An Optimized Protocol for the Generation of Vascular Organoids from Human Induced Pluripotent Stem Cells

Summary

This protocol presents a method for generating vascular organoids using human induced pluripotent stem cells.

Abstract

Vascular organoids derived from human pluripotent stem cells (hPSCs) have emerged as powerful three-dimensional models for studying vascular development, disease mechanisms, and drug responses. Current vascular organoid protocols enable the generation of self-organizing endothelial–pericyte networks; however, batch-to-batch variability, inconsistent organoid formation, and inadequate pericyte coverage can compromise reproducibility and scalability. This study presents an optimized protocol for generating vascular organoids from human induced pluripotent stem cells (hiPSCs) with improved reproducibility and structural consistency. Key modifications include refined embryoid body formation conditions, optimized growth factor concentrations and timing during mesoderm induction and vascular specification, and improved three-dimensional culture conditions that promote robust pericyte recruitment and endothelial–pericyte interactions. The optimized organoids exhibit highly branched CD31-positive endothelial networks consistently ensheathed by PDGFR-β-positive pericytes, with significantly reduced batch-to-batch variability compared with the original protocol. Detailed, step-by-step procedures are provided for organoid generation, whole-mount immunofluorescence characterization, and quality control assessment. This optimized method enables the scalable production of high-quality vascular organoids suitable for studying vascular cell interactions, modeling disease, and screening compounds, thereby lowering the technical barrier for laboratories seeking to adopt this model system.

Introduction

Three-dimensional (3D) organoids have emerged as powerful tools for modeling human organ development and disease in vitro, offering superior structural biomimicry and pathophysiological relevance compared with conventional two-dimensional cell culture models^1,2. In recent years, human pluripotent stem cell (hPSC)-derived organoids have been successfully established for multiple tissue types, including the intestine³, brain^4,5, kidney⁶, liver^7,8, and retina⁹. However, a major challenge in organoid-based studies is insufficient vascularization during long-term culture, which can lead to hypoxia and necrosis, thereby impeding organoid expansion and differentiation^10,11. To address this limitation, various co-culture strategies have been introduced, including incorporating endothelial cells or mesenchymal stem cells to promote vascular network formation^12,13. However, conventional two-dimensional cultures and animal models do not fully recapitulate human vascular biology, whereas co-culture approaches introduce additional experimental complexity¹⁴. Vascular organoids capable of autonomous vascularization, therefore, offer a promising human-relevant alternative¹⁵.

Vascular network formation involves both vasculogenesis and angiogenesis, which are orchestrated through interactions between endothelial cells and mural cells, such as pericytes and vascular smooth muscle cells, and are tightly regulated by multiple signaling molecules, including VEGF (vascular endothelial growth factor), TGF-β (transforming growth factor-β), PDGF (platelet-derived growth factor), and angiopoietins¹⁶. In 2019, Wimmer and colleagues established a method for generating self-assembling 3D blood vessel organoids directly from hPSCs^17,18. This system simultaneously generates endothelial cells and pericytes, resulting in microvascular networks with lumen formation and basement membrane coverage. Building on this platform, vascular organoids have been used to model diabetic vasculopathy under high-glucose and pro-inflammatory cytokine stimulation (TNF-α and IL-6); these organoids recapitulate basement membrane thickening, a hallmark of diabetic microvascular pathology¹⁷. In addition, this model has been used to study hereditary vascular diseases and to serve as a platform for vascular toxicity screening¹⁹. More recently, several method-oriented studies have sought to optimize vascular organoid generation. For example, advances in single-cell RNA sequencing have identified key regulatory factors, such as MECOM and LEF1, thereby providing a molecular framework for characterizing and optimizing human vascular organoid formation. In addition, Xu et al. introduced microwells to facilitate homogeneous embryoid body formation and incorporated the CEPT cocktail to improve cell viability, thereby effectively reducing batch-to-batch variability²⁰.

