jomar Bioscience	s1077-25mg	Selective p38 MAP kinase inhibitor
Scale
Scalpel handle	Livingstone	WBLDHDL03
Scalpels, #11 blade	Medical and Surgical Requisites	EU-211-1
Serological pipettes (5, 10, 25 mL)
Specimen Waste Bags	Medical Search	SU09125X16
Urine specimen jar
Y27632	Jomar Bioscience	s1049-10mg	Selective ROCK inhibitor. Increases survival of dissociated epithelial cells