To improve the homogeneity, reproducibility, and throughput of vascular organoid generation, the original protocol described by Wimmer et al.^17,18 was systematically optimized to address several key limitations. First, the original protocol employs natural aggregation in low-attachment plates for embryoid body formation, which can result in substantial batch-to-batch variability in aggregate size and, consequently, heterogeneous differentiation efficiency. In the optimized protocol, polyvinyl alcohol (PVA) is combined with orbital shaking during aggregate formation to promote uniform cell aggregation and improve consistency in embryoid body size. Second, the original protocol uses complete medium replacement during the vascular network formation stage, which may cause substantial fluctuations in growth factor concentrations and compromise sustained vascular sprouting and pericyte recruitment. In contrast, the optimized protocol retains one-third of the conditioned medium during each medium change to maintain endogenous pro-angiogenic factors and stabilize the culture environment. Third, the original protocol requires multiple plate transfers during organoid isolation and maturation, increasing handling requirements and the risk of organoid loss. The optimized protocol bypasses these steps by transferring isolated organoids individually into wells of 96-well low-attachment plates, thereby simplifying the procedure and increasing throughput. Collectively, this optimized protocol provides detailed step-by-step procedures and critical quality-control checkpoints to lower the technical barrier to adopting this model system and facilitate the broader application of vascular organoids in studies of vascular development, disease mechanisms, and high-throughput drug screening.

Protocol

The use of hPSCs in this study was approved by the Biomedical Ethics Committee of Anhui University (BECAHU_2026-010).

1. Maintenance of human induced pluripotent stem cells

NOTE: Maintain human induced pluripotent stem cells (hiPSCs) under feeder-free conditions. Perform all procedures in a Class II biological safety cabinet using aseptic technique.

1. Preparation of basement membrane matrix-coated culture plates
   1. Remove a vial of basement membrane matrix from −20 °C and thaw it at 4 °C overnight for approximately 12–16 h. Avoid repeated freeze–thaw cycles.
   2. Mix the thawed basement membrane matrix thoroughly under sterile conditions and divide it into single-use aliquots (e.g., 1 mL per tube). Store the aliquots at −20 °C for up to 2 years.
      NOTE: Keep basement membrane matrix, pipettes, tips, and tubes cold during handling to prevent premature solidification. Avoid introducing air bubbles.
   3. Thaw the required basement membrane matrix aliquot on ice. Dilute the basement membrane matrix 1:200 in prechilled DMEM (5 µL of basement membrane matrix per 1 mL of DMEM; final concentration, approximately 5 µg/mL).
   4. Mix the solution gently by pipetting until no visible particulates remain.
   5. Add 0.5 mL of diluted basement membrane matrix to each well of a 12-well plate. Gently tilt and rotate the plate to coat the entire bottom surface.
   6. Incubate the coated plate at 37 °C in a 5% CO₂ incubator for at least 2 h or at 4 °C overnight.
      NOTE: Store sealed coated plates at 4 °C for 1–2 weeks. Equilibrate the plate to room temperature and aspirate the coating solution immediately before use.
2. Thawing of hiPSCs
   1. Remove a cryovial of frozen hiPSCs from liquid nitrogen and immediately place it in a 37 °C water bath. Gently agitate the vial and remove it within 1 min, when only a few small ice crystals remain.
      NOTE: Avoid prolonged exposure to dimethyl sulfoxide (DMSO) at 37 °C and incomplete thawing, as both can reduce cell viability.
   2. Disinfect the outside of the cryovial with 70% ethanol and open it inside the biological safety cabinet.
   3. Slowly transfer the cell suspension dropwise into a 15 mL conical tube containing 3 mL of prewarmed chemically defined hPSC maintenance medium, or an equivalent chemically defined maintenance medium. Mix gently during transfer to dilute the DMSO gradually and minimize osmotic stress.
   4. Centrifuge the cells at 300 × g for 3 min at room temperature (22–24 °C) using a swinging-bucket rotor.
   5. Carefully aspirate the supernatant without disturbing the cell pellet. Gently resuspend the pellet in 1 mL of prewarmed chemically defined hPSC maintenance medium.
      NOTE: Use a P1000 pipette and pipette gently 2–3 times to minimize mechanical shear and bubble formation.
3. Plating and culture of hiPSCs
   1. Remove the precoated 12-well plate from the incubator and aspirate the coating solution. Do not wash the wells.
   2. Seed approximately 1 × 10⁵–2 × 10⁵ viable cells per well for routine recovery after thawing. Adjust the density according to the hiPSC line and experimental requirements.
   3. Dispense the cell suspension slowly along the wall of each well without touching the coated surface with the pipette tip.
   4. Add Y-27632 to a final concentration of 5 µM to promote cell survival and attachment after thawing.
      NOTE: Use Y-27632 only in the initial plating medium. Replace it with Y-27632-free chemically defined hPSC maintenance medium the following day.
   5. Incubate the cells at 37 °C with 5% CO₂ and saturated humidity.
   6. Examine cell attachment and morphology after approximately 16–24 h. Aspirate the medium and replace it with fresh, prewarmed chemically defined hPSC maintenance medium without Y-27632.
   7. Replace the medium daily or every other day until the cells reach approximately 70%–80% confluency for passaging or differentiation.
      NOTE: Examine the cultures daily. Use colonies with compact, well-defined borders, a high nuclear-to-cytoplasmic ratio, and minimal spontaneous differentiation.

2. Generation and differentiation of hiPSC aggregates

NOTE: Maintain hiPSCs under feeder-free conditions in chemically defined hPSC maintenance medium on basement membrane matrix-coated plates. Passage the cells every 4–5 days using 0.02% EDTA. Use cultures with appropriate seeding density and colony size and minimal spontaneous differentiation to maximize vascular differentiation efficiency.

1. Preparation of a single-cell suspension
   1. Aspirate the chemically defined hPSC maintenance medium and wash the hiPSCs with 0.5 mL of DMEM per well of a 6-well plate.
   2. Aspirate the DMEM and add 0.5 mL of 0.02% EDTA per well. Incubate at room temperature for 2–4 min and monitor the cells until they begin to round up.
   3. Detach the cells using a P1000 pipette. Pipette gently 2–3 times to dissociate the aggregates into a single-cell suspension.
   4. Resuspend the cells in 5 mL of chemically defined hPSC maintenance medium and transfer the suspension to a 15 mL conical tube.
   5. Mix 10 µL of the cell suspension with 10 µL of 0.1% Trypan blue solution. Load 10 µL of the mixture onto a counting chamber and determine the number of viable cells.
   6. Calculate the volume of cell suspension required to obtain 5 × 10⁵ viable cells per well.
   7. Centrifuge the cells at 300 × g for 5 min at room temperature (22–23 °C). Aspirate the supernatant.
   8. Resuspend the cells in fresh chemically defined hPSC maintenance medium containing 50 µM Y-27632 at a volume of 3 mL per well.
2. Aggregate formation
   1. Add 3 mL of the cell suspension to each well of a low-attachment 6-well plate. Add PVA to a final concentration of 0.25%.
   2. Place the plate on an orbital shaker at 50 rpm and incubate at 37 °C for 1 day.
   3. Confirm the formation of aggregates with smooth borders and a diameter of approximately 50–100 µm or greater before proceeding.
   4. NOTE: Optimize the aggregation period for individual hiPSC lines. Extend incubation to 2–3 days when necessary to obtain aggregates of the desired size.
3. Mesoderm induction
   1. On Day 0, transfer the aggregates to a 15 mL conical tube using a sterile 10 mL disposable pipette. Retain the original low-attachment plate for replating.
   2. Allow the aggregates to settle by gravity for approximately 15–20 min.
   3. NOTE: Handle aggregates gently and use gravity sedimentation rather than centrifugation to minimize mechanical damage.
   4. Carefully aspirate the supernatant after the aggregates have settled.
   5. Resuspend the aggregates in 3 mL per well of N2B27 medium supplemented with 12 µM CHIR99021 and 30 ng/mL BMP-4.
   6. Transfer the aggregates back to the low-attachment plate and incubate at 37 °C.
   7. Pipette the aggregates gently once daily with a P1000 pipette to minimize excessive aggregate fusion. Continue mesoderm induction for 3 days.
4. Vascular lineage induction
   1. On Day 3, transfer the aggregates to a 15 mL conical tube and allow them to settle by gravity. Retain the original 6-well plate.
   2. Carefully aspirate the supernatant after the aggregates have settled.
   3. Resuspend the aggregates in 3 mL per well of N2B27 medium supplemented with 100 ng/mL VEGF-A and 2 µM forskolin.
   4. Transfer the aggregates back to the low-attachment 6-well plate and incubate for an additional 2 days.

3. Generation of vascular networks and organoids

1. Preparation of the collagen I–basement membrane matrix
   1. Collect the aggregates generated in one 6-well plate for embedding into one 12-well plate. Use approximately 40–60 aggregates per well.
   2. Prepare sufficient collagen I–basement membrane matrix mixture to add two 0.5 mL layers to each well of the 12-well plate.
   3. Prepare 12 mL of matrix for one 12-well plate. Keep all components on ice, prepare the mixture immediately before use, and avoid introducing air bubbles.
   4. Add 0.5 mL of matrix to each well of the 12-well plate. Gently swirl the plate to distribute the matrix evenly.
   5. Incubate the plate at 37 °C for 2 h to allow the first matrix layer to solidify.
2. Embedding of aggregates
   1. Transfer the aggregates to a 15 mL conical tube using a sterile 10 mL disposable pipette and allow them to settle by gravity.
   2. Aspirate the supernatant and wash the aggregates twice with 1 mL of N2B27 medium. Remove as much residual medium as possible after the final wash.
   3. Prepare a fresh collagen I–basement membrane matrix mixture and keep it on ice.
   4. Resuspend the washed aggregates gently in the freshly prepared matrix.
   5. Remove the 12-well plate containing the solidified first layer from the incubator.
   6. Add 0.5 mL of the aggregate-containing matrix to each well.
   7. Rock the plate gently for no longer than 1 min to distribute the aggregates evenly.
   8. Immediately incubate the plate at 37 °C for 2 h to allow the second layer to solidify.
3. Induction of vascular sprouting
   1. Prewarm N2B27 medium supplemented with 15% FBS, 100 ng/mL VEGF-A, and 100 ng/mL FGF-2 to 37 °C.
   2. Add the prewarmed supplemented medium to each well to induce vascular differentiation.
   3. Monitor the cultures for vascular sprouting over the next 1–3 days.
   4. NOTE: Prewarm the medium to 37 °C before adding it to the cultures to minimize gel matrix destabilization.
4. Medium exchange and culture maintenance
   1. Perform the first medium change 3 days after embedding.
   2. Replace the medium every other day thereafter using fresh N2B27 medium containing 15% FBS, 100 ng/mL VEGF-A, and 100 ng/mL FGF-2.
   3. Retain one-third of the conditioned medium during each medium change and replace the remaining two-thirds with fresh supplemented medium.
5. Proceed with vascular organoid isolation on Day 10.

4. Establishment of vascular organoids and preparation for staining

1. Isolation of individual vascular networks
   1. Insert the rounded end of a sterile spatula along the edge of the well and gently detach the gel matrix from the bottom.
   2. Continue moving the spatula from the well edge toward the center until the entire gel matrix floats freely.
   3. Use two sterile needles to excise individual vascular networks from the gel. Remove as much excess matrix as possible without damaging the vascular network.
2. Culture of isolated organoids
   1. Transfer each isolated organoid directly into an individual well of a low-attachment 96-well plate using a sterile disposable pipette.
   2. Add 100 µL of N2B27 medium supplemented with 15% FBS, 100 ng/mL VEGF-A, and 100 ng/mL FGF-2 to each well.
   3. Incubate the organoids for an additional 5 days and replace the supplemented medium daily.
   4. Confirm that the organoids exhibit a rounded and fully encapsulated morphology by Day 5 of maturation.
3. Fixation of vascular organoids
   1. Transfer morphologically mature organoids to 2 mL microcentrifuge tubes. Process no more than approximately 60 organoids per tube.
   2. Carefully aspirate the medium and add 4% paraformaldehyde (PFA) in PBS.
   3. Fix the organoids at room temperature for 1 h.
      CAUTION: PFA is toxic and a suspected carcinogen. Handle PFA in a chemical fume hood while wearing appropriate personal protective equipment.
   4. Aspirate the PFA and replace it with PBS.
      PAUSE POINT: Store fixed vascular organoids in PBS at 4 °C for up to 1 month.

5. Staining and confocal imaging of vascular organoids

1. Blocking and permeabilization
   1. Add 2 mL of blocking buffer to each tube containing fixed vascular organoids.
   2. Position the tubes vertically on an orbital shaker and incubate with agitation at room temperature for 2 h to block and permeabilize the organoids.
2. Primary antibody incubation
   1. Aspirate the blocking buffer and add 50–100 µL of primary antibody diluted 1:100 in blocking buffer.
   2. Position the tubes upright on a shaker and incubate with agitation at 4 °C overnight.
3. Washing and secondary antibody incubation
   1. Wash the samples three times with PBST for 10–15 min per wash. Agitate the vertically positioned tubes during each wash.
   2. Add fluorescent secondary antibody diluted 1:300 in blocking buffer.
   3. Incubate the samples at room temperature for 2 h.
4. Nuclear counterstaining and final washing
   1. Wash the samples twice with PBST for 10–15 min per wash.
   2. Add Hoechst solution diluted 1:5,000 in PBS and incubate at room temperature for 10–15 min.
   3. Wash the samples with PBS for 10–15 min.
5. Confocal imaging
   1. Transfer the stained organoids to confocal imaging dishes.
   2. Acquire confocal images using consistent imaging parameters across experimental groups.
6. Quantitative image analysis of vascular organoids
   NOTE: Perform quantitative image analysis using Fiji/ImageJ and AngioTool. Maintain identical microscope acquisition settings across all experimental groups.
   1. Aggregate diameter measurement
      1. Acquire bright-field images of Day 0 hiPSC aggregates using an inverted microscope with a calibrated scale. Image well-separated aggregates and include a scale bar.
      2. Import each image into Fiji/ImageJ. Draw a line along the scale bar using the Straight Line tool and select Analyze > Set Scale.
      3. Enter the known distance and unit (e.g., 100 µm) and select Global to apply the calibration to all images in the session.
      4. Draw a line across the widest diameter of each aggregate using the Straight Line tool. Select Analyze > Measure or press Ctrl+M and record the diameter in µm.
      5. Measure at least 30 aggregates per batch.
      6. For nonspherical aggregates, outline each aggregate using the Oval or Polygon Selection tool. Select Analyze > Set Measurements > Feret's diameter and record the measurement.
      7. Export the measurements from the Results table and compile the data in a spreadsheet.
      8. Calculate the mean, standard deviation (SD), and coefficient of variation (CV) as CV = SD/mean × 100% for each batch.
   2. Organoid diameter measurement
      1. Acquire bright-field images of mature vascular organoids on Day 15 using an inverted microscope with a calibrated scale. Include a scale bar in each image.
      2. Import each image into Fiji/ImageJ and calibrate the scale as described in step
      3. Draw a line across the widest diameter of each organoid using the Straight Line tool. Select Analyze > Measure or press Ctrl+M and record the diameter in µm.
      4. Measure at least 30 organoids per batch.
      5. For irregularly shaped organoids, measure the longest and shortest diameters and calculate their average, or determine Feret's diameter as described in step 5.6.1.6.
      6. Export the measurements and calculate the mean, SD, and CV for each batch.
   3. Quantification of CD31-positive vascular area
      1. Acquire confocal z-stack images of Day 15 whole-mount organoids stained for CD31. Use identical laser power, gain, offset, and pinhole settings for all samples.
      2. Generate maximum-intensity projections (MIPs) from the z-stacks using the microscope software or Fiji/ImageJ.
      3. Apply an intensity threshold to segment the CD31-positive vascular network while excluding background signal.
      4. Select Process > Noise > Remove Outliers and use a radius of 2–5 pixels to remove small artifacts. Select Process > Binary > Fill Holes to fill small gaps within the vascular network.
      5. Select Analyze > Set Measurements and enable Area and Area fraction. Select Analyze > Measure to determine the vascular area.
      6. Calculate the CD31-positive vascular area as a percentage of the total organoid area.
      7. Analyze at least three organoids per batch from three independent batches. Export the measurements for statistical analysis.
   4. Quantification of PDGFR-β-positive pericyte coverage
      1. Acquire confocal z-stack images of Day 15 whole-mount organoids stained for CD31 and PDGFR-β. Generate MIPs from the z-stacks.
      2. Import the CD31 and PDGFR-β MIPs into Fiji/ImageJ and verify image alignment.
      3. Threshold the CD31 channel to generate a binary mask defining the vascular network.
      4. Threshold the PDGFR-β channel using the same segmentation approach to generate a binary mask of the pericyte signal.
      5. Select Process > Image Calculator and apply the AND operation to the CD31 and PDGFR-β masks to identify overlapping pixels.
      6. Measure the area of the overlapping signal.
      7. Calculate pericyte coverage as (PDGFR-β-positive area within CD31-positive vessels/total CD31-positive vascular area) × 100%.
      8. Analyze at least three organoids per batch from three independent batches. Export the measurements for statistical analysis.
   5. Branching index analysis using AngioTool
      1. Acquire confocal z-stack images of Day 15 whole-mount organoids stained for CD31. Generate MIPs from the z-stacks.
      2. Launch AngioTool and load the MIP image.
      3. Set the vessel diameter according to the typical vessel diameter observed in the images.
      4. Adjust the intensity threshold to segment the vascular network while excluding the background signal. Set the minimum vessel size to exclude small artifacts.
      5. Run the analysis and record vessel area, vessel density, total vessel length, number of junctions, and branching index.
      6. Verify the output image by confirming that the vasculature is outlined in yellow, the vascular skeleton is displayed in red, and branching points are marked with blue dots.
      7. Export the quantitative results from AngioTool.
      8. Analyze at least three fields of view per batch from three independent batches and compile the data for statistical analysis.

6. Quantitative real-time PCR analysis of vascular organoids

NOTE: Perform all RNA isolation and downstream procedures under RNase-free conditions.

1. RNA extraction
   1. Collect 5–10 mature Day 15 vascular organoids in a 1.5 mL RNase-free microcentrifuge tube. Remove the residual medium completely.
      NOTE: Pool multiple organoids when necessary to obtain sufficient RNA.
   2. Add 350 µL of lysis buffer supplemented with 1% β-mercaptoethanol.
   3. Pipette the samples 10–15 times until the organoids are completely lysed.
   4. Transfer the lysate to a genomic DNA elimination spin column positioned in a collection tube. Centrifuge at 11,000 × g for 30 s at room temperature.
   5. Discard the column and retain the flow-through.
   6. Add an equal volume of 70% ethanol, approximately 350 µL, to the flow-through and mix thoroughly by pipetting.
   7. Transfer up to 700 µL of the mixture to an RNA spin column positioned in a collection tube. Centrifuge at 11,000 × g for 30 s and discard the flow-through.
   8. Add 350 µL of wash buffer 1 to the column. Centrifuge at 11,000 × g for 30 s and discard the flow-through.
   9. Prepare the DNase I mixture by combining 10 µL of DNase I with 70 µL of DNase digestion buffer per sample.
   10. Add 80 µL of the DNase I mixture directly to the column membrane and incubate at room temperature for 15 min.
   11. Add 350 µL of wash buffer 1 to the column. Centrifuge at 11,000 × g for 30 s and discard the flow-through.
   12. Add 500 µL of wash buffer 2 to the column. Centrifuge at 11,000 × g for 30 s and discard the flow-through.
   13. Repeat the wash with 500 µL of wash buffer 2.
   14. Centrifuge the empty column at 11,000 × g for 2 min to dry the membrane.
   15. Transfer the column to a fresh 1.5 mL RNase-free microcentrifuge tube. Add 30–50 µL of RNase-free water directly to the membrane and incubate at room temperature for 1 min.
   16. Centrifuge at 11,000 × g for 1 min to elute the RNA. Store the RNA at −80 °C until use.
       NOTE: Assess RNA concentration and purity by spectrophotometry. Use an A260/A280 ratio of 1.8–2.1 as the acceptable range.
2. cDNA synthesis
   1. Prepare the reverse- reaction mixture on ice.
   2. Combine 600 ng of total RNA with 4 µL of RT master mix containing reverse transcriptase, buffer, dNTPs, and random primers. Add RNase-free water to a final volume of 20 µL.
   3. Incubate the reaction at 37 °C for 15 min, followed by 85 °C for 5 s.
   4. Store the resulting cDNA at −20 °C until use.
3. Quantitative real-time PCR
   1. Prepare 9 µL of qPCR master mix per reaction by combining 5 µL of 2× SYBR Green master mix, 3 µL of RNase-free water, and 1 µL of premixed forward and reverse primers at 10 µM each.
   2. Dispense 9 µL of master mix into each well of a 96-well qPCR plate.
   3. Add 1 µL of cDNA to each well to obtain a final reaction volume of 10 µL.
   4. Seal the plate with optical adhesive film and centrifuge at 1,000 × g for 1 min.
   5. Perform initial denaturation at 95 °C for 30 s.
   6. Perform 40 amplification cycles of 95 °C for 5 s followed by 60 °C for 30 s.
   7. Perform melt-curve analysis to confirm amplicon specificity.
      NOTE: Run each sample in duplicate or triplicate.
   8. Calculate relative target-gene expression using the 2^−ΔΔCt method. Normalize Ct values to the housekeeping gene (e.g., GAPDH) and calibrate expression against undifferentiated hiPSC samples.

7. Statistical analysis

1. Present quantitative data as the mean ± standard deviation (SD) from at least three independent biological replicates (n ≥ 3) per condition.
2. Treat each independent batch as one biological replicate for aggregate and organoid diameter measurements, and average the 30 individual structures measured within each batch to obtain a single batch-level mean.
3. Average vascular area, pericyte coverage, and branching index measurements from multiple organoids or fields within each batch to obtain a single batch-level value, and use these batch-level values for statistical comparisons.
4. Compare the optimized protocol with the original Wimmer protocol using an unpaired two-tailed Student's t-test.
5. Define statistical significance as p < 0.05. Indicate significance levels as p < 0.05, p < 0.01, and p < 0.001, as applicable.
6. Perform all statistical analyses using statistical analysis software version 10.

Results

The optimized protocol described here enables the generation of mature vascular organoids from hiPSCs within approximately 15 days using a stepwise differentiation strategy (Figure 1A). Briefly, hiPSC aggregates were generated in low-attachment plates supplemented with 0.25% PVA on an orbital shaker (Day −1). Mesoderm induction was initiated on Day 0 using CHIR99021 (12 µM) and BMP-4 (30 ng/mL) for 3 days, followed by vascular lineage specification with VEGF-A (100 ng/mL) and forskolin (2 µM...

Discussion

The optimized protocol described herein introduces several key improvements to the vascular organoid differentiation system originally established by Wimmer et al.¹⁸, with the aim of enhancing the homogeneity, reproducibility, and structural integrity of vascular organoid generation. Compared with the original protocol, the optimized method provides significant improvements in the following aspects. First, optimizing aggregate formation conditions substantially improves the homogeneity of embryoid...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.22 μm Bottle-Top Filter	Nalgene	FPE204030	For sterile filtration of media and buffers
15 mL Conical Tube	SparkJade	GD0001	Sterile
2 mL Microcentrifuge Tube	Eppendorf	22363204	Sterile
2-Mercaptoethanol (1000×)	Gibco	21985023	Use in fume hood
Alexa Fluor 488 AffiniPure F2 Fragment Donkey Anti-Rabbit IgG (H+L)	Lablead	Y1104	Secondary antibody; dilute 1:300
Alexa Fluor 594 AffiniPure F(ab′)2 Fragment Donkey Anti-Goat IgG (H+L)	Lablead	Y1007	Secondary antibody; dilute 1:300
Anti-CD31 antibody	Cell Signaling	3528s	Primary antibody; dilute 1:100 in blocking buffer
Anti PDGFR β antibody	Zenbio	347269	Primary antibody; dilute 1:100 in blocking buffer
Automated Cell Counter	Life Technologies	Countess™ II	Includes counting chamber slides
B27 Supplement (50×)	Gibco	17504044	For N2B27 medium; thaw at 4ºC, aliquot and store at −20ºC, avoid freeze-thaw
Benchtop Centrifuge	Zonkia	SC-3610	With swinging-bucket rotor
Biological Safety Cabinet (Class II)	Thermo Fisher	NA	NA
BMP4	Proteintech	HZ-1045	Reconstitute in sterile ddH2O to 100 ng/μL, aliquot and store at −20 °C
Bovine Serum Albumin (BSA)	Biosharp	9048-46-8	For blocking buffer
CHIR99021	MCE	HY-10182	Prepare 10 mM stock in DMSO, store at −20ºC for up to 1 year
CO2 Incubator	Fisher Scientific	LS-CO150	37 ºC, 5% CO2, humidified
Confocal Imaging Dish	Biosharp	BS-20-GJM	For imaging after staining
Confocal Microscope	LEICA	NA	Equipped with 10×, 20×, 63× objectives and ZEN software
DMEM	Vivacell	C3113-0500	For Matrigel dilution and collagen I solution
DMEM/F-12, GlutaMAX Supplement	Gibco	10565018	For Matrigel dilution and N2B27 medium
EDTA (0.5 M, pH 8.0)	Invitrogen	AM9260G	Dilute with PBS to 0.02% for passaging
Fetal Bovine Serum (FBS)	Gibco	10270-106	For N2B27 complete differentiation medium (15% final)
FGF-2	Novoprotein	C046	Prepare 1 mg/mL stock in sterile PBS, aliquot and store at −20ºC
Forskolin	R&D Systems	1099	Prepare 10 mM stock in DMSO, store at −20ºC
HEPES (1M)	Gibco	15630080	For collagen I solution
Hoechst	Beyotime	C1071S-4	Prepare 1 mg/mL stock in deionized water, aliquot and store at −20ºC
Inverted Microscope	LEICA	DMIL LED	Routine cell observation
Matrigel (growth factor-reduced)	STEEMA	BM05	Store at −20ºC after aliquoting, avoid freeze-thaw; handle on ice
MEM Non-Essential Amino Acids Solution (NEAA, 100×)	Gibco	11140035	For N2B27 medium
N2 Supplement (100×)	Gibco	17502048	For N2B27 medium; store at −20ºC protected from light
Neurobasal Medium	Gibco	21103049	For N2B27 medium
Nuclease Free Water (DNase/Rnase Free)	Lablead	Doo55WJ	For RNA/cDNA preparation and qPCR
Orbital Shaker	Joan Lab	NA	For washing steps
Paraformaldehyde (4% solution)	Servicebio	G1101	Dilute with PBS to 4% for fixation; use in fume hood
Penicillin-Streptomycin (100×)	Gibco	15140122	For N2B27 medium
PGM1 Medium	Cellapy	A1517001	hiPSC expansion medium; store at 4ºC protected from light, stable for 2 weeks
PGM1 Supplement (25×)	Cellapy	A1517101	For use with PGM1 Basal Medium
Polyvinyl Alcohol (PVA)	Sigma-Aldrich	P8136	Dissolve in PGM1 at 0.25% (w/v) to promote aggregation
PureCol (Bovine Type I Collagen, 3 mg/mL)	Advanced BioMatrix	5005	For collagen I-Matrigel mixture; test batch for polymerization capacity
qPCR Thermal Cycler	Thermo Fisher	StepOnePlus™	With melt curve capability; for real-time PCR detection
Reverse Transcription Kit	Accurate Biology	AG11706	For cDNA synthesis from total RNA; contains RT master mix with reverse transcriptase, buffer, dNTPs, and random primers
RNA Extraction Kit	Yeasen	19221ES50	For total RNA isolation from vascular organoids; includes gDNA elimination columns and RNase-free DNase I
Sodium Bicarbonate (7.5%)	Gibco	25080094	For collagen I solution
Sodium Deoxycholate	Sigma-Aldrich	D6750	Prepare 1% (w/v) solution in deionized water for blocking buffer
Sodium Hydroxide (1.0 N)	Sigma-Aldrich	S2770	For pH adjustment of collagen I solution
Sterile Disposable Pipettes (10 mL)	Bkmamlab	110205007	Individually wrapped, sterile
Sterile Filter Pipette Tips (1000, 200, 20 μL)	Biozym	VT0270/VT0250/VT0220	Sterile, with filter
Sterile Needle (30 G)	NA	NA	For dissecting vascular networks
Sterile Spatula (Round End)	Fisher Scientific	21-401-5	Stainless steel
SYBR Green Master Mix	Accurate Biology	AG11733	2× concentration for qPCR; contains hot-start DNA polymerase, buffer, dNTPs, and SYBR Green dye
Tissue Culture-Treated 12-Well Plate	BD Falcon	353043	For gel embedding and vascular network culture
Tissue Culture-Treated 6-Well Plate	Eppendorf	30720113	Standard TC-treated
Triton X-100	Sigma-Aldrich	T9084	For blocking buffer
Trypan Blue (0.4%)	Thermo Fisher	15250061	For cell counting
Tween 20	Sigma-Aldrich	P7949	For 0.05% PBS-T wash solution
Ultra-Low Attachment 6-Well Plate	Corning	3471	For aggregate formation and differentiation
Ultra-Low Attachment Round-Bottom 96-Well Plate	Beyotime	FULA962-6pcs	For suspension culture of vascular organoids
VEGF-A	Proteintech	HZ-1038	Prepare 1 mg/mL stock in sterile PBS, aliquot and store at −20ºC
Water Bath	Blue pard	HWS-24	37ºC
Y-27632	MCE	HY-10071	Prepare 10 mM stock in sterile PBS or DMSO, store at −20ºC; working concentrations: 5 μM (thawing), 50 μM (aggregate formation)